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5,035 | https://bio-protocol.org/en/bpdetail?id=5035&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Approach for Electrophysiological Studies of Spinal Lamina X Neurons
VK Volodymyr Krotov
PB Pavel Belan
NV Nana Voitenko
Published: Vol 14, Iss 14, Jul 20, 2024
DOI: 10.21769/BioProtoc.5035 Views: 316
Reviewed by: Sarajo MohantaMayank Gautam Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Pain Oct 2022
Abstract
Despite playing diverse physiological roles, the area surrounding the central canal, lamina X, remains one of the least studied spinal cord regions. Technical challenges and limitations of the commonly used experimental approaches are the main difficulties that hamper lamina X research. In the current protocol, we describe a reliable method for functional investigation of lamina X neurons that requires neither time-consuming slicing nor sophisticated in vivo experiments. Our approach relies on ex vivo hemisected spinal cord preparation that preserves the rostrocaudal and mediolateral spinal architecture as well as the dorsal roots, and infrared LED oblique illumination for visually guided patch clamp in thick blocks of tissue. When coupled with electric stimulation of the spared dorsal roots, electrophysiological recordings provide information on primary afferent inputs to lamina X neurons from myelinated and non-myelinated fibers and allow estimating primary afferent–driven presynaptic inhibition. Overall, we describe a simple, time-efficient, inexpensive, and versatile approach for lamina X research.
Key features
• Quick and easy preparation procedure that grants access to lamina X neurons without spinal cord slicing
• Preserved rostrocaudal and mediolateral connectivity and preserved primary afferent supply
• Ability to perform electrophysiological recordings in combination with dorsal root stimulations allowing to study afferent inputs and presynaptic inhibition of lamina X neurons
Keywords: Spinal cord Lamina X Electrophysiology Dorsal root stimulation Postsynaptic currents Presynaptic inhibition
Graphical overview
Preparation of ex vivo hemisected spinal cord and electrophysiological recordings from lamina X neurons
Background
Based on its cytoarchitectonics, the spinal cord is subdivided into ten regions called laminae [1]. Spinal laminae are studied to a different extent: while superficial ones have been explored in much detail, our knowledge about the deeper laminae is unsatisfactory. In particular, lamina X surrounding the central canal remains one of the least studied spinal regions despite playing diverse physiological roles. Evidence indicates that lamina X is crucial for visceral nociception [2–6] and participates in somatosensory processing [4,7,8], locomotion [9], and autonomic regulation [10]. Yet, reports on lamina X are scarce and derive mostly from morphological and immunohistochemical data. Functional investigations of lamina X are even rarer given that they mainly rely on two approaches. The first one involves in vivo electrophysiological recordings using either an extracellular electrode [4,7] or a blind patch clamp technique [10,11]. Both methods require post hoc tissue analysis to confirm recordings from lamina X cells. The other approach utilizes visually guided patch clamp in spinal cord slices [13–23] in which intrinsic spinal cord connectivity and primary afferent inputs are disrupted. Both mentioned methods are technically challenging and have inherent flaws that limit the functional investigation of lamina X. Thus, an experimental approach circumventing these limitations would greatly boost the lamina X research.
Here, we provide a detailed protocol for electrophysiological studies of lamina X neurons. The described procedure is easy and reliable. It allows recording over prolonged periods (up to 6–8 h) and requires only very basic equipment for ex vivo electrophysiology. The main advantage of our approach is the use of a spinal cord hemisected at the sagittal midline, which fully preserves rostrocaudal and mediolateral spinal architecture and spares the dorsal roots. Unlike spinal cord slices, the hemisected preparation is produced without any specialized device and needs little to no incubation; the preparation is ready for use within half an hour after decapitation. For cell visualization, we use infrared LED oblique illumination, which is specifically designed to work with thick blocks of tissue [24,25]. A combination of hemisected spinal cord preparation together with oblique LED illumination gives a unique opportunity to perform visually guided patch clamp in practically intact tissue. This is particularly beneficial for studying primary afferent inputs to lamina X and their modulation. Patch clamp experiments coupled with electrical dorsal root stimulation via suction electrodes enable recording elicited postsynaptic currents in lamina X neurons. Different parameters of electrical stimuli allow the examination of direct synaptic inputs from thinly myelinated A fibers and non-myelinated C fibers as well as their primary afferent–driven presynaptic inhibition. The current protocol is suitable for using both young rats and mice of various genetic backgrounds. Additionally, the protocol may be applied for studying various cell populations (such as spinothalamic, spinohypothalamic, and sympathetic preganglionic neurons) preliminary labeled with retrograde dyes [26]. Finally, the preserved rostrocaudal architecture of the hemisected preparation allows complementing the protocol with the stimulation of descending (top-down) fibers of the spinal cord [27].
Materials and reagents
Biological materials
Wistar rats P7–14 (Charles River, strain code: 003)
Reagents
Sucrose (Sigma-Aldrich, catalog number: S0389)
Glucose (Sigma-Aldrich, catalog number: G8270)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
Sodium bicarbonate (NaHCO3) (Sigma-Aldrich, catalog number: S0751)
Sodium monophosphate (NaH2PO4) (Sigma-Aldrich, catalog number: S6040)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9333)
Magnesium chloride (MgCl2·6H2O) (Sigma-Aldrich, catalog number: M2670)
Calcium chloride (CaCl2·2H2O) (Sigma-Aldrich, catalog number: C7902)
Potassium gluconate (Sigma-Aldrich, catalog number: G4500)
Sodium ATP (Na2ATP) (Sigma-Aldrich, catalog number: A3377)
Sodium GTP (NaGTP) (Sigma-Aldrich, catalog number: G8877)
HEPES (Sigma-Aldrich, catalog number: H3375)
EGTA (Sigma-Aldrich, catalog number: E0396)
Sodium ascorbate (Sigma-Aldrich, catalog number: 11140)
Sodium pyruvate (Sigma-Aldrich, catalog number: P2256)
Isoflurane (Abbvie, catalog number: B506)
95% O2 and 5% CO2 gas mixture
Solutions
Sucrose dissection solution (see Recipes)
Krebs bicarbonate solution (see Recipes)
Potassium gluconate intracellular solution (see Recipes)
Recipes
Sucrose dissection solution (final volume 0.5 L, in ddH2O)
Reagent Final concentration Quantity
Sucrose 200 mM 34,230 mg
Glucose 11 mM 991 mg
NaHCO3 26 mM 1092 mg
NaH2PO4 1.2 mM 72 mg
KCl 2 mM 74.5 mg
MgCl2·6H2O 7 mM 712 mg
CaCl2·2H2O 0.5 mM 37 mg
Sodium ascorbate (optional) 5 mM 495 mg
Sodium pyruvate (optional) 3 mM 168 mg
pH 7.3–7.4 when bubbled with 95% O2 and 5% CO2; osmolarity 310–320 mOsm/kg. This solution may be used for up to two weeks after preparation if kept at 4–8 °C.
Krebs bicarbonate recording solution (final volume 0.5 L, in ddH2O)
Critical: To avoid precipitation, the solution must be bubbled with 95% O2 and 5% CO2 gas mixture before adding calcium chloride.
Reagent Final concentration Quantity or Volume
NaCl 125 mM 3,653 mg
Glucose 10 mM 901 mg
NaHCO3 26 mM 1092 mg
NaH2PO4 1.25 mM 75 mg
KCl 2.5 mM 93 mg
MgCl2·6H2O 1 mM 102 mg
CaCl2·2H2O 2 mM 147 mg
pH 7.3–7.4 when bubbled with 95% O2 and 5% CO2; osmolarity 300–310 mOsm/kg. This solution may be used for up to two weeks after preparation if kept at 4–8 °C.
Potassium gluconate intracellular solution (final volume 50 mL, in ddH2O)
Reagent Final concentration Quantity or Volume
Potassium gluconate 145 mM 1,698 mg
MgCl2·6H2O 2.5 mM 25.5 mg
Sodium ATP 2 mM 55 mg
Sodium GTP 0.5 mM 13 mg
HEPES 10 mM 119 mg
EGTA 0.5 mM 9.5 mg
pH 7.3 adjusted with KOH; osmolarity 280–290 mOsm/kg. Make 1 mL aliquots and freeze them at -20 °C.
Laboratory supplies
Dissection dish with Sylgard-lined bottom (Living Systems Instrumentation, catalog number: DD-90-S-BLK)
25 G hypodermic needles (BD, catalog number: 300400)
30 G hypodermic needles (BD, catalog number: 304000)
Super glue (cyanoacrylate, water-resistant gel)
Glass capillaries with filament O.D. 1.5 mm, I.D. 0.86 mm (e.g., Sutter Instruments, catalog number: BF-150-86-10; Harvard Apparatus, catalog number: GC150F-10)
Thin-walled glass capillaries without filament O.D. 1.5 mm, I.D. 1.17 mm (Warner Instruments, catalog number: G150T-3)
1 mL Luer-Lok syringes (BD, catalog number: 309628)
2.5 mL Luer-Lok syringes (BD, catalog number: 300185)
Silicon tubing (VWR, catalog number: 228-0701)
Three-way tap (BD Connecta, catalog number: 394601)
Patch pipette fillers (WPI, catalog number: MF28G67-5)
Sterile syringe filters 0.22 μm (Thermo Scientific, catalog number: 171-0020)
Equipment
Dissection instruments
Dissection pad (could be made from Styrofoam tightly wrapped in foil)
Anesthesia induction chamber (VetEquip, US) or any other available
Big scissors (F.S.T., catalog number 14000-20)
Small scissors (F.S.T., catalog number 14058-11)
Coarse forceps (F.S.T., catalog number 11651-10)
Big spring scissors (F.S.T., catalog number 15025-10)
Small spring scissors (F.S.T., catalog number 15000-12)
Two curved forceps (F.S.T., catalog number 11063-07)
Small metal plate (preferably gold, should fit in experimental chamber)
Visualization and illumination
Stereomicroscope (Olympus, model: SZX7)
Light source (AmScope, model: 6 W LED Dual Gooseneck Illuminator)
Upright microscope (Olympus, model: BX50WI)
60× water immersion objective (Olympus, model: LUMPlanFl 60×/0.90 W)
Low magnification (4–5×) objective (Carl Zeiss, model: Epiplan; Olympus, model Plan N)
Eyepiece graticule (any fitting the microscope)
Infrared-sensitive camera (Olympus, model: OLY-150IR or Dage-MTI, model: IR-2000)
Narrow beam (± 3°) infrared light emitting diode (Osram, models: SFH4550, SFH4545)
White light emitting diode (Dialight, model: 5219901802F)
Power supply unit (AimTTI, model: QL355T)
Perfusion
Gravity-fed perfusion system (any self-made or commercially available)
Peristaltic pump (Gilson, model: Minipuls 3)
Recording chamber (any fitting the microscope)
Electrophysiology
Patch clamp amplifier with respective headstage (Molecular Devices, models: Multiclamp 700B, Axopatch 200B)
Digitizer (Molecular Devices, models: Digidata 1440, Digidata 1550)
PC computer with Windows operating system
Patch clamp micromanipulator (e.g., Scientifica, model: PatchStar; Sutter Instruments, model: MP-225)
Constant current stimulator (e.g., A.M.P.I., model: ISO-FLEX; A-M Systems, model: 2100; Digitimer, model: DS3)
Vibration isolation table (CleanBench, model: TMC)
Faraday cage (Sutter Instruments, model: AT-36FC)
Small three-axis micromanipulators (Narishige, model: UN-3C)
Pipette holders (Molecular devices, model: 1-HL-U)
Pipette puller (Sutter Instruments, models: P-87, P-97)
Silver wire for electrodes (WPI, product number: AGW1510)
Ag/AgCl pellets (WPI, product number: EP1)
BNC cables
Bunsen burner
Software and datasets
pClamp (version 10.7, Molecular Devices, July 2016)
Clampfit (version 10.7, Molecular Devices, July 2016)
Origin (version 2022, OriginLab, June 2022)
MiniAnalysis (version 6.0.7, Synaptosoft Inc.)
Procedure
Critical: Before the start of the experiments, make sure both sucrose dissection solution and Krebs bicarbonate solution are at room temperature (20–23 °C) and have been bubbled with 95% O2 and 5% CO2 gas mixture for at least 15 min. Changes by 2 °C or more significantly impact the conduction velocity of primary afferent fibers, therefore compromising the classification of the monosynaptic components of primary afferent input to lamina X neurons. Cooled (4–8 °C) solutions compromise the quality of ex vivo spinal cord preparation.
Ex vivo spinal cord preparation
Prepare dissection instruments and place them next to the stereomicroscope.
Deeply anesthetize the animal with isoflurane. Ensure loss of pedal reflex by pinching the hind paw. Quickly decapitate the animal with big scissors and drain the blood for 10 s.
Excise the spinal column with attached ribs.
Place the decapitated rat on the dissection pad and pin its paws with 25G needles as shown in Figure 1.
Figure 1. Excision of the spinal cord with attached ribs. Schematic depiction of the excision process. Red dots: sites to grasp using forceps. Dash lines: lines of cut. Numerals denote the order of cuts for a right-handed person.
Remove the skin from the back to expose the spinal column and the ribs. Using coarse forceps, pull the skin up and insert one branch of small scissors underneath it. Push the scissors to either side and cut the skin along the flank holding the scissors parallel to the dissection pad. Then, cut the skin along the other flank. Finally, cut the skin near the tail (Figure 1).
Hold the scissors vertically with the branches open and facing down. Thrust the scissors into the rat’s back by 3–5 mm at the level of the hips so that the branches are around the spinal column. Cut the spinal column.
Using coarse forceps, grab the cut part spinal column, pull it up, and cut the tissues rostrally along each side of the spinal column until the ribs are reached.
Cut the ribs on each side and cut any thoracic and abdominal viscera that might be attached to the spinal column or the rib cage.
Note: The ribs are necessary for pinning the preparation to the dissection dish. Therefore, do not cut them too short.
Hold the spinal column vertically and cut it at the lower cervical/upper thoracic level.
Put the excised spinal column into a dissection dish with a Sylgard-lined bottom filled with sucrose solution continuously bubbled with 95% O2 and 5% CO2 gas mixture.
Using 30G needles, pin the excised tissue dorsal side down (ventral side up) rostrally to the experimenter as depicted in Figure 2.
Focus the stereomicroscope on the preparation (8–10× magnification).
Using coarse forceps, grab the ribs near the rostral part of the spinal column and slightly pull it up to see the opening. Insert one branch of big spring scissors inside the opening, press the blade against the vertebrae, and cut them. Keeping the branches closed, move the scissors to the side to evacuate them. Then, cut the other side of the vertebrae in a similar manner.
Grasp the cut ventral part of the spinal column with coarse forceps and slightly pull it upward and caudally. This allows us to clearly see the space between the spinal cord and the vertebrae where the branch of the scissors needs to be inserted. Continue cutting the vertebrae as described above, keeping the scissors parallel to the dissecting dish (Figure 2). Move the forceps as you cut and take extra care once lumbar enlargement is reached. Cut any dura mater (an opaque sheath covering the spinal cord) attached to the ventral part of the spinal column. Once cauda equina (a group of nerves and nerve roots stemming from the distal end of the spinal cord) is exposed, cut off the ventral part of the spinal column and remove it from the dish.
Using small spring scissors, cut the dura mater along the midline all the way to the caudal segment of the spinal cord. Gently pull the dura mater with curved forceps if necessary.
Remove the spinal cord from the spinal column (Figure 2).
Cut the cauda equina.
Using the side of the vertebrae as a rail, cut thoracic dorsal and ventral roots on each side of the spinal cord.
Using curved forceps, pull the spinal cord upward and to the side and cut dura mater attached to the dorsal part of the vertebrae. Hold the spinal cord by its very rostral part or, even better, by the dura mater or thoracic roots.
Critical: The spinal cord should always remain in aqueous solution. Do not stretch the preparation.
Cut the lumbar dorsal roots sparing their whole length.
Cut any other tissue that attaches the spinal cord to the inside of the spinal column.
Turn the spinal cord ventral side down and cut the dura mater on the dorsal side along the midline.
Figure 2. Schemes illustrating stages of making hemisected spinal cord preparation. Modified from Krotov et al. [26].
Optional: Cut the very rostral part of the spinal cord if it is severely damaged.
Turn the spinal cord ventral side up. Take big spring scissors, position the branches along the midline, and make a single cut (2–4 mm long).
Note: Hemisecting the spinal cord with scissors/scalpel/blade damages lamina X; use the scissors solely to perform the initial cut.
Take each side of the spinal cord with curved forceps and spread them apart as shown in Figure 2 to expose lamina X. This results in two hemisected spinal cord preparations with spared roots. In approximately 25% of cases, the dorsal roots on one of the hemisected cords are severed. That happens more often if the dura mater is not cut properly.
Note: All the above steps should be completed within 7–10 min after decapitation. Longer preparation time decreases the number of lamina X neurons that are fit for recordings.
Remove the spinal column from the dissection dish.
Cut off damaged parts of the spinal cord, thin sacral segment, remainders of the dura mater, and dorsal/ventral roots not destined for stimulations/recordings. Typically, dorsal L3–L5 roots are spared. Avoid crushing the spinal cord with the forceps. Use dura mater or unwanted roots to handle the spinal cord.
Note: The hemisected cord is slightly curved, and the outer curve corresponds to the dorsal side.
Position the preparation so that the spinal cord and the dorsal roots form an inverted T letter (Figure 3).
Figure 3. Scheme illustrating the stages of gluing hemisected spinal cord preparation to a metal plate
Glue the preparation to the metal plate.
Place the metal plate on the base of the stereomicroscope close to the dissection dish.
Use 25 G needle to spread cyanoacrylate glue. Create a thin layer covering half of the plate (Figure 3).
Using two curved forceps, grab the hemisected spinal cord by its most rostral and caudal parts. Remove the preparation from the aqueous solution and delicately transfer it to the part of the metal plate that is NOT covered with glue. Ensure the gray matter (i.e., the lamina X) faces upward. Do not let the forceps go.
Slightly stretch the preparation, slightly lift above the metal plate, and put the hemisected cord on the glue-covered part of the plate. Do not lift the preparation too high; dorsal roots should stay on the glue-free part of the plate and the drag should help to avoid getting the roots glued (Figure 3).
Use the curved forceps to grab the metal plate and transfer it to the dissection dish.
Note: Minimize exposure to air. Ideally, steps A18c–e should be completed within 10–20 s.
Use a 30 G needle to detach the roots from the plate. The roots should float freely (Figure 4).
Figure 4. Ex vivo hemisected spinal cord preparation. Left: Lumbar segment of hemisected spinal cord with spared dorsal and ventral roots. Right: Ex vivo preparation glued to the metal plate. Notice spared L4 and L5 dorsal roots. Typically, lumbar and lower thoracic (T6–13) segments are available for recordings. The length of the dorsal roots is expected to be 4–6 mm. Modified from Krotov et al. [26].
To avoid unnecessary vibration during recordings, cut caudal and rostral (unless descending stimulation is to be performed) parts of the hemisected cord that stick out of the metal plate or are not glued properly.
Optional: Cut off any damaged ends of the dorsal roots. Cut the root to 2–3 mm length for dorsal root potential recordings. Use sharp scissors to avoid crushing the roots when cutting.
Setting up the perfusion and transferring the preparation to the recording rig
Install the experimental chamber and position the patch clamp reference electrode.
Fill the perfusion system with Krebs bicarbonate solution; at least 30–40 mL is necessary if the solution is recycled.
Note: Incorporate drippers into the outflow to avoid peristaltic pump–associated electrical noise.
Set the flow rate to 1.5–3 mL/min. To achieve that, use glass capillaries for inflow and outflow and fire polish their tips to an appropriate diameter. Alternatively, a flow regulator may be used.
Ensure that the flow through the chamber is stable and continuous. No ripples should be observed in the experimental chamber.
Before transferring the preparation, make sure the solution is thoroughly bubbled with 95% O2 and 5% CO2 gas mixture for at least 5 min.
Bring the dissection dish containing the preparation close to the recording chamber. Use the forceps to grab the metal plate to which the preparation is glued and transfer it to the chamber.
Important: Once in the recording chamber, the entire preparation including the dorsal roots should always be covered with the solution.
Position the metal plate so that the rostrocaudal axis of the preparation is along either the x- or y-axis of the microscope table.
Allow the preparation to accommodate the Krebs bicarbonate recording solution for 10 min before any recordings.
Attaching dorsal roots to suction electrodes
Using white LED as a source of light and a low magnification objective (4–5×), focus on the tip of the dorsal root and assess its width using eyepiece graticule or camera software.
Note: It is possible to use a portable white LED flashlight for attaching dorsal roots and positioning the preparation. However, having a designated stationary white LED is more convenient.
Fabricate a suction electrode by fire-polishing a glass capillary. The opening of the suction electrode should be slightly narrower than the width of the root.
Note 1: It is recommended to manufacture a calibrated set of suction electrodes with various opening diameters beforehand and choose an appropriate one during the experiment, similarly to the recordings from the sciatic nerve [28].
Note 2: Use thin-walled glass capillaries to manufacture suction electrodes with narrow openings.
Insert the suction electrode into the pipette holder mounted on a manipulator (compact one with a magnet base is preferable) and connected to the anode of the constant current stimulator.
Position the manipulator and dip the suction electrode into the experimental chamber. Connect a 2.5- or 5-mL syringe to the pipette holder using silicone tubing; apply negative pressure to draw the solution into the suction electrode until it reaches the AgCl wire.
Position the opening of the suction electrode right next to the tip of the dorsal root. The suction electrode and the root should be aligned; if necessary, push the root with the forceps to achieve this configuration. Apply negative pressure sufficient for the root to go inside the electrode.
Visually check the tightness of the contact between the root and the electrode; it should be as shown in Figure 5. If a substantial proportion of the root enters the suction electrode creating excessive tension, the opening is too wide, and the glass electrode should be fire-polished further. If the root bulges inside the suction electrode, the opening is too narrow.
Note: Given that action potential initiation in response to electrical stimulation of the dorsal root occurs at the opening of the suction electrode [29,30], just the distal part of the root needs to go inside. Longer dorsal roots allow us to more easily distinguish between postsynaptic currents driven by fast-conducting (myelinated) A-fiber and slow-conducting (non-myelinated) C-fiber primary afferents.
Ensure the dorsal root does not pull the hemisected spinal cord. Otherwise, the preparation may move during recordings.
Position the AgCl reference electrode (connected to the cathode of the stimulator) next to the suction electrode.
Repeat steps C1–8 for other root(s).
Optional: For recording dorsal root potentials that spread passively, the proximal part of the root needs to be inside the recording suction electrode (Figure 5) to maximize signal-to-noise ratio.
Figure 5. Ex vivo spinal cord preparation with dorsal roots attached to suction electrodes. The walls of the suction electrodes are highlighted in red. Notice the tight contact between the roots and the electrodes. Top electrode: arrangement for dorsal root stimulation; the distal part of the root is inside the electrode. Bottom electrode: arrangement for dorsal root potential recordings; the proximal part of the root is inside the electrode. The configuration shown in the figure allows recording afferent-driven dorsal root potentials, a generalized readout of primary afferent–driven presynaptic inhibition.
Visualization of lamina X neurons
Visualization of lamina X neurons is performed using oblique infrared illumination (Figure 6) for which the following is necessary:
Narrow beam (± 3–5°) infrared light-emitting diode (IR-LED) connected to an adjusted power supply unit and mounted on a micromanipulator (or a gooseneck) allowing IR-LED to change its position in relation to the ex vivo spinal cord preparation.
Note: For convenience, it is better to mount IR and white LEDs on the same micromanipulator.
60× objective.
Note: It is possible to use a 40× objective; yet, it is less convenient than the 60× one and digital zooming might be required for patching small lamina X neurons.
CCD-camera coupled to a computer (or a separate display in case of an analog camera).
Note: Make sure that the spectral characteristics of IR-LED and CCD-camera match.
Turn these devices on for visualization.
Figure 6. Schematic illustration of the experimental setup for oblique LED illumination. Left: notice the experimental chamber with perfusion inflow and outflow tubes, low-magnification objective, high-magnification water-immersion objective, and white and IR-LED mounted together on a micromanipulator. The LEDs should not be immersed in the solution. Right: notice two LEDs: white LED used together with a low-magnification objective for rough positioning and attaching dorsal roots to suction electrodes, and IR-LED used with a high-magnification objective for cell visualization. The IR-LED should be positioned at an acute (10–20°) angle in relation to the spinal cord preparation. Also, notice ex vivo spinal cord preparation glued to a metal plate; spared dorsal root is attached to the suction electrode. The patch pipette for electrophysiological recordings from lamina X neurons is located in the middle of the dorsoventral axis of the preparation. Modified from Krotov et al. [26].
Install IR-LED so that its beam forms a 10–20° angle with the plane of the experimental chamber (Figure 6).
Note: The exact position of the LED greatly depends on its properties and needs to be found experimentally.
Using a white LED and low-magnification objective, focus on the desired spinal segment approximately at the middle of the dorsoventral axis of the preparation. Once done, turn off the white LED.
Switch to the 60× objective and focus on the surface of the preparation to see the cells. Typically, the cells of the ependyma (Figure 7A), a thin (5–10 μm) neuroepithelial lining of the central canal, are observed first. The ependymal layer is not uniform, and some lamina X neurons may be observed on the very surface of the preparation (Figure 7B). Lamina X neurons are mostly located underneath the ependyma (Figure 7C) and are approximately 20–40 μm long, although some may amount to 70–80 μm.
Critical: Make sure the image is still, i.e., the preparation does not move or wobble. Slow lateral movement indicates that the dorsal root attached to a suction electrode is pulling the preparation. A wobbly image indicates that a) the flow of the solution is unstable or b) the preparation is not glued to the metal plate properly. Apply additional glue and/or stabilize the flow before proceeding further.
Note: The auto-contrast option of the camera should be turned on.
Figure 7. Lamina X cells observed using IR-LED oblique illumination. A. Ependymal cells lining the central canal. B. Lamina X neurons on the surface of hemisected spinal cord preparation. C. Lamina X neurons located underneath the ependymal layer. Modified from Krotov et al. [26].
Adjust the position of the IR-LED for the best quality of the picture. Change the intensity of LED emission if necessary. When adjusted correctly, it is possible to visualize the cells up to 60 μm deep. In younger (P7–9) animals in which the ependyma is less pronounced, it is possible to visualize cells up to 80–100 μm deep.
Note: In oblique illumination, healthy-looking cells are usually perceived as convex, which is convenient for further patching. In case the cells appear concave, mirror the image using camera software or physically turn the camera 180 degrees.
Choose a lamina X neuron for patch clamping. The dorsal and ventral boundaries of lamina X are defined by bundles of parallel fibers spanning in the rostrocaudal direction (Figure 8). If the dorsal root is to be stimulated, choose neurons from the same (in respect to the dorsal root) or rostrally adjacent segments (i.e., if L5 root is stimulated, choose neurons from L4–5 spinal segments), since they receive most of the primary afferent input.
Figure 8. Rostrocaudal fibers of the spinal cord (parallel bundles) defining dorsal and ventral boundaries of the lamina X. Occasionally, cells may be observed in between the parallel fibers (right). Such cells do not belong to lamina X. Modified from Krotov et al. [26].
Patch clamping lamina X neurons
Turn on the amplifier, digitizer, constant current stimulator, and manipulator. Run their respective software. Set the amplifier to voltage-clamp mode.
Using a silicon tube, connect the pipette holder to a three-way tap with an attached 2.5 (or 5 mL) syringe and a mouthpiece (1 mL syringe without the plunger).
Thaw 1 mL vial of potassium gluconate intracellular solution. Fill a 1 mL syringe and put on a syringe filter and a patch pipette filler.
Note: Keep the syringe on ice or in the refrigerator (4–8 °C) during the experiment to avoid aggregation of potassium gluconate and/or NaGTP that may cause pipette blockage.
Use glass capillaries and a pipette puller to fabricate patch pipettes.
Fill a patch pipette with potassium gluconate solution. Insert the pipette into the holder connected to the headstage of the amplifier.
Lift the water immersion objective to create a meniscus that would accommodate the conical part of the patch pipette. Introduce the pipette and focus on its tip.
Visually ensure the opening of the pipette is not blocked. Assess the resistance of the pipette using Membrane test (Bath) in pClamp software. The pipette needs to have a resistance of 3–5 MΩ when filled with potassium gluconate solution.
Continuously apply positive pressure using a 2.5 mL syringe (displace the plunger by 1–1.5 mL). Ensure there is an outflow of solution from the pipette. Compensate offset potential.
Note: When using potassium gluconate solution, applying positive pressure increases the resistance between the patch pipette and the reference electrode.
Approach the surface of the preparation with the pipette: iteratively lower the objective first and catch up with the pipette. Once done, switch the manipulator to a lower speed and position the tip of the pipette right above the center of the soma of a chosen neuron.
Descend the pipette toward the neuron. The flow from the pipette should visibly spread the tissue. In case the ependymal layer prevents that, gently tap on the headstage and reposition the tip of the pipette above the center of the neuron.
Keep descending to the surface of the desired neuron. At some point, the flow from the pipette is going to indent the membrane. Once the indentation is approximately twice the size of the pipette opening, use the mouthpiece to apply negative pressure (as if you were drinking through a straw). Switch the tap to the mouthpiece while applying the negative pressure. At this point, the resistance between the patch pipette and the reference electrode should abruptly increase. When it reaches 200–300 MΩ, release the negative pressure. From this point, the gigaseal (the resistance over 1 GΩ), which defines the cell-attached patch clamp configuration, should establish on its own. If that does not happen, facilitate the gigaseal formation by switching to negative potentials and applying negative pressure with the mouthpiece.
Important: Insufficient indentation compromises the gigaseal formation. Excessive indentation compromises establishing whole-cell patch clamp configuration. In case the gigaseal cannot be achieved on a regular basis, change the puller settings to increase the resistance of the patch pipettes by additional 1–2 MΩ.
Critical: The flow of the potassium gluconate solution from the patch pipette depolarizes surrounding neurons. Therefore, keep the time between penetrating the tissue and establishing the gigaseal to a minimum.
Note: The flow of the solution from the pipette may move the cell. Use the manipulator to always keep the tip of the patch pipette above the center of the cell.
Compensate pipette capacitance and let the contact between the cell and the pipette improve for a couple of minutes. Ideally, the absolute value of the observed current should be below 20 pA or, even better, 10 pA.
Optional: Perform action-potential recordings in the cell-attached configuration. Recordings in the current clamp mode (I = 0) allow further analysis of action potential parameters and are more sensitive to sub-threshold events. Recordings in the voltage clamp mode (V = 0) are less sensitive, hence providing a smoother baseline, and are mostly used for assessing the number of action potentials (see [31] for a detailed description of cell-attached recordings). Typically, recordings in the cell-attached mode are coupled with dorsal root stimulation (see section F).
Note: Ideally, the recording pipette should be filled with filtered extracellular solution to assess genuine unperturbed activity of a neuron. Yet, that approach makes any further whole-cell experiments impossible. Using a high-potassium internal solution may be a reasonable compromise, allowing both to estimate neuronal activity and to conduct further intracellular recordings. Given that in our experiments dorsal root stimulation evoked the same number of spikes in cell-attached and whole-cell configurations, we assume that depolarization of a small membrane patch does not affect the spike generation significantly.
Set the holding potential to -60 (or -70) mV and establish whole-cell patch clamp configuration. For that, apply negative pressure through a mouthpiece and gradually increase it until the flat line on the membrane test (Patch) indicating gigaseal contact changes to a classic whole-cell transient. Once the transient appears, release the negative pressure immediately. Alternatively, apply a small amount of negative pressure and use the zap function of the amplifier. Typically, 50–250 μs zap duration is enough for a breakthrough.
Critical: Monitor series resistance often, as it tends to increase over time (generally, the higher the pipette resistance, the greater the increase in series resistance). Make sure that series resistance does not change by more than 20% to avoid compromising collected data.
Note: Series resistance may vary substantially from cell to cell; larger lamina X neurons exhibit 15–25 MΩ series resistance, and smaller ones exhibit -25–35 MΩ. Typical values of series resistance are an order of magnitude lower than those of membrane resistance.
Optional: Compensate whole-cell transient and compensate series resistance. That does not always work and, often, setting compensation and/or prediction over 70% results in oscillations that may ruin the whole cell configuration.
Electrophysiological recordings and dorsal root stimulation
For recordings, set the digitization at 10–20 kHz and the Bessel filter at 2.4–3 kHz.
In the first minute after establishing whole-cell configuration, run the protocols for determining membrane resistance and capacitance as well as the firing properties of the cell. This information might help in the classification of studied neurons.
For determining resistance and capacitance of the membrane, apply -10 mV hyperpolarizing step for 200 ms at 1 Hz (10–15 trials).
For determining the firing pattern of a neuron, switch to the current clamp mode and inject a negative current sufficient to hyperpolarize the cell to -80 mV (necessary to recover potassium voltage–gated channels from inactivation). Apply incrementally increasing 500 ms steps of current at 0.3 Hz; start with -20 pA and increase the current by 10 or 20 pA with each trial. Do not exceed 400–500 pA current.
Before proceeding to the next recordings, check the series resistance and adjust it if necessary.
For assessing network activity, allow cell dialysis for at least 5 min after establishing whole-cell configuration. Then, record spontaneous excitatory postsynaptic currents (sEPSCs) at -60 or -70 mV in a gap-free mode. At these potentials, sEPSCs appear as negative deflections of current. Positive deflections correspond to spontaneous inhibitory postsynaptic currents; however, they are unlikely to appear, given that the reversal potential for chloride ions is around -80 mV for the combination of external Krebs bicarbonate and internal potassium gluconate solutions. Add TTX (final concentration 1 μm) and D-AP5 (final concentration 20–40 μm) to Krebs bicarbonate solution for recordings of AMPA receptor-mediated miniature postsynaptic currents.
For assessing primary afferent supply of lamina X neurons, combine recordings at -60 or -70 mV with dorsal root stimulation. Connect the stimulator to the digital output of the digitizer. Set the stimulator to square pulses of current. Set the stimulation frequency to 0.1 Hz to avoid slowing down of conduction [32] and the wind-up phenomenon [33]. Apply the following values of stimulus intensity and duration to activate the primary afferents in the dorsal roots:
Use +30–50 μA × 50 μs stimuli to activate thick fast-conducting myelinated low-threshold Aβ-fiber primary afferents.
Use +60–100 μA × 50 μs stimuli to activate Aβ-, thinly myelinated Aδ-, and slow-conducting low-threshold C-fiber primary afferents.
Use +20–150 μA × 1 ms stimuli to activate all primary afferents including high-threshold Aβ, Aδ, and C fibers.
Note: In physiological conditions, primary afferent–driven postsynaptic currents are saturated at 70–100 μA; thus, the 100–150 μA range corresponds to supramaximal stimulation.
Use negative 20–150 μA × 1 ms stimuli to exclusively activate C-fiber primary afferents. Stimuli of negative polarity are necessary to induce an anodal block of fast-conducting A fibers [29,30,35], thus ensuring exclusive C-fiber activation.
Note: The exact stimulus duration necessary to induce anodal block of A fibers depends on the configuration of the suction electrode. In case A fiber–driven postsynaptic currents are observed when stimulating with negative current, adjust the duration in 0.8–1.2 ms range.
Record at least 12–15 trials for any selected stimulation.
For assessing homosegmental A fiber–driven presynaptic inhibition (i.e., decrease of C fiber–driven postsynaptic currents mediated by A fibers from the same dorsal root), perform the recordings as described in the step above. Stimulate the dorsal root at +150 μA × 1 ms first and then at -150 μA × 1 ms.
For assessing heterosegmental C fiber–driven presynaptic inhibition (i.e., decrease of A and C fiber–driven postsynaptic currents mediated by C fibers of the adjacent dorsal root), make paired stimulations. First, perform conditioning C-fiber stimulation (-150 μA × 1 ms) and record elicited postsynaptic currents (this is necessary for subtracting postsynaptic current elicited by conditioning stimulation from the one elicited by the test stimulation). Then, stimulate the adjacent dorsal root (choose one of the patterns from step F5) and record test postsynaptic currents in the absence of conditioning stimulation. Finally, make recordings with paired stimulations; apply conditioning stimulus 80–100 ms before the test one (Figure 9).
Protocols described in steps F3–7 may be repeated in the presence of a desired pharmacological agent.
Record at -10 mV using the protocols described in steps F3–7 to assess spontaneous and primary afferent–driven postsynaptic inhibitory currents.
Critical: Monitor series resistance often. Do not take data for quantitative analysis if series resistance changes by more than 20% during recording.
Figure 9. Electrophysiological recordings for assessing heterosegmental C fiber–driven presynaptic inhibition of the afferent input to lamina X neuron. Unprocessed recording (unfiltered, unadjusted baseline) showing individual traces. The red line is an average of traces recorded in the absence of conditioning stimulation. Notice the decreased amplitude of the monosynaptic component of primary afferent input in the presence of conditioning stimulation. Inset: experimental design of paired dorsal root stimulation coupled with patch clamp recordings. Test stimulation of L4 dorsal root (+150 μA × 1 ms) was performed to induce primary afferent–driven postsynaptic currents. Conditioning stimulation (-150 μA × 1 ms) of L5 dorsal root was performed to observe C fiber–mediated presynaptic inhibition of the primary afferent input from L4 dorsal root. Conditioning stimulus was applied 100 ms prior to the test one. Recordings were made from a lamina X neuron of L4 spinal segment. Modified from Krotov et al. [30].
Optional: To assess generalized presynaptic inhibition in the entire spinal segment, use the dorsal root to record passively spreading dorsal root potentials. For that, ensure the recording suction electrode is attached to the dorsal root as described in step C10. Set the amplifier to current clamp mode. Given that dorsal root potentials rarely exceed 0.2–0.3 mV, use high-gain settings. To elicit primary afferent–driven dorsal root potentials, use stimulation patterns from step F4. Although dorsal root potentials are highly reproducible, record at least 5–10 trials for averaging. Typically, dorsal root potentials peak 50–100 ms after stimulation (Figure 10).
Figure 10. Dorsal root potential recorded from L5 dorsal root. The potential was elicited by supramaximal (+150 μA × 1 ms) L4 dorsal root stimulation. Average of five traces, baseline adjusted to zero.
Data analysis
Data obtained from electrophysiological experiments on lamina X neurons may include recordings of spontaneous/miniature excitatory and inhibitory postsynaptic currents, primary afferent–driven postsynaptic currents elicited by dorsal root stimulation, and action potentials evoked by current injection via patch pipette or dorsal root stimulation. Hemisected spinal cord with spared dorsal roots also allows to perform recordings of compound action potentials driven by primary afferent activation as well as passively spreading dorsal root potentials caused by primary afferent depolarization elicited by the activation of spinal neurons. Spontaneous/miniature currents are better analyzed with MiniAnalysis software, which automatically returns amplitudes and kinetics of individual events. Other types of data such as the integrals and the amplitudes of evoked postsynaptic currents may be analyzed using either Clampfit or Origin software. Identification and classification of monosynaptic components of primary afferent inputs elicited by dorsal root stimulation are performed based on the following criteria [36,37]:
Low failure rates (less than 30%).
Small latency variations (less than 2 ms).
Conduction velocity (CV), calculated as the length of the dorsal root from the opening of the suction electrode to the dorsal root entry zone, divided by the latency of the monosynaptic response with a 1 ms allowance for synaptic transmission:
CV < 0.5 m/s corresponds to C-fiber afferents.
0.6 m/s < CV < 1.4 m/s corresponds to Aδ-fiber afferents.
CV > 3.5 m/s corresponds to Aβ-fiber afferents.
Note: Typically, Aβ afferents do not supply lamina X neurons [38].
Analysis of the amplitudes of monosynaptic components of primary afferent input is necessary for studying presynaptic inhibition.
Homosegmental A fiber–driven presynaptic inhibition: compare monosynaptic inputs to a particular lamina X neuron elicited by 150 μA × 1 ms stimuli of positive and negative polarities. After switching to negative polarity, faster A fiber–driven components disappear while C fiber–driven ones are time-shifted to the right by several milliseconds. Emergence (unmasking) of a new (previously unobserved) monosynaptic C fiber–driven input corresponds to full presynaptic block. An increase in the amplitude of existing monosynaptic C fiber–driven components corresponds to partial presynaptic block.
Heterosegmental presynaptic inhibition: compare monosynaptic components of postsynaptic currents before and after conditioning stimulation of the adjacent dorsal root. Reduced monosynaptic amplitude corresponds to partial presynaptic inhibition, and disappeared monosynaptic component indicates full presynaptic block.
Typically, obtained datasets do not follow normal distribution. Therefore, it is recommended to use non-parametric tests (for instance Mann-Whitney test) for statistical comparisons.
For presentation purposes, the signals may be slightly smoothed (Savitzky-Golay filter) to eliminate fast noise from the peristaltic pump. Dorsal root potential recordings may be filtered for sine wave noise.
Validation of protocol
This protocol has been used and validated in the following research articles:
Krotov et al. (2019). High-threshold primary afferent supply of spinal lamina X neurons. PAIN (Figures 1–4).
Krotov et al. (2022). Segmental and descending control of primary afferent input to the spinal lamina X. PAIN (Figures 1–5).
Krotov et al. (2023). Elucidating afferent-driven presynaptic inhibition of primary afferent input to spinal laminae I and X. Frontiers in Cellular Neuroscience (Figures 1–6).
The part of the current protocol regarding dorsal root stimulations has also been validated in the following articles:
Agashkov et al. [39]; Krotov et al. [34]; Tadokoro et al. [40])
General notes and troubleshooting
General notes
Electrophysiological recordings from lamina X neurons may be coupled with electrical [27] or optogenetic activation of descending (top-down) spinal cord tracts or calcium transient measurements using fluorescent indicators [26].
It is possible to work with defined populations of lamina X neurons using the following techniques: Injections of fluorescent tracer Fluorogold (or its analogues) into the thalamus or hypothalamus retrogradely labels spinothalamic or spinohypothalamic projection neurons in the lamina X [26]. Intraperitoneal injections of Fluorogold retrogradely label sympathetic preganglionic neurons located in thoracic and upper lumbar spinal segments [26]. The use of respective transgenic animals [41] allows to work with cholinergic lamina X neurons that are known to be involved in locomotion [9].
The current protocol may be applicable for electrophysiological studies of lamina X neurons in P7–P15 mice. Yet, the use of mice is more challenging given the smaller size of their spinal cord and dorsal roots in comparison with rats.
P10–12 Sprague-Dowley rats (Charles River, strain code: 001) may be used for the experiments. However, these animals possess a more pronounced ependymal layer and exhibit weaker primary afferent input to lamina X neurons than their age-matched Wistar counterparts.
Visualization of lamina X neurons in animals older than P15 becomes challenging due to the thicker ependymal layer and higher number of commissural fibers.
The current protocol is also suited for the electrophysiological investigation of Clarke column neurons (spinal lamina VII). Their cell bodies may be visualized in the thoracic segments of the spinal cord right next to the dorsal border of lamina X [42].
The current protocol may also be utilized for electrophysiological recordings of dorsal root potentials from hemisected spinal cords of adult mice. A similar approach was used by Zimmerman and colleagues [43].
Acknowledgments
This protocol is adapted from the previously published papers [26,27,30, 38].
The authors would like to thank Prof. Safronov (University of Porto), Dr. Oleh Halaidych and Mr. Andrew Dromaretsky (Bogomoletz Institute of Physiology). The drawing of the rat pup used in the graphical abstract was modified from the original image by Antonis Asiminas (doi.org/10.5281/zenodo.3926273) uploaded to SciDraw. Funding sources: this work was supported by NIH-1R01NS113189-01 grant (PB and NV), NASU 0124U001557, and 0124U001556 grants (PB).
Competing interests
The authors declare no competing interests.
Ethical considerations
All procedures involving animals were carried out with the approval of the Animal Ethics Committee of the Bogomoletz Institute of Physiology (Kyiv, Ukraine) in accordance with the European Commission Directive (86/609/EEC), ethical guidelines of the International Association for the Study of Pain, and the Society for Neuroscience Policies on the Use of Animals and Humans in Neuroscience Research.
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Received: Apr 15, 2024
Accepted: Jun 16, 2024
Available online: Jul 17, 2024
Published: Jul 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Correction Notice: Establishment of an in vitro Differentiation and Dedifferentiation System of Rat Schwann Cells
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DOI: 10.21769/BioProtoc.5036 Views: 163
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The protocol “Establishment of an in vitro Differentiation and Dedifferentiation System of Rat Schwann Cells” (DOI: 10.21769/BioProtoc.4631) has been corrected by the author to include an additional citation relevant to the protocol that had been omitted. Furthermore, the wording of several protocol steps was rephrased. All changes to the text are highlighted in the attached PDF file.
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5,037 | https://bio-protocol.org/en/bpdetail?id=5037&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Well Plate–Based Localized Electroporation Workflow for Rapid Optimization of Intracellular Delivery
CP Cesar A. Patino
SS Sevketcan Sarikaya
PM Prithvijit Mukherjee
Nibir Pathak
HE Horacio D. Espinosa
Published: Vol 14, Iss 14, Jul 20, 2024
DOI: 10.21769/BioProtoc.5037 Views: 592
Reviewed by: David PaulRosario Gomez-GarciaNeha Agarwal
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Original Research Article:
The authors used this protocol in Science Advances Jul 2022
Abstract
Efficient and nontoxic delivery of foreign cargo into cells is a critical step in many biological studies and cell engineering workflows with applications in areas such as biomanufacturing and cell-based therapeutics. However, effective molecular delivery into cells involves optimizing several experimental parameters. In the case of electroporation-based intracellular delivery, there is a need to optimize parameters like pulse voltage, duration, buffer type, and cargo concentration for each unique application. Here, we present the protocol for fabricating and utilizing a high-throughput multi-well localized electroporation device (LEPD) assisted by deep learning–based image analysis to enable rapid optimization of experimental parameters for efficient and nontoxic molecular delivery into cells. The LEPD and the optimization workflow presented herein are relevant to both adherent and suspended cell types and different molecular cargo (DNA, RNA, and proteins). The workflow enables multiplexed combinatorial experiments and can be adapted to cell engineering applications requiring in vitro delivery.
Key features
• A high-throughput multi-well localized electroporation device (LEPD) that can be optimized for both adherent and suspended cell types.
• Allows for multiplexed experiments combined with tailored pulse voltage, duration, buffer type, and cargo concentration.
• Compatible with various molecular cargoes, including DNA, RNA, and proteins, enhancing its versatility for cell engineering applications.
• Integration with deep learning–based image analysis enables rapid optimization of experimental parameters.
Keywords: Electroporation Intracellular delivery Deep learning Multiplexing Cell engineering Transfection
Background
Intracellular delivery of molecular cargoes allows for a wide variety of cell manipulation tasks with applications ranging from cell engineering and gene editing to cell therapy [1,2]. Given the wide implications of intracellular delivery for fundamental biology research and therapeutics, several technologies have emerged to achieve this task with high efficiency and precision that can be classified into two broad categories: i) direct delivery and ii) carrier-mediated delivery [3]. In direct-delivery methods, the cell membrane is permeated by subjecting it to physical disturbances such as mechanical forces [4–7], electric fields [8–10], or thermal shock [11]. Conversely, carrier-mediated delivery methods rely on chemical interactions between the cell membrane and the nanocarrier, which trigger endocytosis and encapsulation of the molecular cargo [3]. Examples of nanocarriers include lipid vesicles [12], spherical nucleic acids [13], and polymer nanoparticles [14]. Direct delivery methods offer more control and tunability over the membrane permeation process, which can result in tighter dosage precision compared with carrier-mediated methods [1,15].
Electroporation is one of the most widely used direct-delivery methods because of its ease of use and high throughput. In bulk electroporation systems, cells are placed in a cuvette with embedded electrodes that subject a suspension of cells to an electric field that induces the formation of pores across the cells when the electric potential across the membrane [transmembrane potential (TMP)] reaches a certain threshold. However, high applied voltages are required to achieve threshold TMP in bulk electroporation systems, which results in high cell mortality. Furthermore, the variability in the spatial distribution of the cells in the cuvette results in heterogeneous electric field conditions from cell to cell, which reduces dosage precision [1]. To circumvent these limitations, researchers developed electroporation systems that confine the electric field to a small fraction area of the cell membrane by interfacing cells with nanoscale orifices prior to the application of electric pulses [9,16,17]. The resulting highly localized electric fields enable the application of much lower voltages compared to bulk systems to achieve threshold TMPs, which results in higher viability. Examples of nanoscale features used in localized electroporation systems include nanochannel membranes [18,19], nanopipettes [20], nanostraws [21], and nanofountain probes [22].
Localized electroporation systems have been shown to be efficient in delivering various payloads (e.g., nucleic acids, plasmids, proteins, and nanoparticles) as well as sampling intracellular contents from cells [23,24]. The performance of localized electroporation systems depends on the electric field conditions, the mechanical properties of the cell membrane, the ionic environment in the surrounding fluid, and the physical properties of the molecular cargo (i.e., size and charge) [2,3]. While these systems offer significant advantages, they also pose challenges in scalability in the number of cells and require high cargo concentrations, which may limit their applicability in certain scenarios. Consequently, several parameters must be optimized to achieve the desired electroporation performance, including both delivery efficiency and cell health. To expedite the optimization process, we designed a well plate–based localized electroporation device (LEPD) equipped with multiplexing capabilities [9]. The multi-well LEPD system allows for tunability in the electric pulse conditions across the rows of the device and the molecular cargo or buffer conditions across columns. Furthermore, we integrated the LEPD with automated imaging and deep-learning enhanced segmentation to analyze the morphology of cells following delivery in addition to delivery efficiency and viability. The integrated approach provides a flexible platform suitable for adherent or suspended cell manipulation and analysis. In this protocol, we detail the experimental workflow of the LEPD spanning from device fabrication to optimization of plasmid delivery.
Materials and reagents
Well-plate electrode assembly
Custom printed circuit board (PCB) with gold electrode pads (Infinitesimal LLC, [email protected])
Custom PCB with through-holes (Infinitesimal LLC, [email protected])
Push-fit PCB receptacle (Mill-Max, catalog number: 0350-0-15-15-07-27-10-0)
Au-coated pins (straight pin or nail-head pin) (Mill-Max, catalog number: 3580-1-00-15-00-00-03-0)
Bottomless well plate (Greiner Bio-One, catalog number: 662000-06)
Biopsy punch, 12 mm (Acuderm, catalog number: P1250)
Double-sided pressure adhesive tape (Adhesives Research, catalog number: MH-90880)
Cutting board
Razor blade
Marker
Device assembly
Track-etched polycarbonate (PC) membranes (Itpi4, catalog number: 1000M25/620N401/7525)
Cloning cylinders (Millipore Sigma, catalog number: CLS31668)
Double-sided pressure adhesive tape (Adhesives Research, catalog number: MH-90880)
Biopsy punch, 6 mm (Miltex, catalog number: 33-36)
Acetone (Sigma-Aldrich, catalog number: 179124)
70% EtOH/H2O
Nitrogen (N2) (Airgas, catalog number: UN1066)
Oxygen (O2) (Airgas, catalog number: UN1072)
Scissors
Electroporation: low conductivity electroporation (EP) buffer (Eppendorf, catalog number: EW-36205-62)
Surface treatment and cell culture
Fibronectin from human plasma (0.1% solution) (Sigma-Aldrich, catalog number: F0895)
Dulbecco’s modified Eagle medium (DMEM), high glucose, pyruvate (Gibco, catalog number: 11995065)
Fetal bovine serum (FBS) (Gibco, catalog number: A3160501)
Penicillin-Streptomycin (Pen-Strep) (10,000 U/mL) (Gibco, catalog number: 15140148)
Trypsin-EDTA (0.25%), phenol red (Gibco, catalog number: 25200056)
NuncTM cell culture–treated multi-dishes, 6-well plates (Thermo Fisher Scientific, catalog number: 140675)
Phosphate buffered saline (PBS) (Gibco, catalog number: 10010023)
HeLa cells (ATCC, catalog number: CCL-2)
Plasmid: eGFP plasmids (Addgene, stock concentration 1,000 ng/μL)
Automated imaging: Hoechst 33342 nuclear stain (Invitrogen, catalog number: H3570)
Equipment
Electronics box for pulse generation and resistance measurements along with pulse control software (Infinitesimal LLC, [email protected])
Inverted microscope (Nikon Ti-E) equipped with 4×, 10×, and 20× objectives, fluorescent light source, an epi-filter cube with a DAPI filter, XYZ motorized stage, and a CMOS camera (Andor Zyla) for image acquisition
Computer for setting pulse parameters and image processing
Laboratory centrifuge (Thermo Fisher Scientific, catalog number: 75007200)
Sonicator (Ultrasonic Cleaning Bath, Branson, catalog number: 3800)
Oxygen plasma cleaner (Harrick Plasma, catalog number: PDC-32G)
Procedure
Well-plate electrode assembly
Place the PCB on the double-sided adhesive tape laid on a flat surface and use a razor blade to carefully cut the tape around the PCB’s edges, ensuring it matches the PCB’s dimensions.
Place the top PCB on top of the tape and use it as a stencil to mark the location of the wells using a marker (Figure 1B).
Use a plotter cutter or a biopsy punch (10–15 mm) to cut holes at the markings from the previous step (Figure 1C).
Remove one side of the adhesive tape by carefully inserting a razor blade at a corner and pulling the film with clean tweezers. Place the bottom PCB adjacent to the tape facing up (Figure 1D).
Use two tweezers to hold the tape from both sides, carefully align the tape to the bottom PCB by ensuring the holes on the tape are concentric to the Au pads on the PCB, and place the tape on the surface of the PCB.
Press down firmly on the adhesive tape to ensure a tight seal by using a rigid cylinder to roll over the adhesive film while applying force onto the surface (Figure 1E).
Remove the remaining protective film from the double-sided adhesive tape by separating the film from the adhesive tape using a razor blade and removing the laminate with a tweezer (Figure 1F).
Place the bottomless well plate on the surface while ensuring proper alignment of the wells to the Au pads. Firmly press down on the well plate to ensure a tight seal (Figure 1G).
Insert the receptacles onto the holes of the top PCB firmly and insert the Au-coated pins into the receptacles (Figure 1H and I).
Carefully position the pin headers at designated connection points on the top PCB. Solder PCB pin headers to the top PCB to connect the assembled well-plate electrodes to the function generator.
The aforementioned steps of well plate assembly are demonstrated in Video 1.
Figure 1. Assembly of well plate electrodes. (A) Materials required for the assembly procedure include an Au pin and receptacle, top and bottom printed circuit board (PCBs), double-sided adhesive tape, and a bottomless well plate. (B) Top PCB used as a stencil to mark the locations to cut holes (C) using a biopsy punch. (D) Tape after removal of the protective film is aligned with the bottom PCB. (E) A roller is used to compress the tape and the bottom PCB to form a tight seal. (F) The remaining protective film is removed from the adhesive tape. (G) The bottomless well plate is aligned to the bottom PCB pads and pressed firmly to ensure a tight seal. (H) Receptacles and pins are inserted into the through-holes of the top PCB. (I) Image of the fully assembled well-plate electrodes.
Video 1. Well-plate electrodes assembly
Device assembly
Immerse the glass cloning cylinders in acetone and sonicate for 10 min. Rinse the cylinders with ethanol and deionized distilled water (DDW) and dry with N2 gas.
Use a 6 mm biopsy punch to cut holes (~10 holes for a 75 mm × 25 mm area) through the double-sided adhesive tape (Figure 2B).
Remove one side of the protective film from the adhesive tape using a razor blade to separate the laminate from the tape at a corner and carefully remove the laminate with tweezers (Figure 2C).
Place the track-etched PC membrane on a clean glass slide and place it in oxygen plasma cleaner for 5 min.
Gently place the PC membrane on top of the tape from step B3 ensuring it remains flat by using the rigid microscope slide as support (Figure 2D and E).
Flip the PC membrane–adhesive tape and remove the remaining protective film from the adhesive tape using a razor blade and a tweezer (Figure 2F).
Place the cloning cylinder on the exposed adhesive tape aligned at the location of the pre-cut holes (Figure 2G and H).
Press down on the top of the glass cylinder to activate the pressure adhesive tape and ensure a good seal between the assembled layers (Figure 2I). Repeat this step for each hole cut in the adhesive tape.
Use scissors to cut the tape to separate each cylinder (Figure 2J) and subsequently trim the edge of the tape that surrounds each cylinder (Figure 2K and L). Spray the scissors with 70% ethanol in DI water to prevent the tape from sticking to the surface of the scissors.
Place the devices in a well-plate dish exposed to UV light for 4 h.
LEPD device fabrication and assembly steps are presented in Video 2.
Figure 2. Fabrication of the localized electroporation device (LEPD). (A) Materials needed for the process. (B) Double-sided tape with 6 mm holes punched through. (C) Peeling one side of the adhesive tape. (D) Placing porous membrane on a glass slide. (E) Pressing porous membrane onto the sticky side of the tape. (F) Peeling off the other side of the tape to expose the other side. (G–I) Placing glass cylinders onto the exposed sticky side of the tape and pressing them down to firmly stick. (J) Cutting out each LEPD. (K) Trimming out the edges of LEPDs. (L) Final result of an individual LEPD after completing the process.
Video 2. Fabrication and assembly of localized electroporation device (LEPD)
Surface treatment and cell culture
Prepare fibronectin solution for surface treatment by diluting the stock solution in PBS (20 µL of fibronectin in 1 mL of PBS) to obtain a final concentration of 20 µg/mL.
Add 100 µL of the prepared solution to each LEPD and incubate for 1 h at room temperature inside a biosafety cabinet.
Carefully discard the excess, unbound fibronectin solution from each LEPD and wash them with 100 µL of PBS two times. This ensures that only the adhered fibronectin layer is retained on the devices, which are now ready for cell culture.
Prepare complete cell culture media by adding 50 mL (10%) of FBS and 1 mL (1%) of Pen-Strep to 449 mL of DMEM.
Use this media for HeLa cell culture in 6-well plates.
Seed cells at a density of 3 × 105 cells/well and use 2 mL of media per well for culturing.
Wait until cells reach confluency (1 × 106 cells/well) in the well plates before dissociating them for plating in the LEPDs.
When cells are well adhered and confluent, gently discard the cell culture media in the 6-well plates and add 1 mL of warm (37 °C) trypsin-EDTA to each well.
Incubate (at 37 °C with 5% CO2) for 5 min.
Gently pipette the trypsin-EDTA to detach and dissociate the cells and transfer the cell suspension to a 15 mL Falcon tube.
Add 4 mL of the complete cell culture media to the cell suspension to neutralize the trypsin.
Centrifuge the cells at 300× g for 5 min.
Discard the supernatant, add 1 mL of fresh complete cell culture media, and resuspend the cells.
Count the cells using a hemocytometer and, if necessary, dilute the cells to obtain a final cell suspension concentration of 200 cells/µL.
Add 20,000 HeLa cells in each LEPD by pipetting 100 μL of the cell suspension solution.
Pipette 100 μL of additional complete cell culture media into each well.
Culture the cells on the membrane surface overnight in an incubator (at 37 °C with 5% CO2) to promote cell adhesion and tight cell membrane and nanopore contact.
Electroporate the adhered cells on the LEPDs the next day.
Note that this process can be adapted for various adherent and suspension cell types. The surface treatment, cell seeding, and culture conditions must be optimized accordingly.
Delivery into suspension cells
Check the cells in the microscope to ensure they look viable and have proper morphology.
Count the cell density using a hemocytometer or automated cell counter.
Take the appropriate volume of cells from the media to plate between 30,000 and 50,000 cells per device (e.g., 24 devices require ~1.2 million cells).
Take the appropriate volume of media that contains the desired number of cells calculated from the cell density obtained from step D2 and place the cells in a Falcon tube.
Place the tube containing the cells in a centrifuge and add a counterweight to balance. Centrifuge at 150× g for 5 min.
Gently remove the cell media from the Falcon tube, leaving the pellet of cells. Add electroporation buffer to the tube: 100 μL of electroporation buffer per device (2.4 mL for 24 devices). Mix the cells in the electroporation buffer.
Dispense 100 μL of electroporation buffer containing the cells in each device inside of a well-plate.
Place the well plate that contains the devices with cells into the centrifuge. Place a counterweight (a well plate with ~200 μL fluid in each well) in the opposite chamber of the centrifuge and centrifuge at 150× g for 5 min.
Optimization of plasmid delivery
Prepare eGFP plasmid solutions of concentration ranging from 100 ng/µL to 350 ng/µL with a step size of 50 ng/µL. The minimum volume of each solution should be 5 µL.
Take the electroporation buffer out of 4 °C storage and allow it to reach room temperature.
Transfer the LEPDs with the cells from the incubator into a biosafety cabinet. Carefully pipette out all the media from the LEPDs and add 200 µL of the electroporation buffer to each LEPD.
If using non-adherent cells, place the LEPDs from step E3 in a well plate and centrifuge them at 150× g for 5 min.
Using a 10 µL pipette, place a 5 µL droplet of the 100 ng/µL eGFP plasmid solution on the bottom gold electrodes of the wells of column 1 of each row of the 24-well LEPD system. Repeat this for the higher concentration eGFP plasmid solutions for the remaining columns going up in concentration to 350 ng/µL for the sixth column (Video 3).
Video 3. Loading, transmitting, and voltage optimization of cargo delivery
Using a pair of sterile tweezers, gently place the LEPDs with the cultured cells onto the wells with the droplet of delivery plasmid on the bottom gold electrode with one LEPD device per well, as shown in Video 3.
Set the pulse conditions on the electronics software (provided by Infinitesimal LLC) to a bi-level pulse with the following parameters: V1 ranging from 10 to 40 V, V2 fixed at 10 V, T1 at 0.5 ms, T2 ranging from 0.5 to 2.5 ms, frequency set at 20 Hz, and pulse count between 100 and 800. For guidance on using the software, refer to Video 3.
Connect the positive terminal of the electronics box to row 1 of the top plate and the negative terminal to the bottom plate using cables with alligator clip connectors.
Check the resistance of the system using the Infinitesimal LLC software and check for loose connections if the resistance is not in the 1–2 MΩ range and is unstable.
Use the electronics software to apply the pulse; the cells in the devices in row 1 will be subjected to localized electroporation.
After the pulse (~2 min) ends, switch the positive terminal to the next row and repeat steps E7–10. These steps need to be repeated until cells in all four rows are subjected to electroporation.
Disconnect the electrical connections, lift the top plate, and transfer all the LEPDs into a new transparent well plate.
Carefully pipette out the electroporation buffer from all the devices, add fresh cell culture media to the LEPDs, and transfer them into the incubator.
This protocol for optimizing plasmid concentration along the columns and pulse parameters along the rows can be repeated for as many combinations as we want to try.
Automated imaging
To clean the bottom of the LEPD membrane that comes in contact with the delivery reagents, prepare a 12-well plate by dispensing 1 mL of PBS in each well. Dip the LEPDs in the wells without fully submerging the devices (three wells sequentially) and transfer the clean LEPDs to a transparent 24-well plate for imaging.
To quantify cell viability and delivery efficiencies, prepare a solution of Hoechst 33342 nuclear stain in PBS (0.1 mg/mL), gently aspirate the fluid from each well (cell media or electroporation buffer) without completely drying the well, and place 100 µL of Hoechst solution prepared in the well for 10 min.
Gently aspirate the Hoechst solution, wash thrice with PBS, and dispense 150 µL of PBS for imaging.
Place the well plate containing the LEPDs in a motorized microscope stage.
Program the stage to move to the center of each well in the plate and use an objective lens with 10× or 20× magnification to capture images of the stained nuclei using a DAPI filter.
Focus on the cells manually using the coarse and fine focus knobs or by programming the microscope with an autofocus routine. Briefly, the autofocus routine consists of calculating the focus score (e.g., normalized variance, Laplacian, or log-histogram) for a Z-stack of images acquired in a single field-of-view, moving to the Z-plane with the highest score, and iteratively reducing the step size and scanning range until achieving optimal focus. To reduce the time of the routine, the user manually focuses on the first well to set a reference Z-plane to be used for the subsequent wells, since the difference in focus is small between wells.
Obtain images with multiple fluorescence and brightfield filters for each field of view. The choice of filters depends on the excitation and emission characteristics of the fluorescent probe to be examined.
For each LEPD, acquire images at multiple fields of view by moving manually or by programming the microscope to move to different locations within each well.
The representative transfection images of adherent and suspended cell lines, HeLa and K562, after plasmid (pmax GFP) delivery are presented in Figure 3A–B. The transfection efficiency appears to be lower in K562 cells, likely due to their non-adherent nature, which hampers efficient contact with the porous cell membrane. Figure 4A–B provides the viabilities images taken 24 h post-transfection for HeLa and HEK 293T cell lines. Further discussion of these images can be found in Patino et al. [9].
Figure 3. Representative efficiency results. Representative images of successful transfection of fluorescent protein-encoding plasmids into (A) HeLa (adherent) and (B) K562 (cells in suspension) transfected with a pmax GFP–encoding plasmid. Scale bars represent 100 µm. Variation of transfection efficiency for HEK 293T cells electroporated in cell culture media (DMEM) and EP buffer using a localized electroporation device (LEPD) with respect to (C) pulse voltage and (D) pulse duration. All error bars indicate the standard error of the mean (SEM) of triplicate samples, n cell > 100 per sample for all bar plots. All transfection efficiencies are normalized with respect to the highest value of efficiency in each plot. The highest efficiencies for plots in A and B are 71.6% and 63.6%, respectively. Figures are adapted from Patino et al. [9].
Data analysis
Image segmentation and data analysis
To identify the cells in the images, a fully convolutional network (FCN) [25] with a U-Net architecture [26] was trained using images and corresponding labels of various cell types and imaging modalities (e.g., fluorescence and phase contrast). The FCN network consists of an encoder-decoder architecture containing 20 hidden layers (e.g., convolution, pooling, and up-convolution) that enables the classification of objects in the images (e.g., cell exterior, nucleus) with pixel-level resolution [20,27].
To train the U-Net for a specific cell type of interest, images of cells were manually labeled using drawing tools in image manipulation software. To facilitate labeling of the cell exterior or internal cell compartments of interest, the cells were stained with fluorescent dyes (e.g., Hoechst nuclear dye or Calcein AM cytosolic dye) and imaged as described in section F.
The images and their corresponding ground truth labels were split into training, validation, and test sets to cross-validate the training process and prevent overfitting.
A weighted soft-max cross-entropy loss function was used to classify each pixel into three categories (interior, exterior, and boundary).
The U-Net was optimized using stochastic gradient descent (momentum = 0.9, learning rate = 1 × 10-4). The training was performed using a graphic processing unit (GPU: NVIDIA) to expedite the process for a large network containing more than 7.5 million trainable parameters.
To assess the performance of the model, the area overlap between the predicted objects and the ground truth labels was measured and used to determine true positives (TP), false positives (FP), true negatives (TN), and false negatives (FN). From these values, the precision, recall, and F1 scores could be calculated as follows:
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For analysis of viability, delivery, and transfection efficiencies, the Hoechst nuclear images were used to segment the nuclei of the cells to determine the number of cells in each image. Moreover, the segmented nuclei were used as a mask to measure the fluorescence intensity of the other color channels (e.g., GFP filter for eGFP transfection and TexasRed filter for quantification of dead cells using propidium iodide). The fluorescence intensity for the respective filters was compared to negative control samples to determine the threshold for a positive signal. Viability and transfection efficiency were calculated as follows:
viability = 1 - (Ndead/Ncells)
efficiency = Ntransfected/Ncells
To extract measurements from the segmented images, a cell analysis software (CellProfiler) was used to obtain numerous features (e.g., shape, intensity, texture) from each cell.
Figure 3C and D illustrate the efficiencies of GFP plasmid delivery into HEK 293T cells under varying voltages and pulse durations, and across different buffer conditions. The post-transfection viability results of HEK 293T cells with different voltages are shown in Figure 4C, where viability remains high between 10 and 20 V and declines at higher voltages of 30 and 40 V in both EP and DMEM buffer. Figure 4D represents the viability results for HeLa, HEK, and K562 cell lines post-transfection. Additional data and further discussions are available in Patino et al. [9].
The features were transformed using the generalized logarithm (glog) method and standardized using the median and median absolute deviation (MAD) to obtain the robust Z score (R.Z. score). In contrast to the Z-score, the R.Z. score is not sensitive to outliers.
To analyze the features, correlation matrices and 2D feature projection maps (U-Map, t-SNE) can be applied.
Figure 4. Representative viability results. Representative composite fluorescence micrographs of propidium iodide (PI) viability assay conducted on (A) HeLa and (B) HEK 293T 24 h after being treated with the localized electroporation device (LEPD); pseudo colors: magenta—Hoechst, yellow—PI. Scale bar = 100 µm. (C) Viability after plasmid delivery across different voltages: Bar plots show the mean viability of HEK 293T cells transfected using the LEPD in both DMEM and EP buffer in the voltage range 10–40 V. (D) Viability of various continuous cell lines 24 h after treatment with LEPD and LIPO. All error bars indicate the standard error of the mean of triplicate samples, ncell > 100 per sample for all bar plots. Figures are adapted from Patino et al. [9].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Patino et al. [9]. Multiplexed high-throughput localized electroporation workflow with deep learning–based analysis for cell engineering. Sci Adv. (Figures 4–7).
Acknowledgments
Research reported in this publication was supported by the NIH R21 award number 1R21GM132709-01. This protocol is derived from the original paper [9].
Competing interests
The authors declare that they have no competing interests.
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Boukany, P. E., Morss, A., Liao, W. c., Henslee, B., Jung, H., Zhang, X., Yu, B., Wang, X., Wu, Y., Li, L., et al. (2011). Nanochannel electroporation delivers precise amounts of biomolecules into living cells. Nat Nanotechnol. 6(11): 747–754.
Kang, W., Yavari, F., Minary-Jolandan, M., Giraldo-Vela, J. P., Safi, A., McNaughton, R. L., Parpoil, V. and Espinosa, H. D. (2013). Nanofountain Probe Electroporation (NFP-E) of Single Cells. Nano Lett. 13(6): 2448–2457.
Cao, Y., Ma, E., Cestellos-Blanco, S., Zhang, B., Qiu, R., Su, Y., Doudna, J. A. and Yang, P. (2019). Nontoxic nanopore electroporation for effective intracellular delivery of biological macromolecules. Proc Natl Acad Sci USA. 116(16): 7899–7904.
Kang, W., Giraldo-Vela, J. P., Nathamgari, S. S. P., McGuire, T., McNaughton, R. L., Kessler, J. A. and Espinosa, H. D. (2014). Microfluidic device for stem cell differentiation and localized electroporation of postmitotic neurons. Lab Chip 14(23): 4486–4495.
Mukherjee, P., Patino, C. A., Pathak, N., Lemaitre, V. and Espinosa, H. D. (2022). Deep Learning‐Assisted Automated Single Cell Electroporation Platform for Effective Genetic Manipulation of Hard‐to‐Transfect Cells. Small 18(20): e202107795.
Xie, X., Xu, A. M., Leal-Ortiz, S., Cao, Y., Garner, C. C. and Melosh, N. A. (2013). Nanostraw–Electroporation System for Highly Efficient Intracellular Delivery and Transfection. ACS Nano. 7(5): 4351–4358.
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Mukherjee, P., Berns, E. J., Patino, C. A., Hakim Moully, E., Chang, L., Nathamgari, S. S. P., Kessler, J. A., Mrksich, M. and Espinosa, H. D. (2020). Temporal Sampling of Enzymes from Live Cells by Localized Electroporation and Quantification of Activity by SAMDI Mass Spectrometry. Small 16(26): e202000584.
Patino, C. A., Mukherjee, P., Berns, E. J., Moully, E. H., Stan, L., Mrksich, M. and Espinosa, H. D. (2022). High-Throughput Microfluidics Platform for Intracellular Delivery and Sampling of Biomolecules from Live Cells. ACS Nano. 16(5): 7937–7946.
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Article Information
Publication history
Received: May 23, 2024
Accepted: Jun 3, 2024
Available online: Jul 5, 2024
Published: Jul 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Biochemistry > Protein > Expression
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5,038 | https://bio-protocol.org/en/bpdetail?id=5038&type=0 | # Bio-Protocol Content
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Slot Blot Analysis of Intracellular Glyceraldehyde-Derived Advanced Glycation End Products Using a Novel Lysis Buffer and Polyvinylidene Difluoride Membrane
TT Takanobu Takata
HM Hiroki Murayama
TM Togen Masauji
Published: Vol 14, Iss 14, Jul 20, 2024
DOI: 10.21769/BioProtoc.5038 Views: 605
Reviewed by: Alessandro Didonna Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Diabetology & Metabolic Syndrome Jun 2020
Abstract
Advanced glycation end products (AGEs) are formed through the reaction/modification of proteins by saccharides (e.g., glucose and fructose) and their intermediate/non-enzymatic products [e.g., methylglyoxal and glyceraldehyde (GA)]. In 2017, Dr. Takanobu Takata et al. developed the novel slot blot method to quantify intracellular GA-derived AGEs (GA-AGEs). Although the original method required nitrocellulose membranes, we hypothesized that the modified proteins contained in the AGEs may be effectively probed on polyvinylidene difluoride (PVDF) membranes. Because commercial lysis buffers are unsuitable for this purpose, Dr. Takata developed the slot blot method using an in-house-prepared lysis buffer containing 2-amino-2-hydromethyl-1,3-propanediol (Tris), urea, thiourea, and 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) that effectively probes AGEs onto PVDF membranes. The slot blot method also entails the calculation of Tris, urea, thiourea, and CHAPS concentrations, as well as protein and mass to be probed onto the PVDF membranes. GA-AGE-modified bovine serum albumin (BSA, GA-AGEs-BSA) is used to draw a standard curve and perform neutralization against a non-specific combination of anti-GA-AGEs antibodies, thereby enabling the quantification of GA-AGEs in cell lysates. This paper presents the detailed protocol for slot blot analysis of intracellular GA-AGE levels in C2C12 cells.
Key features
• This protocol leverages the idea that advanced glycation end products are modified proteins.
• The lysis buffer containing Tris, urea, thiourea, and CHAPS enables probing proteins onto PVDF membranes.
• Intracellular GA-AGE levels may be quantified for some cell types using polyclonal anti-GA-AGE antibodies and standard GA-AGE-modified BSA.
• The lysis buffer may be simultaneously prepared with the cell lysate.
• There is no limit to the type of cultured cells used in the preparation of cell lysate.
Keywords: Advanced glycation end product Glyceraldehyde GA-AGE Polyvinylidene difluoride 2-amino-2-hydromethyl-1 3-propanediol Urea Thiourea 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate Slot blot analysis
Graphical overview
Preparation of sample and slot blot analysis with Dr. Takata’s lysis buffer and polyvinylidene difluoride (PVDF) membranes.
Background
The quantification of intracellular advanced glycation end products (AGEs) is particularly useful in the fields of biochemistry, molecular biology, and protein engineering [1]. AGEs are a type of modified proteins that can be extracted from cultured cells and tissues [2] and indirectly measured using the enzyme-linked immunosorbent assay (ELISA) [3,4]. However, some researchers prefer using a slot blot approach [5–7], because identifying or quantifying some types of AGEs may be difficult using ELISA. Previous slot blot methods for the quantification of AGEs were limited by their use of nitrocellulose membranes and radioimmunoprecipitation (RIPA) buffer [5,6]. Although nitrocellulose membranes are useful in column chromatography, polyvinylidene difluoride (PVDF) membranes are more durable and show greater protein adsorption ability [8,9]. However, there is a lack of suitable lysis buffers for probing proteins onto PVDF membranes. Although RIPA buffer is used in western blotting, it is less suitable in slot blot analyses with PVDF membranes [8,9]. Consequently, Dr. Takanobu Takata developed a lysis buffer containing 2-amino-2-hydromethyl-1,3-propanediol (Tris), urea, thiourea, and 3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate (CHAPS) to quantify intracellular AGEs using the slot blot analysis [8]. The improved probing efficacy of this lysis buffer may be related to protein carbamoylation [8,9] and the absence of Triton-X, which can inhibit the probing of proteins onto PVDF membranes [8,9]. In 2017, the novel slot blot method was used to accurately quantify intracellular glyceraldehyde (GA)-derived AGEs (GA-AGEs) [10]. From 2017 to 2022, Takata et al. applied this method in the quantification of intracellular GA-AGEs in cells and tissue lysates of the pancreas [10,11], heart [12,13], skeletal muscles [14], liver [15–19], and bone [20] using (i) the novel lysis buffer, (ii) standard GA-AGEs modified bovine serum albumin (BSA), and (iii) neutralization using anti-GA-AGEs antibodies. In this method, the standard GA-AGEs-BSA is used to estimate standard curves [10]; neutralization using a non-specific combination of polyclonal anti-GA-AGEs antibodies [21] avoids biases in the quantification of GA-AGEs [8, 10–17]. We previously applied this protocol to a type of GA-AGEs [10–17] known as toxic AGEs (TAGE) [21]. However, our slot blot may be applied to various types of AGEs, including 1,5-anhydro-fructose AGEs [22] and modified proteins [8,9], even though various AGE modifications may occur in one protein type or molecule [23]. This study presents a validated protocol for the slot blot analysis of intracellular GA-AGE levels using Dr. Takata’s lysis buffer and a PVDF membrane.
Materials and reagents
Biological materials
C2C12 cell line (KAC, Kyoto, catalog number: EC91031101-F0)
The C2C12 cell line is an immortalized mouse myoblast cell line (ATCC, catalog number: CRL1772)
Reagents
Dulbecco’s modified Eagle’s medium (DMEM) (Sigma-Aldrich, catalog number: D6046-500M)
Penicillin/streptomycin solution (FUJIFILM Wako Pure Chemical Corporation, catalog number: 168-23191)
Fetal bovine serum (FUJIFILM Wako Pure Chemical, catalog number: 554-04855)
Glyceraldehyde (Nacalai Tesque, catalog number: 17014-81)
Phosphate-buffered saline (PBS) without Ca2+ and Mg2+ [PBS(-)], 20× (LSI Medience, catalog number: PM102-PN)
2-amino-2-hydromethyl-1,3-propanediol (Tris base) (Tris) (FUJIFILM Wako Pure Chemical, catalog number: 011-20095)
Urea (FUJIFILM Wako Pure Chemical, catalog number: 217-01215)
Thiourea (FUJIFILM Wako Pure Chemical, catalog number: 206-17355)
CHAPS (DOJINDO, Kumamoto, Japan, catalog number: 349-04722)
Methanol (FUJIFILM Wako Pure Chemical, catalog number: 131-01826)
Protease Inhibitor cocktail cOmplete Tablets EDTA-free, EASY pack (Roche, catalog number: 04-693-132-001)
BSA fraction IV (FUJIFILM Wako Pure Chemical, catalog number: 019-2329)
Bradford dye reagent (Takara Bio, catalog number: T9310A-1)
Skim milk for immunoassay (Nacalai Tesque, catalog number: 31149-75)
Tween-20 (GE Healthcare, catalog number: 17-1316-01)
Polyclonal anti-GA-AGE antibodies (purchased from Prof. Masayoshi Takeuchi, Department of Advanced Medicine, Medical Research Institute, Kanazawa Medical University, Uchinada, Japan; 920-0293).
Note: Prof. Takeuchi successfully prepared the polyclonal antibody for TAGE, despite having limited information on the structure of TAGE [21]. Takeuchi et al. reported the hypothetical structure of TAGE in 2023 [24]. The antibody is preserved at -80 °C.
GA-AGE-BSA, 10 mg/mL (or TAGE-BSA, purchased from Prof. Takeuchi)
Note: Despite limited information on the intra- and intermolecular structure of TAGE-BSA, the polyclonal anti-GA-AGE antibody (Reagent 17) developed by Prof. Takeuchi could be probed against the antigen recognition site in TAGE-BSA [21]. The GA-AGEs-BSA was dissolved in PBS and preserved at -30 °C.
Horseradish peroxidase (HRP)-conjugated molecular weight marker (Bionexus, catalog number: BNPM41)
Polyclonal goat anti-rabbit immunological HRP-conjugated antibody (DAKO, catalog number: REF0448)
ImmunoStar LD kit (FUJIFILM Wako Pure Chemical, catalog number: 292-69903)
Milli-Q ultrapure water
Note: Purchased with an RFU554CA ultrapure water system (Advantech Toyo, Tokyo, Japan) at the Kanazawa Medical University.
Solutions
Medium for cell culture (see Recipes)
PBS(-) 1× (see Recipes)
Solution A (see Recipes)
Solution B (see Recipes)
Solution C (see Recipes)
Solution D (see Recipes)
20 mg/mL BSA in Solution D (20 mg/mL BSA solution) (see Recipes).
Diluted BSA solution (see Recipes).
Diluted GA-AGEs-BSA solution (see Recipes)
HRP-conjugated molecular weight marker solution (see Recipes)
PBS-T (see Recipes)
5% SM-PBS-T (see Recipes)
0.5% SM-PBS-T (see Recipes)
Primary antibody solution (see Recipes)
Neutralized primary antibody solution (see Recipes)
Secondary antibody solution (see Recipes)
Recipes
Medium for cell incubation
Mix the reagents on a laminar flow hood.
Note: Because this step can easily be performed in-house prior to the experiment, we do not provide a detailed description here. Complete information may be found in our previous studies [10–16]. Media can be stored in the general refrigerator (4 °C).
Reagent Volume Recommended storage condition
DMEM 450 mL 4 °C
Penicillin/streptomycin solution 5 mL -30 °C
Fetal bovine serum 50 mL -30 °C
Total 505 mL
PBS(-) (1×)
Room temperature refers to 22–28 °C.
Reagent Volume Recommended storage condition
PBS(-) (20×) 50 mL Room temperature
Milli-Q water 950 mL Room temperature
Total 1,000 mL
Solution A
Using a 50 mL polypropylene centrifuge tube and serological pipet, dissolve Tris in Milli-Q water (10 mL) to prepare 1 mol/L (M) Tris solution. Solution A may be stored at -30 °C for 6 months.
Note: The pH of the Tris solution was approximately 9.4. Only Tris powder was dissolved in Milli-Q-water, without adding other reagents (e.g., hydrochloride).
Reagent Quantity/volume Recommended storage condition Final concentration
Tris 1.21 g Room temperature 1 mol/L (M)
Milli-Q water 10 mL Room temperature
Total 10 mL
Solution B
Dissolve one tablet of protease inhibitor cocktail (cOmplete Tablets EDTA free) in Milli-Q water (2 mL) within a microcentrifuge tube; transfer the Milli-Q water to the tube using a 100–1,000 μL Gilson PIPETMAN. We recommend Solution B (2 mL) to be prepared and used immediately with every experiment. However, Solution B can be preserved at -30 °C for three months if it is reused. Approximately 100 μL of Solution B were preserved.
Reagent Quantity/Volume Recommended storage condition Final concentration
Protease inhibitor cocktail 1 tablet 4 °C 1 tablet/2 mL
Milli-Q water 2 mL Room temperature
Total 2 mL
Solution C
Solution C contains 30 mM Tris, 7 M urea, 2 M thiourea, and 4% CHAPS. Mix these reagents in a 50 mL polypropylene centrifuge tube and add Milli-Q water to a final volume of 20 mL. Treat 1 M Tris with Milli-Q water using a 200–1,000 μL Gilson PIPETMAN.
Note: The reagents are added to the centrifuge tube in no specific order. At room temperature, urea and thiourea are not easily dissolved in Milli-Q water; accordingly, we recommend using a vortex system (e.g., Vortex-Genie 2) at room temperature. We do not recommend dissolution in a 37 °C CO2 incubator or water bath because urea and thiourea may produce cyanate or isocyanic acid at high temperatures [9]. Solution C can be preserved at -30 °C for three months, though we recommend preparing it immediately before use.
Reagent Quantity/Volume Recommended storage condition Final concentration
1 M Tris 0.6 mL Room temperature 30 mM
Urea 8.40 g Room temperature 7 M
Thiourea 3.04 g Room temperature 2 M
CHAPS 0.80 g Room temperature 4 w/v (%)
Milli-Q water 20 mL Room temperature
Total 20 mL
Solution D
Prepare Solution D by mixing Solutions B and C at a ratio of 1:9. Transfer Solution C using the 200–1,000 μL Gilson PIPETMAN. Transfer Solution D using a 10 mL serological pipette. Solution D contains 27 mM Tris, 6.3 M urea, 1.8 M thiourea, and 3.6 v/w (%) CHAPS.
Note: We recommend dissolution using a vortex system (e.g., Vortex-Genie 2) at room temperature. Solution D can be preserved at -30 °C for three months, though we recommended preparing it immediately before use. Solution D was used as lysis buffer to prepare the cell lysate [8–17], which was prepared from C2C12 cells treated with glyceraldehyde [14]. Although “Modified Solution C” containing 30 mM Tris, 7 M urea, 2 M thiourea, 4% CHAPS, and 4% Solution B may also be used as lysis buffer [18–20,22], we recommend that Solution D be used because it contains sufficient urea, thiourea, and CHAPS to generate the carbamylated AGE proteins [8,9]. The cell lysates and unused Solution D should be preserved at -30 °C until later use in the slot blot experiment. The condition of the unused Solution D and the cell lysates should be arranged.
Reagent Volume Final concentration
Solution B 2 mL 10%
Solution C 18 mL 90%
Total (optional) 20 mL
BSA–Solution D mixture (20 mg/mL BSA solution)
Dissolve 20 mg of BSA in Solution D (10 mL) within a 50 mL polypropylene centrifuge tube. Transfer Solution D using a 200–1,000 μL Gilson PIPETMAN.
Note: BSA cannot be easily dissolved in Solution D; thus, we recommend using a vortex system (e.g., Vortex-Genie 2) at room temperature. The presence of undissolved BSA can be confirmed by the presence of a gelatinous substance. Dissolution by vortex should not be performed at high temperatures (e.g., 37 °C) [9].
Note: We do not recommend commercial BSA for use in this experiment. An in-house solution should preferably be prepared. We believe that BSA fraction IV is suitable for the Bradford method in our protocol.
Reagent Quantity/Volume Final concentration Recommended storage condition
BSA 20 mg 2.0 mg/mL (μg/μL) 4 °C
Milli-Q water 10 mL Room temperature
Total 10 mL
Diluted BSA solution
Dilute 20 mg/mL BSA solution to 0.0625, 0.125, 0.25, 0.5, 1.0, and 1.5 μg/μL in a 1.5 mL microcentrifuge tube; transfer BSA using a 50–200 μL Gilson PIPETMAN. We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) at room temperature. The BSA solution should be made and used immediately with every experiment. BSA solution should be aliquoted (approximately 100 μL) and preserved at -80 °C if it will be reused. However, we recommend preparing and using a fresh BSA solution per experiment.
Note: Prepare from 20 mg/mL BSA solution (Recipe 7).
Volume of reagents Volume of Solution D Total volume Final concentration
150 μL of 2.0 μg/μL BSA solution 50 μL 200 μL 1.5 μg/μL
100 μL of 2.0 μg/μL BSA solution 100 μL 200 μL 1.0 μg/μL
100 μL of 1.0 μg/μL BSA solution 100 μL 200 μL 0.5 μg/μL
100 μL of 0.5 μg/μL BSA solution 100 μL 200 μL 0.25 μg/μL
100 μL of 0.25 μg/μL BSA solution 100 μL 200 μL 0.125 μg/μL
100 μL of 0.125 μg/μL BSA solution 100 μL 200 μL 0.0625 μg/μL
Diluted GA-AGEs-BSA solution
Dilute 10 mg/mL GA-AGEs-BSA in PBS(-) within a 1.5 mL microcentrifuge tube; transfer the solution using a 50–200 μL or 1–10 μL Gilson PIPETMAN.
Note: We recommend the diluted GA-AGEs-BSA solution to be prepared before being used.
Volume of reagents Volume of PBS(-) Total volume Final concentration
2.0 μL of 10 mg/mL GA-AGEs-BSA 98 μL 100 μL 200 ng/μL
20 μL of 200 ng/μL GA-AGEs-BSA 60 μL 80 μL 50 ng/μL
60 μL of 50 ng/μL GA-AGEs-BSA 40 μL 100 μL 30 ng/μL
30 μL of 50 ng/μL GA-AGEs-BSA 70 μL 100 μL 15 ng/μL
10 μL of 50 ng/μL GA-AGEs-BSA 90 μL 100 μL 5 ng/μL
10 μL of 15 ng/μL GA-AGEs-BSA 90 μL 100 μL 1.5 ng/μL
10 μL of 5 ng/μL GA-AGEs-BSA 90 μL 100 μL 0.5 ng/μL
HRP-conjugated molecular weight marker solution
Dilute the HRP-conjugated molecular weight marker in PBS(-) within a 1.5 mL microcentrifuge tube; transfer the solution using the 200–1,000 μL or 1–10 μL Gilson PIPETMAN.
Note: We recommend dissolution to be performed using a vortex system (e.g., Vortex-Genie 2) at room temperature and the diluted GA-AGEs-BSA solution to be prepared at the time of use.
Reagent Volume Recommended storage condition
HRP-conjugated molecular weight marker 3 μL -30 °C
PBS(-) 197 μL Room temperature
Total 200 μL
PBS-T
Dissolve Tween-20 in PBS(-). Transfer Tween-20 using the 200–1,000 μL Gilson PIPETMAN.
Reagent Volume Recommended storage condition
Tween-20 0.5 mL Room temperature
PBS(-) 1,000 mL Room temperature
Total 1,000.5 mL
5% SM-PBS-T for immunoassay
Dissolve skim milk for immunoassay in PBS-T within a 50 mL polypropylene centrifuge tube; transfer PBS-T using the 10 mL serological pipette.
Note: We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) at room temperature and that 5% SM-PBS-T be prepared before use. However, the solution may be preserved at 4 °C for two days.
Reagent Quantity/Volume Recommended storage condition
Skim milk for immunoassay 2.5 g 4 °C
PBS-T 50 mL Room temperature
Total 50 mL
0.5% SM-PBS-T
Dilute 5% SM-PBS-T to 0.5% SM-PBS-T in a 50 mL polypropylene centrifuge tube; transfer PBS-T using the 10 mL serological pipette.
Note: We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) at room temperature and that 5% SM-PBS-T be prepared before use, which may occur at room temperature. However, the solution may be preserved at 4 °C for two days.
Reagent Volume Recommended storage condition
5% SM-PBS-T 5 mL Room temperature
PBS-T 45 mL Room temperature
Total 50 mL
Primary antibody solution
Mix 5 mL of 0.5% SM-PBS-T (Recipe 13) using a 10 mL serological pipette with 5 μL of polyclonal anti-GA-AGE antibody (using a 2–20 μL Gilson PIPETMAN) within a 15 mL polypropylene centrifuge tube.
Note: We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) for 10 s at room temperature. The polyclonal anti-GA-AGE antibody, which was preserved at -80 °C, should first be defrosted at 0 °C (in ice) and may then be preserved at 4 °C for half a year.
Reagent Volume Recommended storage condition Dilution ratio
Polyclonal anti-GA-AGEs antibody 5 μL -80 °C 1:1,000
0.5% SM-PBS-T 5.0 mL (5,000 μL) Room temperature
Total (optional) 5.005 mL (5,005 μL)
Neutralized primary antibody solution
Mix 5 μL of polyclonal anti-GA-AGEs antibody (2–20 μL Gilson PIPETMAN) with 125 μL of 10 mg/mL GA-AGEs-BSA (50–200 μL Gilson PIPETMAN) and 4.875 mL of 0.5% SM-PBS-T (10 mL serological pipette) within a 15 mL polypropylene centrifuge tube.
Note: We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) for 10 s at room temperature. Polyclonal anti-GA-AGEs and GA-AGEs-BSA were preserved at -80 °C and -30 °C, respectively, and defrosted on ice. We recommend that 10 mg/mL GA-AGEs-BSA be divided into 130 μL aliquots and preserved at -30 °C in advance. Moreover, we recommended the divided and preserved GA-AGEs-BSA to be used, and one of them should be for one experiment (they should not be refrozen and reused).
Reagent Volume Recommended storage condition Dilution ratio/Final concentration
Polyclonal anti-GA-AGEs antibody 5 μL -80 °C 1:1,000
10 mg/mL GA-AGEs-BSA 125 μL -30 °C 1/40 (250 μg/mL)
0.5% SM-PBS-T 4.875 mL (4,875 μL) Room temperature
Total 5.005 mL (5,005 μL)
Secondary antibody solution
Mix 5 mL of 0.5% SM-PBS-T (10 mL serological pipette) with 2.5 μL of polyclonal goat anti-rabbit immunological HRP-conjugated antibody (1–10 μL Gilson PIPETMAN) within a 15 mL polypropylene centrifuge tube. We recommend that dissolution be performed using a vortex system (e.g., Vortex-Genie 2) for 10 s at room temperature.
Reagent Volume Recommended storage condition Dilution ratio
Polyclonal goat anti-rabbit immunological HRP-conjugated antibody 2.5 μL 4 °C 1:2,000
0.5% SM-PBS-T 5 mL (5000 μL) Room temperature
Total 5.0025 mL (5002.5 μL)
Laboratory supplies
60 mm dish (BM Equipment, catalog number: 93060)
1.5 mL microcentrifuge tube (Thermo Fisher Scientific, catalog number: 3451)
2.0 mL microcentrifuge tube (Watson, catalog number: 132-6201)
50 mL polypropylene centrifuge tube (TrueLine; Nippon Genetics, catalog number: TR2004)
15 mL polypropylene centrifuge tube (TrueLine; Nippon Genetics, catalog number: TR2000)
10 mL serological pipette (BM Equipment, catalog number: 207500-SLP-10)
Rubber bulb for 10 mL serological pipette (AS ONE, catalog number: 6-356-4)
Disposable polypropylene tray (AS ONE, catalog number: 1-3145-03)
Dispenser (TPP, BM Equipment, catalog number: 99010)
1–10 μL pre-sterilized tip (10 μL) (BM Equipment, catalog number: W10-RS)
50–200 μL pre-sterilized tip (200 μL) (Watson, catalog number: 62-0887-34)
200–1,000 μL pre-sterilized tip (1,000 μL) (Watson, catalog number: 38688239)
96-well microplates (Becton Dickinson, catalog number: REF353072)
25 mL reagent reservoir (BM Equipment, catalog number: BM-0850-1)
PVDF membrane (pore size: 0.45 μm) (Merck Millipore, catalog number: IPVH00010)
Filter paper (9 cm × 12 cm) (Bio-Rad Laboratories, catalog number: 1620161)
Hybri-Bag (hard type) (Cosmo Bio, catalog number: S1001)
Gilson PIPETMAN (GILSON, model type: 100–1,000 μL)
Gilson PIPETMAN (GILSON, model type: 50–200 μL)
Gilson PIPETMAN (GILSON, model type: 2–20 μL)
Gilson PIPETMAN (GILSON, model type: 1–10 μL)
Nichipet 7000 range: 50–200 μL (Nichiryo, Tokyo, Japan)
Equipment
Ultrapure Water System (Advance Toyo, model: RFU554CA)
Flask-trap aspirator (1,000 mL) (Biosan, Riza, Latvia; model number: FTA-1)
Centrifuge (Eppendorf, model: 5415R)
Microplate reader (Bio-Rad, model: iMark)
Vortex-Genie 2 (M&S Instruments, catalog number: 33230217)
Imager (M&S Instruments, model: Fusion FX)
AS-200 Heat sealer (AS ONE, catalog number: H221049/01167C)
Seesaw shaker (BIO CRAFT, Tokyo, model: BC-700)
Aspirator with water pump (AS ONE, catalog number: 1-689-02)
Bio-Dot SF microfiltration apparatus (48 lanes) (Bio-Rad; Figure 1)
Figure 1. Bio-Dot SF microfiltration apparatus (48 lanes) containing sample template and equipped with sealing screws, sealing gasket, gasket support plate, vacuum manifold, and tubing with flow valve
Software and datasets
Excel software (Microsoft, Redmond, WA, USA. version 2010, 2013, 2016)
Note: The software was installed on a standard personal computer (OS: Windows 7, 10).
FUSION FX software (M&S Instruments, version: 17.03)
Note: The software was installed on a standard personal computer (OS: Windows 7) with Fusion FX imager (M&S Instruments) and was used since November 2017 at Kanazawa Medical University [11,13–16,22]. However, other chemiluminescence imagers and software may be used [10,12,17–20].
Stat FX software (Artech, Osaka, Japan, version: 6)
Note: When the slot blot analysis was performed in more than three independent experiments, Statflex (version 6) was used to perform statistical analysis [10–16,18–20].
Procedure
Incubation of C2C12 cell line and treatment with glyceraldehyde
Note: Here, we only provide the simplified procedure for the slot blot analysis. A detailed description may be found in the Materials and Methods in Takata et al. [14] (DOI: 10.1186/s13098-020-00561-z).
Seed 1.9 × 104 cells/cm2 onto a 60 mm dish and incubate in a CO2 incubator for 24 h on DMEM supplemented with penicillin/streptomycin and fetal bovine serum.
After changing the medium, treat the cells with 0, 0.5, 1, 1.5, and 2 mM glyceraldehyde and incubate in the CO2 incubator for 24 h.
Note: The medium should be refreshed 24 h after seeding.
Removing medium and washing cells with PBS(-)
Remove culture medium by decantation and aspirate the residual medium with a 1,000 mL trap-flask aspirator. Perform this step twice.
Note: After decantation, ~300 μL of the culture medium should remain.
Add 7.0 mL of PBS(-), decant, and aspirate the residual PBS(-) with the flask-trap aspirator. Perform this step twice.
Note: Transfer PBS(-) using a 10 mL serological pipette at 22–28 °C Decantation removes ~300 μL of PBS(-).
Preparation of cell lysates
Add 300 μL of Solution D into a 60 mm dish and scrape and move the cells into a 1.5 mL microcentrifuge tube.
Add cells to Solution D and harvest with a dispenser.
Incubate cells in Solution D on ice for 20 min, during which the cell suspension is subjected to five pipetting operations, repeated three times.
Note: This operation is performed with 5–6 min intervals.
Centrifuge cells at 10,000× g for 15 min at 4 °C with the 5415R centrifuge.
Collect the supernatants in the 1.5 mL microcentrifuge tube.
Note: Cell lysates are preserved at -80 °C. Both cell lysates and Solution D are preserved at -80 °C until the protein concentrations are measured, because the components of Solution D should remain in the same conditions.
Measurement of protein concentration in cell lysates (see General note 1)
Add 4 μL of BSA in Solution D (final 0–2.0 μg/μL) to a 96-well microplate (N = 2).
Add 4 μL of cell lysate (cells treated with 0, 0.5, 1, 1.5, and 2 mM glyceraldehyde) to a 96-well microplate (N = 2).
Add Bradford dye reagent to a 25 mL reagent reservoir using the 10 mL serological pipette.
Note: Bradford dye reagent should be preserved at 4 °C. However, we recommend it to be kept at 22–28 °C for 30–60 min before being added into BSA in Solution D and the cell lysates.
Transfer 200 μL of Bradford dye reagent using the Nichipet 7000 to the BSA in Solution D and cell lysate in the 96-well microplates.
After 5 and 10 min, measure absorbance (595 nm) using the iMark microplate reader.
PVDF membrane and filter papers incubated in methanol and/or PBS(-)
Cut a PVDF membrane into 9 cm × 12 cm sections and incubate in methanol for 1 min at room temperature.
Note: This operation should be performed in a fume hood to avoid exposure to methanol.
Submerge the PVDF membrane sections in PBS(-) at room temperature.
Submerge also three filter papers (9 cm × 12 cm) in PBS(-) at room temperature.
Incubate both the PVDF membrane sections and three filter papers in the PBS(-) for 1 h at room temperature.
Note: Because the PVDF membrane may be hydrophobic after incubation in methanol, excess methanol should be removed using the three filter papers, and the membrane sections should be sufficiently submerged in the PBS(-).
PVDF membrane and filter papers prepared for slot blot apparatus
Set the sealing gasket onto the vacuum manifold (Figure 1).
Set the gasket support plate onto the vacuum manifold (Figure 1).
Set each filter paper (a total of three) onto the gasket support plate (Figures 1, 2).
Set the PVDF membrane onto the filter papers (Figure 2).
Incubate both the PVDF membrane and filter papers in PBS(-) for 1 h at room temperature.
Fix the sample template with attached sealing screw to the sealing gasket and tighten the four screws (Figures 1–3).
Note: Any air between the PVDF membranes and filter papers should be removed. Do not dry the PVDF membrane before adding the PBS(-) in section G. If section G cannot be performed before the PVDF membrane is dried, a little PBS(-) may be added to it. Because the PVDF membrane may be hydrophobic after incubation in methanol, excess methanol should be removed using the three filter papers, and the membrane sections should be sufficiently submerged in the PBS(-). Ensure that the four screws are tightened appropriately.
Figure 2. PVDF membranes and filter papers set onto slot blot apparatus. The white closed square represents the PVDF membrane, the gray closed squares represent the filter papers, and the blue closed square represents the sealing gasket.
Figure 3. Slot blot apparatus with 48 lanes for PVDF membrane with sample template attached using sealing screws
Preparation of sample solution for slot blot
For the absolute quantification of the intracellular content of GA-AGEs in C2C12 [14], we selected samples containing 2 μg of protein.
Note: To prepare the sample solution against the PVDF lanes for both “Primary antibody” and “Neutralized primary antibody,” the samples were prepared in duplicate.
Calculate the volume of the 2 μg of protein samples and move to a 1.5 mL microcentrifuge tube. Consequently, the samples with lower protein concentrations were collected in a greater volume (maximum volume of Solution D in all samples).
Calculate the volume of Solution D to be added to the other samples to reach the maximum volume.
Note: If the volume of one sample (2 μg of protein, the lowest protein concentration) was 10 μL and that of the other samples (2 μg of protein each) was 1, 3, 5, and 7 μL, Solution D was added to a final volume of 10 μL (including the cell lysates). The 1–10 μL Gilson PIPETMAN should preferably be used.
After Solution D is added to the cell lysates, incubate for 5 min at room temperature.
Add PBS(-) to the cell lysates to a final volume of 200 μL for the slot blot analysis.
Note: The 50–200-μL Gilson PIPETMAN should preferably be used.
Preparation of standard GA-AGEs-BSA solution for slot blot
We prepared 2 μL of GA-AGEs-BSA solution (0, 0.5, 1.5, 5, 15, 30, and 50 ng/μL, see Recipe 9) in a 1.5 mL microcentrifuge tube.
Note: To prepare the standard GA-AGEs-BSA solution against the PVDF lanes for both “Primary antibody” and “Neutralized primary antibody,” the samples were prepared in duplicate. The 1–10 μL Gilson PIPETMAN should preferably be used.
Add Solution D against the GA-AGEs-BSA solution.
Note: If the volume of the sample solution is 10 μL, the same volume of Solution D must be added against the GA-AGEs-BSA solution.
After Solution D is added to the cell lysates, incubate for 5 min at room temperature.
Add PBS(-) to the cell lysates up to a final total volume of 200 μL for the slot blot analysis.
Note: The 50–200 μL Gilson PIPETMAN should preferably be used.
Preparation of HRP-conjugated marker solution for slot blot
We prepared 3 μL of the HRP-conjugated marker solution (Recipe 10) in a 1.5 mL microcentrifuge tube.
Note: To prepare the HRP-conjugated marker solution against the PVDF lanes for both “Primary antibody” and “Neutralized primary antibody,” the samples must be prepared in duplicate. The 1–10 μL Gilson PIPETMAN should preferably be used.
Add Solution D against the HRP-conjugated marker solution.
Note: If the volume of the sample solution is 10 μL, the same volume of Solution D is added against the HRP-conjugated molecular weight marker solution.
After Solution D is added to the HRP-conjugated marker solution, incubate for 5 min at room temperature.
Add PBS(-) to the cell lysates up to a final volume of 200 μL for the slot blot analysis.
Note: The 50–200 μL Gilson PIPETMAN should be used.
Selecting lanes for sample, GA-AGEs-BSA, and HRP-conjugated marker solution
The selection of lanes for the sample, GA-AGEs-BSA, and HRP-conjugated marker solution is presented in Figure 4.
Note: The addition of samples onto the PVDF membrane is duplicated (N = 2) for both “Primary antibody” and “Neutralized primary antibody” [14]. However, the method by which one sample is applied onto each PVDF membrane (N = 1) can be performed [10,12].
Figure 4. Setup for the application of standard GA-AGEs-BSA, HRP-conjugated marker, and sample solutions onto the PVDF membrane (setup explained in depth in Takata et al. [14]). White open squares indicate slot lanes. L1, L4: 0, 1, 3, 10, 30, 60, and 100 ng of GA-AGEs-BSA aliquots and HRP-conjugated marker solution. L2, L3, L5, and L6: C2C12 cell lysate samples treated with 0, 0.5, 1, 1.5, and 2 mM glyceraldehyde for 24 h. (This figure was reproduced from Takata [8]; copyright belongs to Takata T.)
PBS(-) wash of the PVDF membrane in slot blot apparatus
Add 100 μL of PBS(-) to the slot blot apparatus without water aspiration.
Note: The Nichipet 7000 was used.
Sample, GA-AGEs-BSA, and HRP-conjugated marker solution applied onto the PVDF membrane and removed with water aspiration
Apply 200 μL of sample, GA-AGEs-BSA, and HRP-conjugated marker solutions.
Note: The 50–200 μL Gilson PIPETMAN should preferably be used. This operation should be performed with the water aspirator (Figure 5).
Place the slot blot apparatus on a stand near the water aspirator.
Note: Any stand may be used for the experiment, as long as it has a height of 25–30 cm (Figure 6).
Connect a tube from the flow valve to the water aspirator (Figure 6).
Vacuum the sample, GA-AGEs-BSA, and HRP-conjugated marker solutions with water aspiration while opening one valve for air (Figure 7A).
For complete sample addition, we recommend vacuuming with water aspiration while the valve is closed for air (3–5 s) (Figure 7B).
Note: Although water aspiration pressure is not specified, this can be estimated. Vacuuming with water aspiration was performed at the Kanazawa Medical University, where the water supply is collected in a water tank and redistributed between laboratories. Here, the water pressure is consistent with that of a typical household (0.15–0.74 MPa) or corporate water supply system in Uchinada (0.20–0.23 MPa), according to data from the Ministry of Health, Labour, and Welfare in Japan. Therefore, the water supply system at Kanazawa Medical University has been adjusted to a pressure of 0.20–0.23 MPa.
Figure 5. Water aspirator (AS ONE) in the laboratory at Kanazawa Medical University
Figure 6. Slot blot apparatus on a stand; the large tube runs between the flow valve and water aspirator
Figure 7. Valve between slot blot apparatus and vacuum with water aspirator. A. The vacuuming with one valve opened against air. B. The vacuuming with the valve closed against air (This figure was reproduced from Takata et al. [9]. Copyright belongs to Takata T., Masauji T., and Motoo Y., who permitted the use of this image.)
PBS(-) wash of the PVDF membrane in slot blot apparatus following section L
Add 200 μL of PBS(-) to the slot blot apparatus.
Note: The Nichipet 7000 was used.
Vacuum PBS(-) with water aspiration while opening one valve for air (Figure 7A).
For complete sample addition, we recommend vacuuming with water aspiration while the valve is closed for air (3–5 s) (Figure 7B).
Milli-Q water wash of the PVDF membrane
Remove the PVDF membrane from the slot blot apparatus.
Incubate the PVDF membrane in Milli-Q water in a disposable polypropylene tray (AS ONE) for 1 min at room temperature.
PVDF membrane dissection and incubation in 5% SM-PBS-T
Cut the PVDF membrane into smaller sections (9 cm × 6 cm).
For blocking, incubate each PVDF membrane section in 5% SM-PBS-T (15 mL) for 30 min at room temperature (see Recipe 12).
Note: Each PVDF section was placed into a disposable polypropylene tray and processed using the seesaw shaker (BIO CRAFT).
PVDF membrane incubation in primary and neutralized primary antibody solutions
Make two sample packs using the Hybri-Bag (hard type).
Note: The pack should contain the PVDF membrane section (size: 9 cm × 6 cm) and ~5 mL of solution.
Wash each PVDF membrane with 0.5% SM-PBS-T (10 mL) for 5 min at room temperature. Perform this operation twice (see Recipe 13).
Note: Each PVDF membrane section is placed into a disposable polypropylene tray and incubated on the seesaw shaker (BIO CRAFT) in a refrigerator or cooled room.
Pack one PVDF membrane with primary antibody solution (see Recipe 14) and another with neutralized primary antibody solution (see Recipe 15).
Note: The primary and neutralized antibody solutions must be prepared and incubated on the seesaw shaker for 1 h at room temperature before this step.
Incubate both packs overnight at 4 °C.
Note: Incubation on the seesaw shaker (BIO CRAFT) could be performed in a refrigerator or a cooled room.
PVDF membrane incubation in secondary antibody solution
Remove each PVDF membrane from the primary or neutralized primary antibody solution.
Incubate each PVDF membrane in 0.5% SM-PBS-T (10 mL) for 10 min at room temperature, three times.
Note: Each PVDF membrane section is placed into a disposable polypropylene tray and incubated on the seesaw shaker (BIO CRAFT) in a refrigerator or cooled room.
Make two sample packs using the Hybri-Bag.
Note: The pack should contain the PVDF membrane section (9 cm × 6 cm) and ~5 mL of solution.
Pack each PVDF membrane with a secondary antibody solution (see Recipe 16).
Incubate each PVDF membrane for 1 h at room temperature.
Note: The incubation on the seesaw shaker (BIO CRAFT) should be performed.
PVDF membrane incubation in PBS-T and PBS(-)
Remove each PVDF membrane from the secondary antibody solution.
Incubate each PVDF membrane in PBS-T (15 mL) twice for 10 min at room temperature.
Note: Each PVDF is placed into a disposable polypropylene tray and processed using the seesaw shaker (BIO CRAFT).
Pause point: The PVDF membranes could be incubated in PBS-T on the seesaw shaker for 3–12 h at room temperature before the next step.
Incubate each PVDF membrane section in PBS(-) (15 mL) at room temperature.
Note: Incubation is generally performed for 10 min before section S.
Chemiluminescence imaging of bands on PVDF membrane (see General note 2)
Make two sample packs using the Hybri-Bag.
Take 500 μL of reagents A and B of an ImmunoStar LD kit on the commercial sheet (e.g., polyvinylidene chloride sheet). Perform this operation twice.
Mix Reagents A and B.
Smear each PVDF membrane with the reagent mixture for 10 s at room temperature.
Note: The volume of the mixture of reagents A and B in the ImmunoStar LD kit is 1,000 μL. The area of the PVDF membrane is approximately 27 cm2 (9 × 3 cm). The reagent solution density is 1,000 μL/27 cm2 = 37.0 μL/cm2.
After removing the reagent mixture, place each PVDF membrane section on the sheet for 30 s at room temperature.
Note: This procedure processes two PVDF membranes simultaneously.
Pack each PVDF membrane and analyze using the Fusion FX imager.
Obtain the images of the two PVDF membranes in the standard mode.
Note: We recommend an exposure time of 0.5–30 s. Two PVDF membranes should be exposed at the same time (e.g., 10, 15, 20, and 30 s). We recommend that images be obtained in standard, high, or ultra mode (if it is difficult to obtain images).
Data analysis
Calculation of protein concentration in cell lysates
A standard curve was drawn based on the absorbance data measured using the iMark microplate reader (595 nm). The average concentration of the standard BSA solution was calculated from duplicate measurements (Figure 8). Similarly, the average value for the samples was calculated from duplicate measurements. The sample protein concentration was averaged from two standard curves corresponding to the 5- and 10-min Bradford dye treatments. Note: If the absorbance of all samples is less than one of 1.5 μg/μL, the range from 0 to 1.5 μg/μL can be selected to draw the standard curve, whose value of correlation coefficient (R2) is high grade.
Figure 8. Standard curve of BSA in Solution D. The average concentration of the standard BSA solution was calculated from duplicate measurements. The examination for this example data was performed on June 29, 2022. ABS; absorbance. R2; correlation coefficient. A. The range was from 0 to 2.0 μg/μL. Left: Measurement of absorbance for Bradford dye treatment for 5 min. Right: Measurement of absorbance for Bradford dye treatment for 10 min. B. The range was from 0 to 1.5 μg/μL. Left: Measurement of absorbance for Bradford dye treatment for 5 min. Right: Measurement of absorbance for Bradford dye treatment for 10 min.
Detection of chemiluminescence bands on the PVDF membrane
To detect the chemiluminescence band image, images were exposed with the chemiluminescence mode of Fusion FX equipment, and the image was reflected in the general personal computer that contained Fusion FX software (version 17.03). The example chemiluminescence bands image of GA-AGEs-BSA and intracellular GA-AGEs in cell lysates on PVDF membranes are published in Supplementary Figure 1a in reference [14]. The model images are described in Figure 9.
Figure 9. Chemiluminescence detection of standard GA-AGEs-BSA aliquots, HRP-conjugated marker solution, and sample solution on PVDF membranes. Both membrane sides were simultaneously exposed using the Fusion FX imager. A. (L1–L3): Anti-GA-AGE antibody was probed onto the PVDF membrane. B. (L4–L6): Neutralized anti-GA-AGE antibody was probed onto the PVDF membrane. Closed gray and black squares indicate bands on the PVDF membrane. L1, L4: 0, 1, 3, 10, 30, 60, and 100 ng of GA-AGEs-BSA aliquots and HRP-conjugated marker solution. L2, L3, L5, L6: Cell lysate samples of C2C12 cells treated with 0, 0.5, 1, 1.5, and 2 mM glyceraldehyde for 24 h. (This figure is from Takata [8]; copyright belongs to Dr. Takata.)
Supplementary information
The HRP-conjugated marker solution was used as a control. The corrected luminance value (arbitrary unit, AU) was calculated as follows: Calibration-corrected luminance value = (HRP Left – Blank Left) / (HRP Right – Blank Right).
Note: We performed three independent experiments. Statistical analysis of one-way ANOVA and Tukey tests were used to quantify intracellular GA-AGE levels [14].
Validation of protocol
This protocol or parts of it have been used and validated in Takata et al. [14] (DOI: 10.1186/s13098-020-00561-z).
The chemiluminescence bands on PVDF membranes are shown in Supplementary Figure 1a of Takata et al. [14].
The detection of bands of standard GA-AGEs-BSA and intracellular GA-AGE using the novel slot blot approach was validated in various cells under the same or similar protocols [10–13,15–17].
General notes and troubleshooting
General notes
Measurement of protein concentration in cell lysates
We discourage the use of commercial standard BSA because it contains various compounds that may affect absorbance. Instead, we recommend that BSA powder be dissolved in Solution D.
Note: This method can be applied to tissue lysates [15,16].
Chemiluminescence imaging of bands on PVDF membranes
Although we used the Fusion FX imager and its software [14], other imaging equipment may be used [10,12,17].
For example, a LAS4000 system (GE Healthcare) was used when this novel slot blot approach was developed in 2017 [10].
Limitation
The volume and mass of samples should be less than 500 μL and 10 μg of protein, respectively.
Troubleshooting
Problem 1: Bands are invisible in chemiluminescence images (Case 1).
Possible cause: Proteins in cell lysates are unable to be probed onto the PVDF membrane because it was completely dried before the GA-AGEs-BSA, HRP-conjugated marker, and sample solution were added to it.
Solution: GA-AGEs-BSA, HRP-conjugated marker, and sample solutions should be rapidly added onto the PVDF membrane after the PBS(-) wash.
Problem 2: Bands are invisible in chemiluminescence images (Case 2).
Possible cause: Proteins in cell lysates are unable to be probed onto the PVDF membrane because Solution D contains Triton-X.
Solution: Confirm the composition of Solution D, which should not contain Triton-X.
Problem 3: Excessive background noise in PVDF membrane detection.
Possible cause: The volume of Solution D added onto the PVDF membrane may be excessive.
Solution: We recommend using 5–15 μL of Solution D, while cell lysates should be prepared with 150–200 μL of Solution D.
Acknowledgments
This protocol was modified from [10] in which Dr. Takata’s lysis buffer and PVDF membrane were first used in the slot blot analysis for quantification of intracellular GA-AGEs.
Author Contributions: Manuscript writing: T.T., H.M., and T.M. Figure preparation: T.T. Funding: This work was supported by grants from JSPS KAKENHI (grant number JP21K11607, to T.T. and grant number JP24K14802, to T.T.). Copyright: Yoshiharu Motoo, M.D., Ph.D. (Department of Internal Medicine, Fukui Saiseikai Hospital; Watanabecho, Fukui, Japan 918-8503) who has the copyright of the image in Figure 7 as well as T.T. and T.M. (Copyright is shared by three authors of Ref. 9) permitted use of it.
Competing interests
The authors declare no competing interests.
Ethical considerations
This protocol used the C2C12 cell line. No animal or clinical experiments were performed.
References
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Article Information
Publication history
Received: Apr 6, 2024
Accepted: Jun 23, 2024
Available online: Jul 5, 2024
Published: Jul 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Biochemistry > Carbohydrate > Glucose
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Original Research Article:
The authors used this protocol in Protein Science Sep 2023
Abstract
Peripheral membrane proteins (PMPs) are a subgroup of membrane-associated proteins that are water-soluble and bind to membranes, often reversibly, to perform their function. These proteins have been extensively studied in the aqueous state, but there is often a lack of high-resolution structural and functional studies of these proteins in the membrane-bound state. Currently, nuclear magnetic resonance (NMR) is among the best-equipped methods to study these relatively small proteins and domains, but current models have some disadvantages that prevent a full understanding of PMP interactions with membranes and lipids. Micelles, bicelles, and nanodiscs are all available for NMR observation but are based on synthetic lipids that may destabilize proteins or are too large to accommodate straightforward structural analysis. This protocol introduces a method for forming reverse micelles using lipids from natural sources, here called native reverse micelles. This technique allows the PMPs to embed within a shell of naturally derived lipids surrounding a small water core solubilized in an alkane solvent. PMP embedment in the lipid shell mimics binding to a cellular membrane. Here, naturally derived lipids from soy, bovine heart, and porcine brain are used in conjunction with n-dodecylphosphocholine (DPC) to encapsulate a PMP from either concentrated or dried protein, resulting in reverse micelles that may be confirmed via dynamic light scattering and NMR. This protocol allows for high-quality NMR data of PMPs interacting with membrane lipids within a biologically accurate environment.
Key features
• This protocol describes using natural lipids to construct reverse micelles for high-resolution NMR studies of proteins.
• Initial optimization of encapsulation conditions proceeds through visual assessment, with dynamic light scattering (DLS) to measure size distribution, and NMR to observe protein behavior.
• Membrane-interacting proteins are encapsulated in their membrane-bound state. Proteins that do not interact with membranes are housed in their water-solubilized state.
• Structural, functional, and inhibitory studies may be performed on native reverse micelle-encapsulated proteins.
Keywords: Membrane models Peripheral membrane proteins Protein NMR Protein–membrane interactions Reverse micelles
Graphical overview
Background
Peripheral membrane proteins (PMPs) are water-soluble proteins that can bind, often reversibly, with membranes. Interactions between PMPs and their target membranes are of great interest due to the central role of these proteins in disease and biology. Currently, methods to accurately observe the protein–membrane interaction, especially at high resolution, are limited. Nuclear magnetic resonance (NMR) is well-suited to observe these interactions using micelles, bicelles, and nanodiscs. However, these membrane models are most commonly formulated from artificial components, limiting understanding in a biological context. Reverse micelles (RMs) have a long history in the study of proteins, including being used to house membrane-associated and membrane-integral proteins for NMR [1,2]. RMs house proteins within a nanoscale pool of water, surrounded by a hollow shell of surfactants with the hydrophilic headgroups interacting with the water and the hydrophobic tails pointing outward and serving to solubilize the RM in an alkane solvent. High-quality NMR spectra of proteins encapsulated within RMs are routinely attainable, enabled by the low-viscosity alkane solvent [3]. Recent developments of RMs have improved their properties as membrane mimetics through formulations using phosphocholine-based surfactants. Utilizing membrane-mimicking RMs (mmRMs), the PMP of interest can be observed interacting with the surfactant shell. This allows the mapping of PMP-membrane interaction surfaces and enables structural and functional studies using NMR. RMs formulated with naturally derived lipids represent an advancement in technology, allowing the study of PMPs and other proteins in a more native-like membrane mimetic [4]. The lipid extractions used for native reverse micelles (nRMs), including soy lecithin, bovine heart lipids, and porcine brain lipids, account for a wide range of lipid compositions. These compositions introduce a heterogeneous mixture of lipid headgroup and tail types, reflecting the complexity of biological membranes. Conversely, a formulation of RMs made entirely from phosphocholine-based surfactants can be used as a standard background to explore specific lipid interactions or for structural and functional studies that require a more homogeneous membrane model [5]. Previous applications of RMs to protein studies may also be extended to nRMs, including structural determination of PMPs [6], hydration dynamics measurements [7], experimental cosolvent mapping [8], and enhanced fragment screening [9]. Integral membrane proteins and proteins anchored through lipidations have previously been housed in RMs and would benefit from study in the more native-like environment described here [1,2]. The following protocol describes the construction of reverse micelles from lipids extracted from native sources or phosphocholine-based surfactants, for high-resolution NMR study of encapsulated PMPs in their membrane-bound state.
Materials and reagents
Biological materials
Protein of interest; in this protocol, glutathione peroxidase 4 (GPx4) is used as an example for encapsulation in reverse micelles. Protein was produced as previously described using recombinant 15N or 13C-15N isotopic labeling, expression, and purification from BL21(DE3) E. coli [10].
Reagents
Lecithin (VWR, catalog number: 10791-822)
Ethylenediaminetetraacetic acid (EDTA) (Fisher, catalog number: AAJ15694AE)
Sodium chloride (NaCl) (Fisher, catalog number: S271-1)
Bis-Tris (VWR, catalog number: 6976-37-0)
Tris base (Sigma-Aldrich, catalog number: 77-86-1)
Chloroform (Fisher, catalog number: 67-66-3)
Methanol (EM Science, catalog number: 67-56-1)
Distilled water
Hydrochloric acid (HCl) (Fisher, catalog number: SA49)
Dithiothreitol (DTT) (GoldBio, catalog number: DTT10)
Sodium sulfate (VWR, catalog number: 7757-82-6)
Brain total lipid extract (porcine) (PBL) (Avanti Polar Lipids, catalog number: 131101C)
Heart total extract (bovine) (BHL) (Avanti Polar Lipids, catalog number: 171201P)
Pentane (Alfa Aesar, catalog number: AA32449K2)
1-Hexanol (Sigma-Aldrich, catalog number: 111-27-3)
1,2-dilauroyl-sn-glycero-3-phosphocholine (DLPC) (Avanti Polar Lipids, catalog number: 850335C)
n-dodecylphosphocholine (DPC) (Avanti Polar Lipids, catalog number: 850336P)
D-pentane (Sigma-Aldrich, catalog number: 490482)
D2O (Sigma-Aldrich, catalog number: 7789-20-0)
Trimethyl phosphate (Sigma-Aldrich, catalog number: 512-56-1)
Bradford dye (Thermo Fisher, catalog number: 23238)
Solutions
5 M NaCl stock solution (see Recipes)
500 mM Bis-Tris pH 6.0 stock solution (see Recipes)
1 M DTT stock solution (see Recipes)
200 mM EDTA pH 6.0 stock solution (see Recipes)
6 M HCl stock solution (see Recipes)
Protein buffer (see Recipes)
Dilute protein buffer (see Recipes)
Mobile phase for soy lecithin chelation (see Recipes)
Recipes
5 M NaCl stock solution
*Note: Initially, add half of the distilled H2O to dissolve the NaCl; the rest of the volume should slowly be added as NaCl dissolves. The stock should be brought up to the final volume once all NaCl is dissolved to account for any volume displacement caused by the solid powder.
Reagent Final concentration Quantity or Volume
NaCl 5 M 146.1 g
H2O n/a *see Note
Total n/a 500 mL
500 mM Bis-Tris pH 6.0 stock solution
*Note: Initially, add half of the distilled H2O to dissolve the Bis-Tris; most of the rest should slowly be added as Bis-Tris dissolves. Once the Bis-Tris is dissolved, add small increments of 6 M HCl into the solution to bring the pH to 6.0. The stock should be brought up to the final volume with distilled H2O once the Bis-Tris is pH-corrected to account for any volume displacement caused by the solid powder and HCl.
Reagent Final concentration Quantity or Volume
Bis-Tris pH 6.0 500 mM 52.3 g
H2O n/a *see Note
pH corrected by HCl *see Note
Total n/a 500 mL
1 M DTT stock solution
*Note: Initially, add half of the distilled H2O to dissolve the DTT; the rest should slowly be added as DTT dissolves. The stock should be brought up to the final volume once all DTT is dissolved to account for any volume displacement caused by the solid powder.
Reagent Final concentration Quantity or Volume
DTT 1 M 1.54 g
H2O n/a *see Note
pH adjusted using HCl
Total n/a 10 mL
200 mM EDTA pH 6.0 stock solution
*Note: Initially, add half of the distilled H2O to dissolve the EDTA; most of the rest should slowly be added as EDTA dissolves. Once the EDTA is dissolved, add small increments of 6 M HCl to the solution to bring the pH to 6.0. The stock should be brought up to the final volume with distilled H2O once the EDTA is pH-corrected to account for any volume displacement caused by the solid powder and HCl.
Reagent Final concentration Quantity or Volume
EDTA 200 mM 2.92 g
H2O n/a *see Note
pH adjusted using HCl
Total n/a 50 mL
6 M HCl stock solution
Reagent Final concentration Quantity or Volume
HCl, 12 N 6 M 100 mL
H2O n/a 100 mL
Total 6 M 200 mL
Protein buffer
Reagent Final concentration Quantity or Volume
Bis-Tris pH 6.0, 500 mM stock 20 mM 2 mL
NaCl, 5 M stock 100 mM 1 mL
DTT, 1 M stock 20 mM 1 mL
H2O n/a 46 mL
Total n/a 50 mL
Dilute protein buffer
Reagent Final concentration Quantity or Volume
Bis-Tris pH 6.0, 500 mM stock 20 μM 2.4 μL
NaCl, 5 M stock 100 μM 1.2 μL
DTT, 1 M stock 20 μM 1.2 μL
H2O n/a 59.952 mL
Total n/a 60 mL
Mobile phase for soy lecithin chelation
Note: Dry the solvents needed with sodium sulfate overnight and decant before the mobile phase is made.
Reagent Final concentration Quantity or Volume
Chloroform n/a 10 mL
Methanol n/a 5 mL
Total n/a 15 mL
Laboratory supplies
1.5 mL glass vials with screw top lid (Sigma-Aldrich, catalog number: 854171)
Pierce Concentrator, PES membrane, 10K MWCO 0.5 mL (Thermo Fisher, catalog number: 88513)
Spin-XR UF 20 10K MWCO PES membrane 20 mL (Corning, catalog number: 431488)
Amicon Ultra-15 centrifugal filters regenerated cellulose membrane 3K 20 mL (Millipore Sigma, catalog number: UFC901024)
NMR tubes (New Era Enterprises, Inc., catalog number: NE-UL5-7)
1.5 mL microcentrifuge tubes (Cell treat, catalog number: 229441)
20 mL scintillation vials (Wheaton, catalog number: 03-341-25K)
Pipettes (Corning, catalog numbers: 4075, 4071, 4072, 4074)
Pipette tips (Fisherbrand/Olympus, catalog numbers: 02-717-134/22-119B, 24-150RL, 23-404)
Glass pipettes (Fisherbrand, catalog number: 13-678-6B)
Separation funnel (Chem Glass, catalog number: CG1743-09)
25 mL glass graduated cylinder (Chem Glass, catalog number: CG-8242-25)
Stir bar (Fisher Scientific, catalog number: 16-800-507)
Pipette bulbs (Fisherbrand, catalog number: 03-448-21)
Teflon tape (SPBel-Art, catalog number: 240200000)
Parafilm (Bemis Parafilm, catalog number: PM996)
Equipment
Pioneer analytical balance (Ohaus, model number: PX84/E)
Navigator analytical scale (Ohaus, model number: NV222)
Eppendorf centrifuge 5425 (Eppendorf, catalog number: 5405000646)
Eppendorf thermomixer (Eppendorf)
Note: The model used in this protocol was discontinued; a comparable model would be the Eppendorf Thermomixer, catalog number: 5384000020.
LSE digital dry bath (Corning, catalog number: 6875-SB)
Hereaus megafuge 8 centrifuge (Thermo Scientific, catalog number: 75007210)
ST plus series centrifuge (Sorvall, catalog number: 75009909)
Speed vacuum concentrator (Savant, catalog number: SVC100H)
nXDS vacuum pump (Edwards, catalog number: A73501983)
4 °C refrigerator (VWR, catalog number: 10819-904)
-80 °C freezer (Fisherbrand, catalog number: IUE30086FA)
Zetasizer nano (Malvern Panalytical, model: ZEN3600)
2800 ultrasonic bath (Branson, catalog number: M2800)
LSE vortex mixer (Corning, catalog number: 6778)
Mini centrifuge (Corning, catalog number: 6770)
AG centrifuge (Eppendorf, catalog number: 022620100)
Laboratory fume hood (Hamilton Laboratory Solutions, model: 61L)
Advance stir plate (VWR, catalog number: 76557-502)
Genesys 150 UV spectrophotometer (Thermo Scientific, catalog number: 840-300000)
pH meter (Oakton, catalog number: EW-35413-20)
High-field (≥ 500 MHz) NMR spectrometer, equipped with triple-resonance inverse probe
Software and datasets
Prism v10.2 (GraphPad, 02/5/2024)
NMRPipe (v11.4) [11]
NMRFAM-Sparky (v3.12, 04/15/2015)
Bruker Topspin (v4.1.3)
Procedure
Reverse micelle calculations
Determine the desired surfactant concentration.
Note: This concentration may be tested for the protein of interest and the specific experiments to be performed. For the protein used here, GPx4 (18.6 kDa), a 75 mM surfactant concentration is used, while another common concentration is 150 mM. The concentration of 75 mM provides slightly better spectroscopic properties in the form of slightly narrower NMR line widths, while 150 mM allows for a higher protein concentration due to the increased aqueous phase volume.
Calculate the mass of surfactants needed for the desired ratio based on molecular weight. For a 50:50 molar percent ratio of natural lipid (or DLPC) to DPC:
DPC (MW: 351.462): 6.6 mg
DLPC (MW: 621.826): 11.6 mg
or
Soy lecithin: 12.1 mg
or
Porcine brain lipids: 12.1 mg
or
Bovine heart lipids: 12.1 mg
Note: All compositions can form RMs at a 50:50 molar percent ratio of natural lipid (or DLPC) to DPC, but DLPC:DPC and lecithin:DPC work well up to 70:30 molar percent ratios. Soy lecithin, porcine brain, and bovine heart lipids are complex lipid mixtures, and the molar percent ratios are estimated as outlined in Walters et al. [4].
Calculate the volume of water phase needed in relation to the total volume of the sample (500 μL) and the target water loading (W0). W0 is the molar ratio of water to surfactant.
W
0
=
H
2
O
s
u
r
f
a
c
t
a
n
t
s
W0 = 20: 13.5 μL
Note: W0 values differ based on the size of encapsulated protein, desired water dynamics, and target RM size. GPx4 encapsulates at W0 equal to and greater than 20, while ubiquitin (8.6 kDa) can be encapsulated at a W0 as low as 10. The range of 10–25 is typical in these systems.
Protein preparation: concentration
There are two ways to prepare the protein for encapsulation: concentration or vacuum concentration/lyophilization. Each has its advantages and disadvantages. Every protein should be prepared both ways initially to determine the best method. Concentration is a faster process, and the protein can be encapsulated the day of, but it is more likely to aggregate during concentration.
Use an appropriate 1.5 mL spin concentrator.
In this example, for GPx4, a 10K MWCO PES membrane is appropriate.
Verify if the initial protein concentration is ~1.2 mg/mL or higher through Bradford assay. Immediately before encapsulation, concentrate protein to the desired volume based on the W0.
Note: Some proteins may survive concentrating at room temperature and others may need to be at 4 °C. This will be protein-dependent and will need to be evaluated for each protein.
Protein preparation: vacuum concentration
If concentration does not result in optimal encapsulation, vacuum concentration (or lyophilization) results in higher encapsulation efficacy for many proteins but has to be prepared the day before sample preparation.
The day before RM sample preparation, use a PES spin concentrator (10K MWCO, 20 mL) and buffer-exchange the protein sample from the original buffer to dilute buffer.
Prepare 60 mL of dilute protein buffer.
Add protein samples at 1.2 mg/mL (initial volume is 1 mL) and concentrate to 500 μL by centrifuging at 3,260× g until 500 μL is reached.
Note: Some protein may be lost throughout the buffer exchange process. To account for this loss, up to 2-fold more protein is needed before concentration. Protein concentration may be verified through a Bradford assay after exchange.
Add dilute protein buffer to protein sample up to 20 mL and centrifuge until the volume is at 500 μL.
Repeat two more times.
Add a final 500 μL of protein volume to a 1.5 mL glass vial.
Freeze protein solution for at least 10 min in a -80 °C freezer.
Add vial to speed vacuum concentrator and allow to sublimate overnight, leaving only protein and dilute buffer components.
Soy lecithin metal contamination removal
This step is important when using soy lecithin to encapsulate protein in RMs. Without this step, the excess metal in the lecithin will induce line broadening in the 31P spectrum and may impact protein spectra (Supplementary Figure S2 [4]). Perform this step in a fume hood.
Dry chloroform and methanol overnight with sodium sulfate. Decant the sodium sulfate via gravity filtration from the solvents the following day.
Make 15 mL of mobile phase.
Weigh 1 g of soy lecithin into a 20 mL scintillation vial.
Add the mobile phase to the lecithin and allow to stir on a stir plate until dissolved.
Add 5 mL of 200 mM EDTA pH 6.0.
Allow to chelate by stirring for 24 h with a stir bar on a stir plate with the lid and parafilm.
Add contents to a separatory funnel.
Shake the funnel and release pressure as needed by slightly opening the spigot of the separatory funnel. Shake until fully mixed.
Allow contents to separate; then, remove the bottom organic layer into a graduated cylinder.
Add additional mobile phase up to 15 mL and transfer to a clean 20 mL scintillation vial.
Add 5 mL of 200 mM EDTA pH 6.0.
Allow to stir overnight.
Repeat steps D6–8.
Transfer the organic layer into a pre-weighed 20 mL scintillation vial and allow it to sit open to evaporate off the solvents overnight on a 37 °C heat block.
Note: Dry lecithin overnight in a speed vacuum concentrator before use to remove excess moisture. The lecithin will be slightly oily and tacky.
To verify heavy metal extraction, add 75 mM lecithin (24.1 mg) to 500 μL of pentane and 13.5 μL of distilled water (W0 = 20). Titrate 1-hexanol in 50 mM increments (3.125 μL of 8 M 1-hexanol) until visual clarity is reached.
Perform 31P 1D NMR scan to verify narrow lines and seven individual peaks corresponding to the different headgroup types using trimethyl phosphate as the 31P standard.
DLPC:DPC RM
The night before experimentation, place all surfactants in the speed vacuum concentrator to remove excess moisture.
Weigh out surfactants based on the decided ratio into a 1.5 mL glass vial. For a 50:50 molar percent ratio DLPC:DPC RM, measure out 11.6 mg of DLPC and 6.6 mg of DPC.
Add 500 μL of pentane to surfactants and vortex.
Add 200 mM 1-hexanol (12.5 μL of 8 M 1-hexanol) to the solution and vortex until most solids are in solution.
Add the desired water volume, including protein if needed.
If concentrating the protein, add directly from the spin concentrator to the solution.
If adding from vacuum-concentrated or lyophilized protein, add protein buffer to the protein and vortex until the protein is mostly in solution. Then, add the surfactants and solvent mixture to the vial containing the protein solution and vortex.
Titrate and vortex in between each addition with additional 1-hexanol in 200 mM increments (12.5 μL of 8 M 1-hexanol) until 800 mM total 1-hexanol is reached; then, titrate with additional 50 mM 1-hexanol increments (3.125 μL of 8 M 1-hexanol). Titrate until there is a visually clear solution and then add an additional 50 mM 1-hexanol.
Notes:
If adding from vacuum concentrated or lyophilized protein, the solution will need to be transferred twice (once into the original surfactant vial and once back into the protein vial) after the first and second additions of additional 1-hexanol from this step. This ensures that all surfactants and proteins are solubilized into the reverse micelle sample.
The typical needed range for 1-hexanol is 800 mM to 1.2 M, varying between different proteins and different sample conditions.
Pause point: Once visual clarity is reached, the RM will remain stable for an extended period of time (up to several months for most stable proteins). When preparing samples a day ahead, it is recommended that, after formation, they be left on a thermomixer at room temperature until needed. Samples may also be stored in refrigerated conditions to prolong viability.
Lecthin:DPC RM
The night before experimentation, place all surfactants in the speed vacuum concentrator to remove excess moisture.
Weigh out surfactants based on the decided ratio into a 1.5 mL glass vial.
For a 50:50 lecithin:DPC RM, measure out 12.1 mg of lecithin and 6.6 mg of DPC.
For a 70:30 lecithin:DPC RM, measure out 16.9 mg of lecithin and 3.95 mg of DPC.
Add 500 μL of pentane to surfactants and vortex.
Add 50 mM 1-hexanol (3.125 μL of 8 M 1-hexanol) to the solution and vortex until most solids are in solution.
Add the desired water volume, including protein if needed.
If concentrating the protein, add directly from the spin concentrator to the solution.
If adding from vacuum-concentrated or lyophilized protein, add protein buffer to the protein and vortex until the protein is solubilized. Then, add the surfactants and solvent mixture to the vial with protein and vortex.
Titrate with additional 1-hexanol in 50 mM increments (3.125 μL of 8 M 1-hexanol), vortexing in between each addition until there is a visually clear solution. Add an additional 50 mM 1-hexanol.
Notes:
If adding from vacuum-concentrated or lyophilized protein, the solution will need to be transferred twice (once into the original surfactant vial and once back into the protein vial) after the first and second additions of additional 1-hexanol from this step. This ensures that all surfactants and proteins are solubilized into the reverse micelle sample.
The typical range needed for 1-hexanol is 400–700 mM, which varies according to the protein and sample conditions used.
Figure 1 demonstrates the formation of lecithin:DPC nRMs. There is a need for proper addition of hexanol as well as a proper W0. The W0 must be carefully determined; if too little volume is added, the nRM will not form, and if too much volume is added, the solution will phase-separate (Figure 1A). If an insufficient amount of hexanol is titrated into the nRMs, visual clarity will not be reached and the nRMs will not form (Figure 1B). With the proper amount of hexanol, surfactants, and W0, a visually transparent solution will be acquired with fully formed nRMs (Figure 1C).
Figure 1. Two examples of empty native reverse micelle (nRM) formulations that do not solubilize and a fully solubilized nRM sample. A. 75 mM lecithin:DPC nRM with a W0 = 100 and 1 M hexanol displays a water content that is too large for nRM formation. Phase separation occurs and is observable by the separated aqueous layer at the bottom of the vial. B. 75 mM lecithin:DPC nRM with a W0 = 20 and 600 mM hexanol shows a hexanol concentration that is too low. nRM is not visually transparent; encapsulation and formation of nRMs is not complete, often remedied by the addition of more hexanol. C. 75 mM lecithin:DPC nRM with a W0 = 20 and 1 M hexanol. nRM is visually transparent; this indicates complete formation of nRMs.
PBL:DPC and BHL:DPC RMs
The PBL:DPC and BHL:DPC RMs can be formed with the same protocol. We describe the protocol for a PBL:DPC RM in this section, but the PBL can be substituted for the BHL with no additional alterations to the protocol.
The night before experimentation, place all surfactants in the speed vacuum concentrator to remove excess moisture.
Weigh out DPC based on the decided ratio into a 1.5 mL glass vial.
For a 50:50 molar percent ratio PBL:DPC RM, measure out 6.6 mg of DPC.
Make 100 mg/mL stock of PBL in pentane and calculate the needed PBL for 50:50 molar percent ratio.
For this ratio, 22.5 mg/mL is needed, which is equal to 112 μL of PBL solution.
Add 388 μL of pentane to DPC and vortex.
Add 112 μL of the PBL/pentane solution to the vial and vortex.
Add 50 mM 1-hexanol (3.125 μL of 8 M 1-hexanol) to the solution and vortex until most solids are in solution.
Add the desired water content and protein as needed.
If concentrating the protein, add directly from the spin concentrator to the solution.
If adding from vacuum-concentrated or lyophilized protein, add water to the protein and vortex until the protein is mostly in solution. Then, add the surfactants and solvent mixture to the vial with protein and vortex.
Titrate with additional 1-hexanol in 50 mM increments (3.125 μL of 8 M 1-hexanol), vortexing in between each addition until there is a visually clear solution. Add an additional 50 mM of 1-hexanol.
Notes:
If adding from vacuum-concentrated or lyophilized protein, the solution will need to be transferred twice (once into the original surfactant vial and once back into the protein vial) after the first and second additions of additional 1-hexanol from this step. This ensures that all surfactants and proteins are solubilized.
Optionally, PBL solution may be added to the glass vial the night before and the solvent dried off with nitrogen and then in a speed vacuum concentrator overnight; then, DPC should be added directly into the glass vial with PBL. This will accommodate 500 μL of pentane being added after the lipids and step G4 may be omitted.
NMR sample preparation
Add 10% deuterated solvent as a lock solvent to reverse micelles. For RM with pentane as the solvent, add 50 μL of d-pentane.
If needed, deuterated alkane and surfactant may be used to suppress background signal and suppress artifacts for protein NMR experiments. Deuteration generally does not affect the stability of RM systems. However, at this time, deuterated lipid extracts are not available.
Add the sample into an NMR tube and cover with Teflon tape to reduce evaporation of solvents.
Note: Screw-top NMR tubes, such as Norell, catalog number: C-S-5-600-SC-7, may be used to mitigate evaporation in longer 3D NMR experiments. However, in our experience, a well-sealed standard NMR tube can be used in most cases.
NMR experiments
Add 10% deuterated solvent as a lock solvent to the reverse micelle sample.
For RMs with pentane as the solvent, add 50 μL of d-pentane or d-hexane.
The pentane (or hexane) methyl signal is used for the NMR spectrometer lock.
Standard Bruker pulse sequences are suitable for the approach reported here.
Echo-anti echo gradient–selected NMR experiments are typically used for suppression of solvent and lipid/surfactant signals.
Water suppression using flip-back pulses is generally not necessary and may be turned off due to the low overall concentration of water in the RM system.
Pulse length and water frequency are calibrated as typical for aqueous NMR samples.
15N-HSQC experiments of encapsulated proteins are typically collected to confirm encapsulation and map membrane interactions of PMPs. The first increment of 15N-HSQC experiments may be used to track sample stability over time (Figure 2). 2D-15N-HSQC experiments provide amino acid resolution, which may be compared to a reference through calculated chemical shift perturbations (CSPs).
Figure 2. Overlay of the first increment of 15N-HSQC experiments of GPx4 in a 50:50 molar percent ratio lecithin:DPC native reverse micelle (nRM) with 100 μM of GPx4 in nuclear magnetic resonance (NMR) buffer with 1 M hexanol, collected multiple days apart to check sample stability. The red spectrum was collected on day one, and the blue spectrum was collected on day 4, with the sample stored at room temperature throughout. The overlay indicates that very little protein signal was lost between experiments, confirming protein stability within the nRM.
DLS experiments
Collect DLS measurements in a quartz cuvette at room temperature. A typical sample volume is 500 μL. Hexane may be used for DLS, since signal loss from slow tumbling is not a concern and hexane evaporates more slowly than pentane.
Use published viscosity and dielectric constant parameters using the volume ratio of hexane to hexanol [12,13].
A typical ratio used for the determination of the viscosity and dielectric constant for a nRM with 1 M hexanol in hexane is 0.875. The viscosity value using these parameters is 0.356 and the dielectric constant is 2.10. The refractive index value is kept constant at 1.38.
Data were collected on a Malvern Zetasizer nano using standard DLS measurement settings.
DLS measurements are typically collected in triplicate to calculate error. Figure 3 demonstrates the DLS measurements of GPx4 encapsulated in a lecithin:DPC nRM.
Figure 3. Dynamic light scattering (DLS) size distribution of 75 mM lecithin:DPC nRM with a W0 = 20 encapsulating 100 μM GPx4 in 20 mM Bis-Tris pH 6.0, 100 mM NaCl, and 10 mM DTT, with 1 M hexanol as the cosurfactant. Average diameter is 6.8 nm with a size distribution of 1.4 nm.
Data analysis
The analysis of the protein NMR spectra of encapsulated PMPs has been previously described in Labrecque et al. [5]. Briefly, all NMR data were processed using the NMRPipe software and visualized and analyzed using the NMRFam-Sparky distribution. The assignments for aqueous GPx4 have been previously published (BMRB: 50955) and the encapsulated protein assignments were transferred from the aqueous assignments as described in Labrecque et al. [5]. The chemical shift perturbations were calculated using the following formula using weighted shifts:
∆1H and ∆15N represent the changes in the 1H and the 15N chemical shifts for each resonance. Resonance CSPs can be mapped to the corresponding residues on the structure of the protein for a view of membrane interaction surfaces. Figure 4 demonstrates the shifting that occurs when GPx4 is encapsulated in a nRM compared to its aqueous state.
Figure 4. 1H-15N HSQC of aqueous GPx4 (black) and encapsulated GPx4 (red). The encapsulation conditions are 75 mM 50:50 molar percent ratio lecithin:DPC with 1 M hexanol (GPx4 buffer: 20 mM Bis-Tris pH 6.0, 100 mM NaCl, and 10 mM DTT). Both HSQCs were collected at 25 °C and 600 MHz on a room temperature probe and processed on NMRPipe and the NMRFAM-Sparky distribution. Contours were lowered to near the noise level in both spectra.
Validation of protocol
The protocol described here has been used and validated in:
Walters et al. [4]. Investigating protein-membrane interactions using native reverse micelles constructed from naturally sourced lipids. Protein Science (Figure 2, panels a–f) (Figure 4, panels a, d) (Figure 5, panels c, f, i) (Supplementary Figure 2, panels a–e).
Labrecque et al. [5]. Membrane-Mimicking Reverse Micelles for High-Resolution Interfacial Study of Proteins and Membranes. Langmuir (Figure 4, panels a–b) (Supplementary Figure 2, panel a).
Successful encapsulation of GPx4, along with other proteins, is shown in Walters et al. [4] and Labrecque et al. [5] for all RM systems described here. The confirmation of encapsulation was determined via protein NMR and the validity of the compositions was confirmed through comparable chemical shift perturbation (CSP) in the active and binding sites of the proteins. Ubiquitin was used as a control protein, and NMR revealed that the formulations of the reverse micelles did not impact the aqueous structure of ubiquitin. DLS confirms the formation of uniformly sized RMs, which increase in diameter as the water loading is increased, as expected.
General notes and troubleshooting
General notes
Pentane is a highly volatile solvent, which makes it difficult to maintain the proper volume. Pipette carefully with either a glass pipette or a micropipette by slowly bringing the solvent into the pipette, and then maintaining exact pressure between vials before ejecting it into the new vial.
Alternatively, hexane may be used as the solvent. This mitigates evaporation but results in more NMR line-broadening for larger protein systems.
Protein stability can be a concern during the formation of the reverse micelles due to the high concentration in the water phase before addition to the surfactant/solvent phase. If protein aggregates, remove aggregate before addition to reverse micelle. If protein concentration is lost during buffer exchange, add additional protein before exchanging to mitigate loss.
If possible, NMR spectra of the protein of interest should be collected in membrane models with little or no curvature, such as isotropic bicelles or nanodiscs, and compared to a spectrum of the protein within nRMs. Major differences in spectra may indicate curvature-dependent conformational differences, and caution is warranted. Smaller spectral differences are likely indicative of differences in lipid content among the membrane models and suggest that the nRM is an adequate model for the protein of interest.
Alkanes and hexanol may bind to or otherwise perturb the proteins of interest. The above-described control using either isotropic bicelles or nanodiscs may also reveal changes in the protein structure from the solvent components. Due to its high water solubility, hexanol may be added to aqueous samples to determine if it is a strong binder to the protein of interest or if it otherwise perturbs the protein structure.
Troubleshooting
Problem 1: Reverse micelle is not reaching visual clarity.
Possible causes: Excess of lipids or protein or non-optimized water loading volume.
Solution: Verify surfactant and protein concentrations. Allow to shake for 1 h or sonicate in a water bath for 15 min. Slowly add additional 1-hexanol titrations. Adjust the water loading value (increase). In some cases, the reverse micelle will not fully clear and the additional hexanol will not improve the visible state of the reverse micelle. If this is the case, test the sample in the NMR for protein signal. Some proteins and sample conditions will produce high-quality NMR spectra without complete visual clarity.
Problem 2: Protein does not encapsulate efficiently, aggregation during concentration.
Possible cause: Sub-optimal stability in the high concentration necessary for delivering concentrated protein.
Solution: If the protein is amenable to vacuum concentration or lyophilization: vacuum concentrate or lyophilize the protein in a glass vial and construct the reverse micelle without the protein. Allow the reverse micelle to form without protein; then, after visual clarity is reached, add reverse micelle sample to the dried protein and vortex. Allow to shake overnight if the addition of protein does not immediately lead to a clear solution. If reverse micelle does not clear by the next day, titrate an additional 1-hexanol in 50 mM increments until visual clarity is reached or high-quality protein NMR spectra are observed.
Problem 3: Reverse micelle sample volume decreased over time.
Possible cause: Pentane is volatile and will rapidly evaporate.
Solution: Leave vials with pentane sealed when not actively adding to the vial. Once the reverse micelle is formed, add additional pentane up to 500 μL and seal the cap with Teflon tape.
Acknowledgments
This research was supported by the National Institute of General Medicine Sciences of the National Institutes of Health under Award Number R35GM147221. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. We would like to thank Courtney Labrecque, Abdul Castillo, Angela Develin, Aubree Nolan, Yun Qu, PhD, and Adam Offenbacher, PhD for their contributions to the original works used in this protocol. We would also like to thank Jake Breeden for the verification of written protocol. The DLPC:DPC reverse micelles were first reported in Labrecque et al. [5] and the native reverse micelles were first reported in Walters et al. [4].
Competing interests
The authors declare that they have no conflicts of interest.
References
Valentine, K. G., Peterson, R. W., Saad, J. S., Summers, M. F., Xu, X., Ames, J. B. and Wand, A. J. (2010). Reverse Micelle Encapsulation of Membrane-Anchored Proteins for Solution NMR Studies. Structure. 18(1): 9–16.
Kielec, J. M., Valentine, K. G., Babu, C. R. and Wand, A. J. (2009). Reverse Micelles in Integral Membrane Protein Structural Biology by Solution NMR Spectroscopy. Structure. 17(3): 345–351.
Wand, A. J., Ehrhardt, M. R. and Flynn, P. F. (1998). High-resolution NMR of encapsulated proteins dissolved in low-viscosity fluids. Proc Natl Acad Sci USA. 95(26): 15299–15302.
Walters, S. H., Castillo, A. J., Develin, A. M., Labrecque, C. L., Qu, Y. and Fuglestad, B. (2023). Investigating protein‐membrane interactions using native reverse micelles constructed from naturally sourced lipids. Protein Sci. 32(11): e4786.
Labrecque, C. L., Nolan, A. L., Develin, A. M., Castillo, A. J., Offenbacher, A. R. and Fuglestad, B. (2022). Membrane-Mimicking Reverse Micelles for High-Resolution Interfacial Study of Proteins and Membranes. Langmuir. 38(12): 3676–3686.
O'Brien, E. S., Nucci, N. V., Fuglestad, B., Tommos, C. and Wand, A. J. (2015). Defining the Apoptotic Trigger. J Biol Chem. 290(52): 30879–30887.
Nucci, N. V., Pometun, M. S. and Wand, A. J. (2011). Site-resolved measurement of water-protein interactions by solution NMR. Nat Struct Mol Biol. 18(2): 245–249.
Fuglestad, B., Kerstetter, N. E. and Wand, A. J. (2019). Site-Resolved and Quantitative Characterization of Very Weak Protein–Ligand Interactions. ACS Chem Biol. 14(7): 1398–1402.
Fuglestad, B., Kerstetter, N. E., Bédard, S. and Wand, A. J. (2019). Extending the Detection Limit in Fragment Screening of Proteins Using Reverse Micelle Encapsulation. ACS Chem Biol. 14: 2224–2232.
Labrecque, C. L. and Fuglestad, B. (2021). Electrostatic Drivers of GPx4 Interactions with Membrane, Lipids, and DNA. Biochemistry. 60(37): 2761–2772.
Delaglio, F., Grzesiek, S., Vuister, G., Zhu, G., Pfeifer, J. and Bax, A. (1995). NMRPipe: A multidimensional spectral processing system based on UNIX pipes. J Biomol NMR. 6(3): 277–293.
Franjo, C., Jimenez, E., Iglesias, T. P., Legido, J. L. and Paz Andrade, M. I. (1995). Viscosities and Densities of Hexane + Butan-1-ol, + Hexan-1-ol, and + Octan-1-ol at 298.15 K. J Chem Eng Data. 40(1): 68–70.
Singh, R. P. and Sinha, C. P. (1982). Dielectric behavior of the binary mixtures of n-hexane with toluene, chlorobenzene and 1-hexanol. J Chem Eng Data. 27(3): 283–287.
Article Information
Publication history
Received: Apr 16, 2024
Accepted: Jun 27, 2024
Available online: Jul 11, 2024
Published: Jul 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Biochemistry > Protein > Structure
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5,040 | https://bio-protocol.org/en/bpdetail?id=5040&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
An Experimental Protocol for the Boyden Chamber Invasion Assay With Absorbance Readout
KB Kathleen C. Brown
AS Amanda M. Sugrue
KM Kushal J. Modi
RL Reagan S. Light
KC Kaitlyn B. Conley
AC Ashley J. Cox
CB Christopher R. Bender
SM Sarah L. Miles
MV Monica A. Valentovic
Piyali Dasgupta
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5040 Views: 584
Reviewed by: Valérian DORMOYChhuttan L MeenaSalah Boudjadi
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Cited by
Original Research Article:
The authors used this protocol in Cell Adhesion & Migration Jan 2017
Abstract
The phenomenon of cell invasion is an essential step in angiogenesis, embryonic development, immune responses, and cancer metastasis. In the course of cancer progression, the ability of neoplastic cells to degrade the basement membrane and penetrate neighboring tissue (or blood vessels and lymph nodes) is an early event of the metastatic cascade. The Boyden chamber assay is one of the most prevalent methods implemented to measure the pro- or anti-invasive effects of drugs, investigate signaling pathways that modulate cell invasion, and characterize the role of extracellular matrix proteins in metastasis. However, the traditional protocol of the Boyden chamber assay has some technical challenges and limitations. One such challenge is that the endpoint of the assay involves photographing and counting stained cells (in multiple fields) on porous filters. This process is very arduous, requires multiple observers, and is very time-consuming. Our improved protocol for the Boyden chamber assay involves lysis of the dye-stained cells and reading the absorbance using an ELISA reader to mitigate this challenge. We believe that our improved Boyden chamber methodology offers a standardized, high-throughput format to evaluate the efficacy of various drugs and test compounds in influencing cellular invasion in normal and diseased states. We believe that our protocol will be useful for researchers working in the fields of immunology, vascular biology, drug discovery, cancer biology, and developmental biology.
Key features
• Measurement of tumor invasion using human cancer cells.
• Ability to measure the pro-invasive/anti-invasive activity of small molecules and biological modifiers.
• Measurement of chemotaxis, chemokines, trafficking of immune cells, and proteolytic activity of matrix metalloproteinases, lysosomal hydrolysates, collagenases, and plasminogen activators in physiological and pathological conditions.
• Investigation of the role of extracellular matrix proteins in the crosstalk between endothelial, epithelial, muscle, or neuronal cells and their adjacent stroma.
Keywords: Invasion Boyden Inserts Matrigel Chemoattractant Metastasis
Graphical overview
A schematic diagram depicting the steps of the Boyden chamber assay
Background
The process of metastasis is responsible for the majority of deaths in cancer patients [1]. Metastasis is a complex multi-step process comprised of primary tumor cell invasion into the surrounding tissue and extravasation into and intravasation out of the blood or lymphatic system, resulting in tumor cell colonization at distant sites and organs in the body [2,3]. Out of these processes, the process of tumor invasion has been the foundation of most drug-discovery studies. Invasion refers to the process by which cancer cells penetrate the basement membrane and launch themselves into circulation in the adjacent blood vessels or lymph nodes [4,5]. The Boyden chamber assay is the most prevalent method to measure this invasion capacity [6,7]. This assay requires two compartments separated by a microporous membrane coated with a Matrigel basement membrane matrix. In most versions of the Boyden chamber assay, cancer cells are placed in the upper compartment, and the lower compartment contains a robust chemoattractant. Under the influence of the chemoattractant, the cancer cells secrete proteases to degrade the Matrigel basement membrane, then migrate through the pores of the membrane into the lower compartment. A challenge to the traditional Boyden chamber assay is the arduous and time-consuming procedures of fixation and staining of the cells on the bottom surface of the porous membrane and subsequent photography and manual cell counting [6]. Our improved method of the Boyden chamber assay allows the results of the assay to be measured on a microplate reader. Therefore, our Boyden chamber assay protocol may be adapted to a high-throughput format. Currently, many research efforts are directed at improving our understanding of the metastatic cascade to advance our ability to design anti-cancer drugs, which will specifically target the process of metastasis. Out of all the events of metastasis, the process of tumor invasion has been the primary focus of drug discovery efforts [8,9]. The overarching goal of our improved Boyden chamber assay is to provide a simple, robust, and reproducible cell-based assay with a large-scale capability of evaluating and measuring the invasive ability of various cancer cells. We believe that our improved protocol of the Boyden chamber assay may pave the way for more rapid identification of novel anti-cancer drugs to combat metastasis.
Materials and reagents
Biological materials
DMS114 human small cell lung cancer (SCLC) cells [American Type Culture Collection (ATCC), catalog number: CRL-2066]. The cells should be grown to 70%–80% confluence in DMS114 culture media (see Recipe 3) and kept in a humidified environment at 37 °C (in the presence of 5% CO2) in a cell culture incubator [10]. Although we are describing the BCA protocol using DMS114 human SCLC cells, the pro-invasive activity of any relevant cancer cell line may be measured using this assay. Normal cells display pro-invasive activity under specific biological conditions. Such conditions may include treatment with growth factors, steroids, and carcinogenic chemicals. The BCA may be used to measure the effect of such chemicals on the invasion of normal cells.
Reagents
CorningTM BioCoatTM MatrigelTM invasion chamber inserts coated with Matrigel matrix. These invasion chambers are sold in two formats. The first is a 6-well format (Fisher Scientific, catalog number: 08-774-121). The second is a 24-well format (Fisher Scientific, catalog number: 08-774-122). Store at -20 °C. We use the 24-well format for the protocol described in the present manuscript
Sterile bovine serum albumin (BSA) solution at a concentration of 30% (w/v) in DPBS (Millipore-Sigma, catalog number: A7596)
Roswell Park Memorial Institute (RPMI)-1640 medium (ATCC, catalog number: 30-2001). Store at 4 °C, 1 M HEPES (Thermo Fisher, catalog number: 15630080)
Fetal bovine serum (FBS) (ATCC, catalog number: 30-2020). Long-term storage for FBS is at -70 °C. In our laboratory, FBS is aliquoted into 50 mL centrifuge tubes and stored at -20 °C. The FBS aliquot is thawed and stored at 4 °C for immediate use
Trypsin-EDTA (0.25% Trypsin, 0.53 mM EDTA) (ATCC, catalog number: 30-2101). Long-term storage for Trypsin-EDTA is at -20 °C. However, Trypsin-EDTA is aliquoted and stored at 4 °C for immediate use
1 M HEPES (Thermo Fisher Scientific, catalog number 15630080). Long-term storage for 1 M HEPES is at -20 °C. However, HEPES is aliquoted and stored at 4 °C for immediate use
Penicillin-streptomycin solution (ATCC, catalog number: 30-2300)
Dulbecco’s phosphate buffered saline (PBS) without calcium and magnesium (Corning, Fisher, catalog number: 21-031-CM). Store at 4 °C
Corning cell counting chamber (Fisher Scientific, catalog number: 07-200-988)
Capsaicin (Sigma-Aldrich, catalog number: M2028-50MG). Capsaicin is the hot and spicy ingredient of chili peppers. It is a potent agonist of transient receptor potential cation channel subfamily V member 1 (TRPV1). Store at 4 °C
PP2 is a potent and selective inhibitor of the Src family tyrosine kinases [11]. PP2 was manufactured by Enzo Biosciences (Enzo Biosciences, catalog number: 50-201-0680) and obtained from Thermo Fisher Scientific. The PP2 is delivered as powder and stored at -20 °C. The PP2 is dissolved in DMSO (Corning DMSO, Fisher, catalog number: MT-25950CQC) at a final concentration of 10 mM. The PP2 solution is aliquoted into microfuge tubes and stored at -20 °C
Dimethyl sulfoxide (DMSO), ACS certified (Fisher BioReagents, catalog number: D128-500). Store at room temperature
Aqua Solutions crystal violet 1% w/v aqueous (Fisher Scientific, catalog number: NC9002731). Store at room temperature
Solutions
Stock capsaicin (100 mM) (see Recipes)
WS-CAP-100X (2 mM) (see Recipes)
VEH-CAP-100X (see Recipes)
Stock PP2 (10 mM) (see Recipes)
WS-PP2-100X (500 µM) (see Recipes)
VEH-PP2 (see Recipes)
Crystal violet staining solution (see Recipes)
DMS114 culture medium (see Recipes)
RPMI-BSA (see Recipes)
Recipes
Reagent Final concentration Solvent Volume of each aliquot Total volume Storage
Stock capsaicin 100 mM DMSO 25 µL 100 µL -20 °C
WS-CAP-100X 2 mM RPMI/0.1% BSA Made fresh before use 100 µL On ice until use
VEH-CAP-100X 2% DMSO (v/v) Serum-free RPMI NA 100 µL Made fresh before use
Stock PP2 10 mM DMSO 25 µL 333 µL -20 °C
WS-PP2-100X 500 µM RPMI/0.1% BSA Made fresh before use 100 µL Made fresh before use
VEH-PP2-100X 5% DMSO (v/v) Serum-free RPMI NA 100 µL On ice until use
Crystal violet staining solution
0.25%
crystal violet
Milli Q Water NA 20 mL Room temperature
Stock capsaicin (100 mM), 1 mL
Weigh 30.5 mg of capsaicin in a sterile autoclaved amber-colored 1.5 mL microfuge tube. In a laminar flow hood, add 1 mL of DMSO to the tube containing capsaicin. Vortex to mix. Aliquot in sterile amber-colored 1.5 mL microfuge tubes. We typically put 25 µL of capsaicin stock solution in each aliquot. Store the aliquots at -20 °C. The aliquots are stable for one month at -20 °C. Laboratory personnel should wear gloves, mask, and face goggles when working with powdered capsaicin. Capsaicin can cause intense irritation if it comes in contact with the skin, face, or eyes.
WS-CAP-100X (2 mM), 100 µL
Take out an aliquot of 100 mM capsaicin solution from -20 °C. Put the tube in a 37 °C water bath for 5 min (until the entire solution is thawed). Vortex briefly to mix. Add 2 µL of 100 mM stock capsaicin to 98 µL of RPMI-BSA (in an autoclaved amber-colored 1.5 mL microfuge tube) in a laminar flow hood. Vortex to mix. Keep the solution on ice until used in the experiment. We make a fresh solution of WS-CAP before every assay and discard any leftover solution.
VEH-CAP-100X, 100 µL
Add 2 µL of DMSO to 98 µL of RPMI-BSA (in an autoclaved amber-colored 1.5 mL microfuge tube) in a laminar flow hood. Vortex to mix. We do not store the VEH-CAP-100X solution. We make a fresh solution of VEH-CAP before every assay and discard any leftover solution.
Stock PP2 (10 mM), 333 µL
Add 333 µL of DMSO to the bottle of 1 mg of PP2 in a laminar flow hood. Vortex to mix. Aliquot in autoclaved amber-colored 1.5 mL microfuge tubes (we put 25 µL of PP2 solution in each aliquot). Store the aliquots in -20 °C. The aliquots are stable for one month at -20 °C.
WS-PP2-100X (500 µM), 1 mL
Take out an aliquot of 10 mM PP2 solution from -20 °C. Put the tube in a 37 °C water bath for 5 min (until the entire solution is thawed). Vortex briefly to mix. Add 5 µL of stock solution of DMSO to 95 µL of RPMI-BSA (in an autoclaved amber-colored 1.5 mL microfuge tube) in a laminar flow hood. Vortex to mix. Keep the solution on ice until it is used for the experiment. We make a fresh solution of WS-PP2 before every assay and throw away the leftover solution.
VEH-PP2, 1 mL
Add 5 µL of DMSO to 95 µL of RPMI-BSA (in an autoclaved amber-colored 1.5 mL microfuge tube) laminar flow hood. Vortex to mix. We make a fresh solution of vehicle before every assay and do not store it.
Crystal violet staining solution, 20 mL
Add 5 mL of 1% crystal violet solution to 15 mL of Milli Q water. Vortex to mix. Store the solution at room temperature.
DMS114 culture medium
DMS114 cells were cultured following the published protocols [10]. All procedures are performed in a laminar flow hood. The DMS114 culture medium consists of RPMI-1640 with 1% Penicillin streptomycin solution (containing 100 units/mL penicillin, 50 µg/mL streptomycin), 25 mM HEPES, and 10% FBS. ATCC RPMI-1640 (30-2001 formulation) comes formulated with 10 mM HEPES. We then add an additional 7.5 mL of 1 M HEPES to 500 mL of RPMI-1640 to obtain a 25 mM concentration of HEPES. Store the media at 4 °C. To note, many laboratories add 50 mL of FBS to the full bottle of media to obtain 10% (v/v). We do not typically add FBS to the entire RPMI bottle. The rationale for this is to facilitate the most efficient use of each RPMI bottle. Many of our experiments require serum-free media, so we add the FBS as needed. The cells should be grown to 70%–80% confluence using the media described above and kept in a sterile cell culture incubator with a humidified environment, set at 37 °C with 5% CO2.
DMS114 culture medium Final concentration in media
Quantity or Volume
added to 500 mL media
RPMI-1640 NA 500 mL
Penicillin-streptomycin solution 100 units/mL penicillin, 50 µg/mL streptomycin 5 mL
HEPES 25 mM 7.5 mL
FBS 10% v/v 50 mL
RPMI-BSA
Add 100 µL of BSA from the bottle containing 30% BSA solution to 30 mL of serum-free RPMI-1640 (at room temperature) in a sterile 50 mL tube. Mix gently to avoid air bubbles. All these procedures should be performed in the laminar flow hood. We make fresh RPMI-BSA before each experiment. Store at room temperature for the duration of the assay.
DMS114 culture medium Final concentration in media Quantity or Volume
RPMI-1640 NA 30 mL
30% BSA solution 0.1% v/v 50 µL
Laboratory supplies
NuncTM EasYFlaskTM T-75 cell culture flasks (Thermo Scientific, catalog number: 156499)
Sterile Corning 24-well polystyrene tissue culture insert companion plate with lid (Corning Life Sciences, catalog number: 353504)
96-well plate, non-treated surface, pack of 25 (Thermo Scientific, catalog number: 12-566-202)
PuritanTM sterile DNA-free standard cotton swab, wood handle (Fisherbrand, catalog number: 22-025-201)
Sterile 15 mL polypropylene centrifuge tubes (Thermo Scientific, Nunc, catalog number: 12-556-017)
1.5 mL natural microcentrifuge tubes (Fisherbrand Premium, catalog number: 05-408-129). Autoclave before use
1.5 mL amber-colored microcentrifuge tubes (Fisherbrand Premium, catalog number: 05-408-134). Autoclave before use
Nunc 15 mL conical sterile polypropylene centrifuge tubes with plastic racks (Thermo Scientific, catalog number: 12-565-269)
Nunc 50 mL conical sterile polypropylene centrifuge tubes with plastic racks (Thermo Scientific, catalog number: 12-565-271)
Low-retention 2 mL microcentrifuge tubes (Fisherbrand, catalog number: 02-681-332)
Curved medium point stainless-steel general-purpose forceps (Fisherbrand, catalog number: 16-100-110). Autoclave the forceps before use with the Boyden chamber assay
Fine point high-precision forceps (Fisherbrand, catalog number: 22-327379). Autoclave the forceps before use with the Boyden chamber assay
Equipment
NU-540 laminar-flow biosafety cabinet (NuAire, Plymouth, MN, model: LabGard® ES NU-540 Class II, Type A2)
Cell culture incubator maintained at 37 °C and 5% CO2 (Thermo Scientific, Waltham, MA, model: Heracell VIOS 150i)
Microplate reader (Agilent, model: BioTek Gen 5)
Benchtop centrifuge (ThermoElectron Corporation, model: IEC Centra CL2)
Gilson PIPETMAN Pipettes [Marshall Scientific, catalog number: F144059M (P1000), F144058M (P200), F144056M (P10), F144054M (P2)]
Software and datasets
Agilent BioTek Gen 5 microplate reader and imager software
GraphPad Prism version 10.2.1
Procedure
Culture of DMS114 cells
The DMS114 human SCLC cell line was obtained from ATCC (Manassas, Virginia).
DMS114 cells were grown to 70%–80% confluence in DMS114 culture medium supplemented with 10% FBS (for instructions on making the media, see Recipe 8). Cells are cultured in T-75 tissue culture flasks. The flasks are kept in a cell culture incubator maintained in a humidified environment at 37 °C and 5% CO2 [10].
Working with Corning BioCoat invasion chambers (containing Matrigel matrix–coated inserts)
We will use the invasion chambers in the 24-well format to perform the Boyden Chamber Assay. A box of Corning BioCoatTM MatrigelTM invasion chamber is comprised of two packets containing two sterile 24-well plates loaded with the invasion chambers. The Corning BioCoatTM MatrigelTM invasion chambers are stored at -20 °C. We require one packet out of this box for the Boyden chamber assay. Keep the packet on the laboratory bench (at room temperature) for approximately 1 h so that the contents of the packet warm up to room temperature (Figure 1).
Figure 1. Packet of Matrigel-coated invasion chambers (in 24-well format)
Note: If you do not plan to use all of the inserts (inside a 24-well tissue culture plate) for your experiment, do not allow them to thaw, as repeated freeze-thaws will damage the Matrigel matrix barrier. Open the package (Figure 1) in a laminar flow hood (under aseptic conditions). With the help of sterile curved stainless-steel forceps, transfer the required number of invasion chamber inserts to a separate 24-well tissue culture companion plate to thaw. Wrap the unused inserts in the original packaging, tape them securely, and quickly return them to the -20 °C freezer.
Take the packet (containing one sterile 24-well plate loaded with the invasion chambers) and open it inside the laminar flow hood. The Corning BioCoat Matrigel invasion chambers consist of a tissue culture–treated sterile 24-well plate containing a total of 12 Matrigel-coated invasion chambers in the middle two rows of the plate (six invasion chamber inserts per row). The picture of the invasion chambers inside the 24-well companion plate is shown in Figure 2A. Figure 2B depicts the sideways view of the Matrigel-coated invasion chambers inside the 24-well plate.
Figure 2. Invasion chambers in a 24-well format. (A) Photograph of Matrigel-coated invasion chambers arranged in a 24-well companion plate. (B) Side-view image of the Matrigel-coated invasion chambers arranged in a 24-well companion plate.
The invasion chamber inserts have a polyethylene terephthalate (PET) porous mesh membrane at the bottom of the inserts. This PET membrane has a pore size of 8 μm. The PET membrane is coated with a thin layer of Matrigel basement membrane matrix (Figure 3).
Figure 3. Matrigel-coated invasion chamber inserts. The bottom of the insert comprises a polyethylene terephthalate (PET) membrane whose pore size is 8 μm and is coated with a thin layer of Matrigel.
Before starting the assay, draw the schema of the assay. Each sample is tested in duplicate (Figure 4).
Figure 4. Schema of the Boyden chamber assay
The inserts should be handled with autoclaved curved stainless-steel forceps, as shown in Figure 5. Do not touch the mesh at the bottom of the insert to avoid damaging the porous membrane or the Matrigel.
Figure 5. The Matrigel-coated invasion chamber insert should be handled with a pair of forceps
The inserts rest on the rims of each well and divide each well into two regions: the apical chamber, which is within the insert, and the basal chamber, which surrounds the outside of the insert (Figure 6).
Figure 6. The apical and basolateral chambers in the Boyden chamber assay
Ensure that the inserts align properly with the positioning notches of the companion 24-well plate. The outline of the rims on the companion plate has been indicated by black marker to better show the location of the notches (Figure 7). The black arrows indicate the positioning notches of the companion 24-well plate.
Figure 7. The positioning notches of the companion 24-well plate are indicated by black arrows
Add 500 µL of serum-free RPMI (at a temperature of 37 °C) to the apical chamber of the inserts using a P1000 Pipetman (Figure 8A and B). Take care not to touch the membrane with the tip or puncture it. Avoid generating air bubbles while adding the media to the apical chamber.
Figure 8. Rehydrating the apical chamber of the insert
Add 500 µL of serum-free RPMI (at a temperature of 37 °C) to the basal chamber (Figure 9). Place the P1000 tip in the empty space beside the insert (between the notches on the insert) and gently dispense the warm serum-free RPMI (Figure 9A–C). Avoid generating air bubbles while adding the media to the basolateral chamber.
Allow the inserts to rehydrate for 2 h in a humidified environment at 37 °C and 5% CO2 in a cell culture incubator (Figure 10). The rehydrated insert should be used immediately after the 2-h rehydration period. Do not wait to use the rehydrated inserts on the next day.
Figure 9. Addition of medium in the basolateral chamber of the invasion chamber. (A) Serum-free RPMI should be added in the space outside the insert or within the notches to avoid air bubbles. (B) Invasion chambers inside the companion plate. (C) Rehydrating the basolateral chamber of the insert.
Figure 10. Schematic diagram outlining the steps for rehydrating the insert
It is vital that there are no air bubbles trapped underneath the PET membrane in the basolateral chamber (Figure 11). If there are air bubbles, gently tap the side of the plate to dislodge the bubbles. Another strategy to remove air bubbles is to tilt the insert chamber at a slight angle to dislodge the bubbles from below the insert.
Figure 11. An air bubble trapped underneath the insert
Performing the Boyden chamber invasion assay: DAY 1
DMS114 cells were cultured in T-75 tissue culture flasks in DMS114 culture medium (see section A). The cells should be grown to 70%–80% confluence in a humidified environment at 37 °C (in the presence of 5% CO2) in a cell culture incubator [10].
Wash cells once with DPBS. Add the relevant volume of pre-warmed trypsin-EDTA solution (see Reagents) to the side wall of the flask. In our laboratory, we add 0.4 mL of trypsin-EDTA for every 10 cm2 of culture area in the flask. Therefore, we will add 3 mL of trypsin-EDTA to a T-75 flask. Gently rock the flask to ensure the trypsin solution covers the entire cell monolayer. Incubate the flask at 37 °C. The flasks should be placed flat (horizontally) in the cell culture incubator. Within 1–2 min, the cells will start rounding up and detaching from the bottom of the flask. Observe the cells using an inverted microscope to monitor the cell detachment process. Gently tap the side of the flask to fully detach the cells if necessary. Almost 90% of the cells should be detached and rounded up within 2 min of treatment with trypsin-EDTA.
Note: Prevent excessive exposure of the cells to trypsin solution (≥ 10 min).
Once cells appear detached, add 1 mL of FBS to inactivate the trypsin. Gently disperse the solution by pipetting up and down (several times) over the cell monolayer surface. Collect the cells in a 15 or 50 mL tube.
Note: Excessive forceful pipetting can cause cell damage.
Gently spin down the cells at 800× g for 5 min. Aspirate the media. Flick the tube to resuspend the cells. This usually loosens the pellet enough to resuspend the cells. Add 10 mL of DPBS to wash the cells and remove any traces of FBS. Centrifuge the cells at 800× g for 5 min. Aspirate the supernatant. Flick the tube to loosen the cell pellet and gently resuspend the cells in 1 mL of RPMI-BSA (Recipe 9).
Note: Do not vortex the cell suspension.
Count the cells using the Corning cell counting chamber. Adjust the concentration of the cells to 5 × 105 cells/mL in a 15 mL centrifuge tube using RPMI-BSA. For performing the assay as described in Figure 4, we will need 10 mL of cell suspension at a density of 5 × 105 cells/mL in RPMI-BSA. This tube will be referred to as TUBE-DMS114-FINAL.
Arrange six autoclaved 2 mL microcentrifuge tubes, labeled A–F, on a rack in the laminar flow hood.
Add the following drugs to the microcentrifuge tubes from step C6, as described in Figure 12 (columns 1–3, colored in green).
Figure 12. Protocol for treating DMS114 human SCLC cells to the indicated drugs (in 2 mL microfuge tubes labeled A–F) is described in Columns 1–3 (colored in green). The final concentrations of the drugs (in the 2 mL microfuge tubes labeled A–F) after the addition of DMS114 cell suspension is represented in Columns 4–5 (colored in yellow). The protocol for the addition of DMS114 cells (treated with/without the indicated drug) into the apical chamber of the Matrigel-coated inserts (fitted in the 24-well companion plate) is depicted in Columns 6–7 (colored in blue). Conc.= Concentration.
Add 1485 µL of DMS114 cell suspension (from TUBE-DMS114-FINAL) to each of the tubes A–E. Mix the cells with the drugs inside the tube by holding the tubes between your thumb and index finger and inverting the tubes 2–3 times. Do not vortex. Figure 12 describes the final concentrations of the drugs in the tubes (columns 4 and 5, colored in yellow).
Remove the rehydrated invasion chambers (along with the 24-well companion plate) from the 37 °C cell culture incubator. Gently remove the media from the apical and basal chamber using a Pipetman. DO NOT aspirate/puncture membrane.
Following the schema of the assay shown in Figure 4, take 500 µL of the cell suspension from TUBE A and add it to the apical chamber of the insert in well B1. Next, take 500 µL of the cell suspension from TUBE A and add it to the apical chamber of the insert in well C1. Continue loading the remaining inserts as described in Figure 12 (columns 6–7, colored in blue).
Add 500 µL of RPMI supplemented with 20% FBS (as the chemoattractant) in the basolateral chamber. Ensure that no air bubbles are trapped underneath the PET membrane in the basolateral chamber (refer to step B11). If there are air bubbles, gently tap the side of the plate to dislodge the bubbles. Another strategy to remove air bubbles is to tilt the insert chamber at a slight angle to dislodge the bubbles from below the insert.
Development of the Boyden chamber invasion assay: DAY 2
After 24 h, take the plate out of the cell culture incubator and place it on the laboratory bench.
Take out the inserts with a pair of forceps (Figure 13A) and gently place them upside down on a paper towel so that the solution inside the apical chamber is drained out on the paper towel (Figure 13B). Use caution not to touch or disrupt the basolateral side of the insert.
Figure 13. After 24 h, the insert is taken out with a pair of forceps (A) and drained on a Kimwipe (B)
Take the sterile DNA-free cotton swab and moisten it with warm serum-free RPMI medium (Figure 14A).
Insert the moistened cotton swab and remove all the cells (and the Matrigel matrix) from the upper surface of the membrane (apical surface) by applying gentle but firm pressure while moving the tip over the membrane. Hold the insert at an angle so the bottom of the membrane is not touching a flat surface and gently swab the upper side of the membrane, taking care to remove the cells but not detaching the insert membrane from the housing. This process is termed as “scrubbing” the membrane. Repeat the scrubbing with a second swab moistened with medium (Figure 14B).
Figure 14. Removing the non-invading cells from the invasion chamber using a moistened cotton swab. (A) A cotton swab moistened with serum-free RPMI. (B) Scrubbing the insert with a moistened cotton swab.
Figure 15. Staining the invasion chambers with crystal violet. (A) A solution of 0.25% crystal violet was added to each well of the companion plate. (B) The inserts were gently lowered into the crystal violet solution for staining.
Take a new companion plate. Add 400 µL of 0.25% crystal violet solution to each well of rows B1–B6 and C1–C6 of the companion plate (Figure 15A). Using a pair of forceps, gently transfer the scrubbed invasion chamber inserts into the companion plate in their corresponding location of the schematic (Figure 4) and incubate the inserts with the dye for 20 min at room temperature (Figure 15B).
Add distilled water (~750 mL) to three 1 L beakers. Using a pair of forceps, lift the insert (Figure 16A) and dip the insert in and out of water (in the first beaker) for about 2 min. Lightly invert the inserts and tap the corner of the inserts gently on a Kimwipe to remove any remaining liquid inside the insert. After this, dip the insert in and out of water (in the second beaker) for about 2 min. Lightly invert the inserts and tap the corner of the inserts gently on a Kimwipe to remove any remaining liquid inside the insert. After this, dip the insert in and out of water (in the third beaker) for about 2 min. Lightly invert the inserts and tap the corner of the inserts gently on a Kimwipe to remove any remaining liquid inside the insert.
Use a moistened sterile cotton swab to scrub the excess dye from the apical surface of the membrane and the walls of the insert (Figure 16B). Step D4 describes the correct procedure to scrub the membrane and the walls of the insert.
Figure 16. Removing the excess dye from the crystal violet-stained insert. (A) The invasion chamber insert stained with crystal violet solution. (B) Scrubbing the insert to remove the excess dye and remaining non-invading cells.
Take a new companion plate. Add 400 µL of DMSO to each well of the companion plate. Using a pair of forceps, gently transfer the invasion chamber inserts into the companion plate in rows B1–B6 and C1–C6. and incubate the inserts with the DMSO on an orbital shaker for 20 min at room temperature (Figure 17). The DMSO will lyse crystal violet–stained cells and a blue color will be observed.
Figure 17. Crystal-violet stained cells (on the underside of the filter) are lysed with DMSO to give a blue color
Remove the inserts from the plate. The old companion plate contains a blue-colored solution. Transfer 100 µL from each sample (in the companion plate) to a 96-well plate (each sample is tested in duplicate) and read the absorbance of the plate at 560 nm. Note that some ELISA readers have the capability of reading 24-well plates. In that case, the absorbance can be measured using a 24-well plate. In our laboratory, we measure the absorbance of the plates immediately after the transfer of the solution to the 96-well plate. We have no knowledge about how long we can wait (after the completion of the assay) to measure the absorbance of the plates at 560 nm.
Data analysis
Each sample/treatment was tested in duplicate in the Boyden chamber assay. The entire assay was repeated six independent times. Subtract the absorbance of the blank wells from all the samples. The absorbance obtained in DMS114 cells treated with the vehicle control (VEH-CAP wells) was taken as 1 and the absorbance obtained in the DMS114 cells treated with 20 µM capsaicin was represented as fold-change of vehicle control. A similar analysis was performed for the PP2-treated DMS114 cells, which represents the positive control for the assay.
Open GraphPad Prism (version 10.1.2). Using the numbers obtained in step 1 (described above), create a column graph of the data.
All data are plotted as mean ± standard deviation (SD). The data should be analyzed by performing a one-way ANOVA followed by Tukey’s post-hoc test. All analyses should be completed using a 95% confidence interval. Data is considered significant when P ≤ 0.05 (Figure 18).
Figure 18. Treatment of DMS114 cells with 20 µM capsaicin potently inhibits invasion as measured in the Boyden chamber assay. (A) The Src inhibitor PP2 was used as the positive control for the assay. (B) An MTT assay was done to demonstrate that none of these compounds inhibited the viability of DMS114 cells. Values represented by the same letter are not statistically significantly different from each other (P ≤ 0.05).
Validation of protocol
This protocol or parts of it have been used and validated in the following research article:
• Hurley et al. [12]. Non-pungent long chain capsaicin-analogs arvanil and olvanil display better anti-invasive activity than capsaicin in human small cell lung cancers. Cell Adhesion and Migration 11: 80–97.
General notes and troubleshooting
General notes
The Boyden chamber experiment is one of the most frequently used experimental techniques to evaluate cell invasion [6,7]. Apart from analyzing the pro-invasive activity of human cancer cells and biological agents, the Boyden chamber assay may be used to assess cell migration, haplotaxis [a directional movement of cells in response to signaling molecules that mediate cell adhesion like the extracellular matrix (ECM)], chemotaxis, cell motility, and chemokinesis (cell migration in response to soluble signaling molecules in the absence of a gradient, usually involving a change in migration speed or cytoskeletal reorganization) [6,7]. In our protocol, the endpoint of the Boyden chamber assay involved colorimetric detection using an ELISA reader. Other methods of endpoint detection (in the Boyden chamber assay) include fluorometric detection, chemiluminescence detection, X-gal staining, cell staining and counting (using microscopy techniques), and measurement of transepithelial electrical resistance (TEER). The Agilent xCELLigence RTCA DP analyzer evaluates cell invasion by the measurement of cellular impedance [13]. A few of these abovementioned Boyden chamber assay modalities have the capacity to be adapted for automation and high-throughput screening strategies.
It is always important to incorporate relevant positive and negative controls while performing a Boyden chamber assay [7,14]. For example, our Boyden chamber assay protocol evaluates the anti-invasive activity of capsaicin (at a concentration of 20 µM). It is possible that the treatment of DMS114 cells with 20 µM capsaicin may induce cell cycle arrest or kill off a substantial number of cells. If cells are quiescent (in G0) phase) they will not travel across the membrane of the invasion chambers. Similarly, if the cells are dying (from treatment with the test drug), they will be unable to invade across the membrane. It must be remembered that such factors may yield false results in the assay. Therefore, when determining the anti-invasive activity of drugs using the Boyden chamber method, an MTT and a proliferation assay must be done in parallel to make sure that the cells being treated remain healthy and viable throughout the assay. It is important to include positive controls in the experiment. For example, we have included the drug PP2, which is known to inhibit the invasion of cancer cells. Another control for the experiment may be primary normal cells, which undergo very little invasion. But it must be remembered that all cells (normal or cancerous) can acquire pro-invasive activity under the relevant physiological conditions.
Lastly, we discuss a few limitations of the Boyden chamber assay. The physiological relevance of transwell inserts to recapitulate invasion in vivo is poor [14,15]. The pore size of the membrane highly influences the number of cells that invade the membrane [14]. In its original format, the assay requires a large number of cells to obtain a sufficient signal, and the movement of cells through the filter cannot be visualized [6]. It can be difficult to control the concentration of the chemokine gradient, and this may produce aberrant or inconsistent results [14]. It is not possible to perform a dynamic determination of cell invasion since it is impossible to visualize the cells while they are migrating across the membrane. The data obtained from the Boyden chamber assay does not shed light on cell–cell and cell–matrix interactions [7,14,15]. An alternate method to measure cell invasion is the spherical invasion assay (SIA). The SIA analyzes the movement of human cancer cells as they migrate from a primary Matrigel layer, across the interface, and travel into a secondary Matrigel layer [16]. The SIA more accurately mirrors the actual process of invasion under physiological conditions [12]. The cells that grow in an ECM retain biological characteristics of tumors, such as responsiveness to diffusion gradient of oxygen, nutrients, and pH [17]. The growth of cells inside the ECM allows for complex cell–cell and cell–matrix interactions. The SIA can be adapted to organoids, spheroids [18], retinal angiogenic sprouts [19], tumor stem cells [20], neurospheres [21], and cells grown on polymeric scaffolds [22]. Such considerations reinforce the idea that the phenomenon of cell invasion should be studied (and validated) by using multiple experimental models. For example, when researchers are evaluating the anti-invasive activity of a drug, they should perform multiple independent assays like the Boyden chamber assay, microfluidic invasion assay, or spherical invasion assay and evaluate whether all these assays yield similar results, in order to infer that the drug has anti-invasive activity in vitro. Finally, the results obtained from the invasion experiments should be validated in an appropriate animal model to arrive at a definitive conclusion regarding the anti-invasive capacity of a drug or test compound.
Troubleshooting
Problem Possible cause Solution
Very low signal or no signal in the assay •Cells not seeded densely enough, resulting in too few cells migrating across the porous membrane •It is a good idea to standardize the optimal seeding density of cell numbers for the Boyden chamber assay. The optimal cell number may vary from cell line to cell line. Consider setting up a calibration assay varying the number of cells in a range of 20,000 cells/mL to 2 million cells/mL and perform the assay. Choose the cell number that yields the highest signal/background ratio.
• Insufficient incubation time. We measured the anti-invasive activity of capsaicin over 24 h. It may be possible that some compounds take longer to inhibit the invasion of cancer cells. Therefore, the optimal incubation time of the assay should be determined by performing a time-kinetics of the experiment. •Increase the incubation times for the assay.
• Insufficient concentration of crystal violet • We use 0.25% crystal violet for the Boyden chamber assay. You can increase the concentration of crystal violet to 0.5%.
• Air bubbles underneath the membrane • It is critical that the BioCoat Matrigel invasion chambers are properly seated in the notches. Make sure that there are no air bubbles underneath the membrane because the cells will not migrate through the dry patches.
• Membrane is damaged
• Do not puncture the membrane with a pipette tip or cotton swab
• Incorporate positive and negative controls for the assay (see General notes). Such controls may involve the use of known anti-invasive drugs and normal human cells (which invade minimally through the membrane).
• Variability in FBS • The chemoattractant in the bottom chamber is media containing 20% FBS. Make sure FBS is sourced from a reliable company. There is inherent lot-to-lot variability in FBS. Aliquot the FBS and store at -70 °C and utilize the same batch of FBS for all assay replicates. Avoid excessive freeze/thaw cycles of the FBS.
• Adding excessive FBS in the apical chamber
• The chemoattractant (media with 20% FBS) should be placed in the lower, or basolateral, chamber. It slowly diffuses into the upper, or apical, chamber setting up a chemotactic gradient for the cells to move towards the membrane, degrade the Matrigel, and travel to the underside of the membrane.
If too much FBS is present in the apical compartment, the chemotactic gradient will be compromised. Resuspend the cells (to be seeded in the apical chamber) in media containing 0.1% BSA. Do not exceed 0.4% FBS in the apical chamber.
• Cells have died or undergone cell cycle arrest during the assay
• The treatment of cells with various drugs (or test agents) may significantly reduce cell viability or cell cycle arrest. In that case, the cells may lack the capacity to invade across the membrane. Performing an MTT and a proliferation assay in parallel to the Boyden chamber assay will ensure that the cells are not quiescent/dying during the assay (see General notes).
• Incorporate positive and negative controls for the assay (see General notes). Such controls may involve the use of known anti-invasive drugs and normal human cells (which invade minimally through the membrane).
• Before lysing the cells with DMSO, look at the invasion chamber with a microscope to ensure that there are cells on the underside of the membrane
High background signal in the assay
• Insufficient washing of the inserts
• Refer to step D4 and D7for directions to efficiently wash the inserts
• Inefficient scrubbing of inserts to remove non-invading cells. • Refer to step D4 and D7for directions to efficiently scrub the inserts
• Impurities on the porous membrane • Avoid touching the membrane of the insert by hand or setting the invasion chamber on a dirty surface.
• Debris in cell culture flask or the cells are stressed out • Make sure that the cells are healthy, and the culture is free of bacterial contamination, debris, and foreign particles. It may be necessary to start a new culture from seed stocks or order new cells.
Variation between replicates
• Pipetting errors, insufficient replicates, improper washing of the inserts
• Use calibrated pipettes and avoid air bubbles
• Measure each sample in duplicate/triplicate
• Ensure that the membrane of the invasion chamber is not damaged. Damage to the membrane by forceps or cotton swabs can contribute to unexpected variation.
• Do not allow the membrane to dry out at any step.
• Refer to step D4 and D7 for directions to efficiently wash the inserts.
Acknowledgments
We acknowledge Dr. S. Chellappan and his laboratory for their continuous support. This work was supported in part by the WV-INBRE Administrative T3 Supplements for Research on Women’s Health in the IDeA States (Grant #. 3P20GM103434-23W1). The women’s Health T3 grant was a supplement of the West Virginia IDeA Network of Biomedical Research Excellence (WV-INBRE) grant (NIH grant P20GM103434; PI: Dr. G. Rankin). P.D. and M.A.V. are supported by a National Institutes of Health R15 Academic Research Enhancement Award (Grants 1R15CA161491-01A1 and 2R15CA161491-02). M.A.V. is also supported by NIH R15AI15197-01 and R15HL145573-01. S.D.R. is a recipient of NSF-SURE and WV-NASA Space Consortium undergraduate fellowships respectively.
Our improved protocol of the Boyden chamber assay was based on the following publications [23–25].
Competing interests
The authors declare no competing interests.
References
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Article Information
Publication history
Received: Apr 10, 2024
Accepted: Jun 30, 2024
Available online: Aug 2, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cancer Biology > Invasion & metastasis > Cell biology assays
Cell Biology > Cell-based analysis > Enzymatic assay
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5,041 | https://bio-protocol.org/en/bpdetail?id=5041&type=0 | # Bio-Protocol Content
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Microscale Thermophoresis (MST) as a Tool to Study Binding Interactions of Oxygen-Sensitive Biohybrids
BJ Bhanu P. Jagilinki *
MW Mark A. Willis *
FM Florence Mus
RS Ritika Sharma
LP Lauren M. Pellows
DM David W. Mulder
ZY Zhi-Yong Yang
LS Lance C. Seefeldt
PK Paul W. King
GD Gordana Dukovic
JP John W. Peters
(*contributed equally to this work)
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5041 Views: 798
Reviewed by: Joana Alexandra Costa ReisPhilipp A.M. SchmidpeterDjamel Eddine Chafai
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Original Research Article:
The authors used this protocol in Nano Letters Nov 2023
Abstract
Microscale thermophoresis (MST) is a technique used to measure the strength of molecular interactions. MST is a thermophoretic-based technique that monitors the change in fluorescence associated with the movement of fluorescent-labeled molecules in response to a temperature gradient triggered by an IR LASER. MST has advantages over other approaches for examining molecular interactions, such as isothermal titration calorimetry, nuclear magnetic resonance, biolayer interferometry, and surface plasmon resonance, requiring a small sample size that does not need to be immobilized and a high-sensitivity fluorescence detection. In addition, since the approach involves the loading of samples into capillaries that can be easily sealed, it can be adapted to analyze oxygen-sensitive samples. In this Bio-protocol, we describe the troubleshooting and optimization we have done to enable the use of MST to examine protein–protein interactions, protein–ligand interactions, and protein–nanocrystal interactions. The salient elements in the developed procedures include 1) loading and sealing capabilities in an anaerobic chamber for analysis using a NanoTemper MST located on the benchtop in air, 2) identification of the optimal reducing agents compatible with data acquisition with effective protection against trace oxygen, and 3) the optimization of data acquisition and analysis procedures. The procedures lay the groundwork to define the determinants of molecular interactions in these technically demanding systems.
Key features
• Established procedures for loading and sealing tubes in an anaerobic chamber for subsequent analysis.
• Sodium dithionite (NaDT) could easily be substituted with one electron-reduced 1,1'-bis(3-sulfonatopropyl)-4,4'-bipyridinium [(SPr)2V•] to perform sensitive biophysical assays on oxygen-sensitive proteins like the MoFe protein.
• Established MST as an experimental tool to quantify binding affinities in novel enzyme–quantum dot biohybrid complexes that are extremely oxygen-sensitive.
Keywords: Microscale thermophoresis Enzyme-quantum dot biohybrids Binding affinities Quantum dots Nanocrystals Nitrogenase Electron transfer Dissociation constant (Kd)
Graphical overview
Flowchart of the experimental workflow to determine the dissociation constant (Kd) between the MoFe protein and quantum dots (QDs). A) Preparation of the MoFe protein for the microscale thermophoresis (MST) experiment: The MoFe protein, which was originally purified in HEPES buffer, was buffer exchanged with MOPS-(SPr)2V• buffer using a Zeba column, diluted, and stored in liquid nitrogen. The MoFe protein in MOPS-(SPr)2V• buffer is diluted in MOPS-(SPr)2V•-Tween buffer and labeled with RED-tris-NTA second-generation dye to achieve the concentration required for the MST experiments. B) Experimental flowchart of the MST experiment: The Red dye–labeled MoFe protein and unlabeled QD solution stocks are prepared separately. Prepare a serial dilution of QD solution and mix them with an equal concentration of Red dye–labeled MoFe protein. Transfer the MoFe protein–QD mixture into capillary tubes individually and seal them with wax. Transfer the capillaries into Monolith [1] and perform the MST experiment, which is followed by data analysis and determination of the dissociation constant (Kd) between the MoFe protein and QDs. The top panel of the graph represents the changes observed in the fluorescence intensity during the MST experiment. The bottom panel is the graph depicting the calculated value of the Kd based on the observed fluorescence changes (Y-axis) as a function of the concentration of the interacting QD (X-axis). The lower part of the bottom panel represents the residuals, depicting the quality of the experimental fit.
Background
Quantification of biomolecular interactions is essential as they are critical to drug design, pharmaceutical, and bioprocessing industries. There are several techniques used to determine the strength of molecular interactions including isothermal titration calorimetry (ITC), surface plasmon resonance (SPR), nuclear magnetic resonance (NMR), and biolayer interferometry (BLI). Although these techniques have wide applications, there are several challenges associated with them in studying oxygen-sensitive samples. Microscale thermophoresis (MST), on the other hand, is a highly sensitive method where the intermolecular interactions are studied based on changes in fluorescence intensity of the target molecule as a function of the temperature-based difference in movement between bound and unbound states across a thermal gradient [1,2]. This Bio-protocol focuses on adapting the technique to analyze molecular interactions for highly oxygen-sensitive metalloenzymes and nanocrystalline quantum dots (QDs). We found that it was suboptimal to operate the NanoTemper MST within an anaerobic chamber, necessitating the development of a procedure to load and seal samples in an anaerobic chamber for analysis on the bench in the air. We achieved this by preparing the samples and sealing the MST capillaries inside the mBraun glove box, using capillary wax. Although this proved to be a successful strategy, there still was a need to protect extremely oxygen-sensitive samples like the MoFe protein and the Fe protein of nitrogenase from trace amounts of oxygen that might be introduced during sample preparation. Strong reducing agents like sodium dithionite (NaDT) are routinely used for this purpose; however, NaDT quenches the intensity of the fluorescence signal at even low concentrations. To overcome this, we have employed a novel, one electron-reduced viologen-based 1,1'-bis(3-sulfonatopropyl)-4,4'-bipyridinium radical anion [(SPr)2V•] that does not quench the fluorescence signal, instead of NaDT as a reducing agent in our experiments [3–5]. To check if the (SPr)2V• is a favorable and viable option, we have tested it for studying MoFe protein and Fe protein interactions. These proteins were selected because they are known to be extremely oxygen-sensitive; if our protocol with (SPr)2V• and sealing of the capillaries inside the mBraun works, this can be adapted as a reliable system to study interactions between any molecules that are oxygen-sensitive. After successfully validating our results, we have further developed MST protocols to study interactions of MoFe protein–cadmium sulfide (CdS) nanocrystal biohybrids. Specifically, we adapted the approach to examine CdS QDs of various sizes, which have been shown to interact with several metalloenzymes where the photoexcited conduction band electrons from the QDs could be transferred to the active site of the enzyme, triggering catalysis [6–9]. This emerging class of novel enzyme-QD biohybrids is of utmost significance as it could redefine bio-catalysis [10]. We have successfully demonstrated light-driven reduction of N2 to NH3 using novel MoFe-CdS nanocrystal biohybrids [11,12]. Although the reactivity of MoFe protein and CdS QD mixtures have been studied, there has been no characterization of the molecular interactions prior to our work [13]. We have now successfully developed the protocols for determining the dissociation constants (Kd) of these novel biohybrids involving MoFe protein and the CdS QDs. Here, we have articulated a detailed protocol for determining the Kd between the MoFe protein and 3.7 nm CdS QDs, which was reported in Pellows et al. [13].
Materials and reagents
Reagents
His-Tag labeling Kit RED-tris-NTA 2nd Generation (NanoTemper, catalog number: MO-L018)
Pierce BCA Protein Assay kit (Thermo Fisher, catalog number: 23225)
Capillary wax (Hampton Research, catalog number: HR4-328)
Liquid nitrogen
Double-distilled Milli-Q water
Sodium chloride (NaCl) (Sigma, catalog number: S9888-5KG)
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Sigma, catalog number: H3375-25G)
3-(N-morpholino) propanesulfonic acid (MOPS) (Sigma, catalog number: M1254-25G)
Sodium dithionite (Fisher Scientific, catalog number: 10274490)
Tween-20 (Sigma, catalog number: P1379-25ML)
One electron-reduced 1,1'-bis(3-sulfonatopropyl)-4,4'-bipyridinium [(SPr)2V•] (details in Procedure)
MoFe protein (details in Procedure)
3.7 nm CdS QD (details in Procedure)
Solutions
HEPES buffer (see Recipes)
MOPS-(SPr)2V• buffer (see Recipes)
MOPS-(SPr)2V•-Tween buffer (see Recipes)
QD buffer (see Recipes)
Recipes
HEPES buffer
Reagent Final concentration Amount
HEPES 50 mM, pH 7 119 mg
NaCl 150 mM 87.6 mg
Sodium dithionite (1 M) 2 mM 20 μL
H2O n/a 9 mL
Make the final volume to 10 mL
MOPS-(SPr)2V• buffer
Reagent Final concentration Amount
MOPS 50 mM, pH 7 104 mg
NaCl 150 mM 87.6 mg
(SPr)2V• 50 mM 1 mM 0.2 mL
H2O n/a 9 mL
Make the final volume to 10 mL
MOPS-(SPr)2V•-Tween buffer
Reagent Final concentration Amount
MOPS 50 mM, pH 7 104 mg
NaCl 10 mM 5.84 mg
(SPr)2V• 50 mM 50 μM 10 μL
Tween-20 (100%) 0.05% 5 μL
H2O n/a 9 mL
Make the final volume to 10 mL
QD buffer
Reagent Final concentration Amount
MOPS 50 mM, pH 7 104 mg
(SPr)2V• 50 mM 50 μM 10 μL
H2O n/a 9 mL
Make the final volume to 10 mL
Note: All buffers must be thoroughly degassed before moving them inside the mBraun glove box. This experiment requires only small amounts of buffer, and (SPr)2V• in buffers can degrade upon long-term storage.
Laboratory supplies
Monolith capillaries (NanoTemper, catalog number: MO-K022)
1.5 mL low protein binding microcentrifuge tubes (Thermo Fisher, catalog number: 90410)
0.2 mL thin-wall low protein binding microcentrifuge tubes (BIOplastics, catalog number: K77301)
0.5 mL low protein binding microcentrifuge tubes (Sarstedt Inc, catalog number: 72.704.600)
0.5 mL Zeba spin desalting columns, 7K MWCO (Thermo Fisher, catalog number: 89882)
50 mL tubes (VWR, catalog number: 89039-658)
50 mL tubes (VWR, catalog number: 89039-666)
10 μL pipette tips (VWR, catalog number: 470146-272)
200 μL pipette tips (VWR, catalog number: 470146-222)
1,000 μL pipette tips (VWR, catalog number: 470146-264)
Fisher brand 96-well PCR tube racks (Fisher Scientific, catalog number: 03-448-20)
VWR 80-well microtube racks (VWR, catalog number: 82024-469)
10 mL Wheaton crimp vials (Sigma, catalog number: DWK223686)
Aluminum crimp seals (VWR, catalog number: 97047-818)
Rubber stoppers (Grainger, catalog number: W224100-202)
500 µL gastight syringe (Hamilton, catalog number: 81265)
20 mL sample vials (Grainger, catalog number: 986710)
Equipment
Nanodrop (Thermo Scientific, NanoDrop One, catalog number: ND-ONEC-W)
Microscale thermophoresis (NanoTemper, model: Monolith NT.115)
Dry bath (VWR, Standard Dry Block Heater, catalog number: 75838-286)
Mini centrifuge (Eppendorf, MiniSpin, catalog number: 022620100)
Glove box (mBraun, model: UNIlab pro SP)
Vortexing machine (VWR, Analog Vortex Mixer, catalog number: 10153-838)
Software and datasets
MO.Control (NanoTemper, version: 1.6.1)
PALMIST software (The University of Texas Southwestern Medical Center, version 1.5.8)
Procedure
The (SPr)2V• and the 3.7 nm CdS QD used in this protocol have been synthesized as described previously [3,13]. The MoFe protein required for this protocol has been purified as previously described [14]. MoFe was initially at a concentration of 50 mg/mL in HEPES buffer, which was buffer exchanged into MOPS-(SPr)2V• and diluted 18 times. This dilution was necessary because MST is a highly sensitive technique that can detect interactions even at the nanomolar range.
MoFe protein buffer exchange
Note: This entire procedure must be performed inside the mBraun glove box.
Break the seal off the bottom of the Zeba spin desalting column and discard it (Note 1).
Slightly loosen the cap on the top of the column.
Place the column in the 1.5 mL collection tube.
Gently centrifuge the column at 1,500× g for 1 min to remove the storage solution (Note 2).
Discard the storage solution and place the column back onto the collection tube.
Equilibrate the column by adding 300 µL of MOPS-(SPr)2V• buffer to the top of it and incubate for 1 min.
Following the incubation, centrifuge the column for 1 min at 1,500× g to drain out the excess buffer.
Discard the buffer collected in the storage tube.
Repeat steps A6–A8 at least three more times to ensure the column is properly equilibrated with the MOPS-(SPr)2V• buffer.
Discard the collection tube and place the equilibrated column onto a clean low protein binding microcentrifuge.
Add 75 µL of concentrated MoFe protein to the top of the column.
Centrifuge the column at 1,500× g for 2 min.
Transfer the buffer-exchanged MoFe protein in MOPS-(SPr)2V• buffer and dilute as necessary to a final concentration of 11.6 μM to an appropriate tube for downstream experiments.
Cryopreserve excess MoFe protein by sealing in a 10 mL Wheaton crimp vial with a rubber stopper and aluminum cap, remove from the mBraun chamber, and use a 500 μL Hamilton syringe to rapidly (minimize O2 exposure) transfer the MoFe protein dropwise into liquid nitrogen. Transfer the frozen MoFe protein pellets into a clean needle-pierced 20 mL sample vial (to allow safe nitrogen gas venting) and store in liquid nitrogen.
MoFe protein concentration determination using bicinchoninic acid (BCA) assay
To determine the protein binding affinities accurately, it is essential to have the accurate value of the concentration of protein being used in the MST experiments. Thus, the concentration of protein being used is quantified using BCA assay (Note 3).
Note: This procedure can be performed on a regular workbench and does not need to be performed inside the mBraun glove box.
Prepare the working reagent (WR) by mixing the reagents A and B in the ratio of 50:1 (Note 4). In a clean 15 mL Falcon tube, dispense 5 mL of reagent A and add 0.1 mL of reagent B (Note 5).
For standards, make at least 10 different known concentrations.
Use the provided 2 mg/mL bovine serum albumin (BSA) as the standard stock solution (Note 6).
Make a dilution series of BSA from 2 to 0.2 mg/mL.
Take 10 clean 0.2 mL Eppendorf tubes and label them as 2.0, 1.8, 1.6, 1.4, 1.2, 1.0, 0.8, 0.6, 0.4, and 0.2.
Add double-distilled Milli-Q water in the following order: For 1.8 add 10 µL, for 1.6 add 20 µL, for 1.4 add 30 µL, for 1.2 add 40 µL, for 1.0 add 50 µL, for 0.8 add 60 µL, for 0.6 add 70 µL, for 0.4 add 80 µL, and for 0.2 add 90 µL. For 2.0, do not add water (Table 1).
Table 1. Reference table to make different known concentrations of the BSA sample
Tube # BSA (µL) Milli-Q water (µL)
2.0 100 0
1.8 90 10
1.6 80 20
1.4 70 30
1.2 60 40
1.0 50 50
0.8 40 60
0.6 30 70
0.4 20 80
0.2 10 90
The final concentration of the MoFe protein was determined to be 11.6 µM. The protein was then cryopreserved in liquid nitrogen until further use.
Now, add BSA to the labeled tubes as follows: For 2.0 add 100 µL, for 1.8 add 90 µL, for 1.6 add 80 µL, for 1.4 add 70 µL, for 1.2 add 60 µL, for 1.0 add 50 µL, for 0.8 add 40 µL, for 0.6 add 30 µL, for 0.4 add 20 µL, and for 0.2 add 10 µL.
Label another set of 12 clean 0.2 mL Eppendorf tubes and label them as 2B, 1.8B, 1.6B, 1.4B, 1.2B, 1.0B, 0.8B, 0.6B, 0.4B, 0.2B, PR1, and PR2 (PR1 & PR2 are the protein samples, whose concentrations are to be determined).
Dispense 25 µL each of all the standards that were made into the respective tubes. For example, dispense 25 µL from 2.0 and add it to 2.0B, and so on.
Use the PR1 and PR2 for the MoFe protein whose concentration is yet to be determined. Dispense 25 µL of the MoFe protein as it is into the PR1. Take a clean 0.2 mL Eppendorf tube and make a 1:1 dilution of the MoFe protein in water by adding 25 µL of the MoFe protein and 25 µL of the double-distilled Milli-Q water. Take 25 µL of the diluted MoFe protein and transfer it into PR2.
To all 12 tubes, add 200 µL of the WR, close the lid, and mix them thoroughly by vortexing.
Incubate the samples at 37 °C for 30 min in a dry bath.
Following the incubation time, cool the samples to room temperature.
Switch on the NanoDrop One and select proteins under the new experiment. In the proteins tab, select protein BCA method and use water as blank (Notes 7 and 8).
Use 2 µL of the sample and record the concentrations at 562 nm for all 12 samples individually (10 standards and 2 MoFe protein samples).
Make a standard curve by plotting the 562 nm absorbance data for each of the BSA standards on the Y-axis vs. the concentration in mg/mL on the X-axis. Use the standard curve to determine the protein concentration of each unknown sample (PR1 and PR2) (Note 9).
Testing of the binding efficiency of His-tagged MoFe protein with the RED-tris-NTA 2nd generation dye
To determine the binding efficiency of the His-tagged MoFe protein with the RED-tris-NTA 2nd generation dye, the MST experiment was performed (Note 10).
Note: This procedure must be performed inside the mBraun glove box until step C13.
Add 25 µL of MOPS-(SPr)2V•-Tween buffer to the vial containing the Red dye provided in the kit to obtain a final stock of 5 µM Red dye.
From the final stock of Red dye, prepare 200 µL of 50 nM working stock of the dye by diluting it 100 times. For this, dispense 2 µL of the 5 µM Red dye into a clean 0.5 mL microcentrifuge tube and add 198 µL of MOPS-(SPr)2V•-Tween buffer.
Dispense 30 µL of 4 µM MoFe protein in MOPS-(SPr)2V•-Tween buffer into a clean 0.2 mL thin-wall low protein binding microcentrifuge tube.
Arrange 16 clean PCR tubes, label them 1–16, and arrange them sequentially on a PCR tube holder.
Dispense 10 µL of MOPS-(SPr)2V•-Tween buffer to all PCR tubes except for the first tube, labeled “1.”
Following this, dispense 20 µL of 4 µM MoFe protein into the first tube, labeled “1.2.”
Make a serial dilution of 1:1 ratio from tube 1. For this, dispense 10 µL from tube 1 and transfer it into tube 2, mix it thoroughly, and using the same tip, transfer 10 µL from tube 2 into tube 3. Repeat this process until tube 16. After thoroughly mixing the sample in the final tube (16), remove 10 µL from that tube and discard it.
Now, add 10 µL of the 50 nM Red dye to all 16 tubes.
Incubate the samples at room temperature for 30 min.
Following the incubation, centrifuge the samples at 14,000× g for 10 min.
Transfer the sample from the PCR tubes into 16 clean Monolith capillaries sequentially.
Allow the samples to load into the capillaries via capillary suction (Figure 1A).
Following the transfer, gently hold the top end of the capillary with forceps and slowly dip the bottom end into the molten wax. Hold the capillary at 90° and allow the wax to solidify to seal one end (Figure 1B). Now, gently reverse the direction of the capillary tube to hold the sealed end of the capillary with the forceps. Gently dip the opposite end into molten wax and seal the other end (Figure 1C) (Note 11).
Figure 1. Workflow for the sealing of the capillary tubes. A) Allow the sample to load into the capillary via capillary suction. B) Hold the top end of the capillary at 90° with clean forceps and allow the wax to solidify to seal one end. C) Similarly, seal the other end by reversing the capillary.
Place the capillaries on the Monolith sample tray and remove the tray from the mBraun glove box.
Switch on the Monolith NT.115, gently place the sample tray (with the capillaries) in the tray holder, and close the lid.
Open the MO.Control version 1.6.1 software and open a new experiment. Under select excitation color tab, select Nano-RED and, in the next screen under choose an experiment tab, select expert mode.
In the expert mode, add the following experimental parameters, as shown in Figure 2: Excitation power, 80%; MST-power, medium; Before MST, 3 s, MST-on time, 15 s; and After MST, 1 s.
After setting up the experimental parameters, click Add dilution series at the bottom of the screen, highlighted in a green rectangular box in Figure 2.
Figure 2. Screenshot of the MO.Control version 1.6.1 software (NanoTemper) with the desired experimental parameters. After updating these parameters, click the Add dilution series button at the end of the window, highlighted in green.
Set the following parameters: Highest concentration for MoFe protein, 2,000 nM; Dilution, 1:1; Red dye concentration is maintained constant at 25 nM as described in Figure 3. After inputting these parameters, click Add at the bottom of the sub-window highlighted in a green rectangle box (Figure 3).
Figure 3. Screenshot of the MO.Control version 1.6.1 software depicting the information related to the dilution factor. After updating the necessary information, press the Add button, highlighted in a green box.
Following this, all experimental parameters, including the concentrations of the target (Red dye) and the ligand (MoFe protein), are reflected appropriately in the next screen as depicted in Figure 4. Now, start the experiment by clicking the Start measurement at the bottom of the screen, indicated in the green rectangular box (Figure 4).
Figure 4. Screenshot of the MO.Control version 1.6.1 software showing the final experimental parameters. Make sure all the experimental parameters reflect the originally planned experiment. Click the Start measurement button at the bottom of the window to begin the experiment.
Following the experiment, open the lid, gently remove the sample tray, and discard the capillaries. Now, analyze the dissociation constant (Kd) of the Red dye with the His-tagged MoFe protein using the PALMIST software. Note that the interaction between the Red dye and His tag is a reversible reaction. (For data analysis, refer to Data analysis section).
Determining the dissociation constant (Kd) between the Red dye–labeled MoFe protein with the 3.7 nm CdS QD
To determine the dissociation constant (Kd) between the His-tagged MoFe protein and the 3.7 nm CdS QD, MST was performed.
Prepare 100 µL of a 100 nM working stock of the Red dye solution. Add 2 µL of the 5 µM Red dye into a clean 0.5 mL microcentrifuge tube and add 98 µL of MOPS-(SPr)2V•-Tween buffer.
Dilute the MoFe protein in MOPS-(SPr)2V•-Tween buffer to a final concentration of 800 nM from its original stock of 11.6 µM in MOPS-(SPr)2V• buffer.
Transfer 90 µL of 800 nM MoFe protein in MOPS-(SPr)2V•-Tween buffer into a clean 0.5 mL microcentrifuge tube.
Add 90 µL of 100 nM Red dye solution in MOPS-(SPr)2V•-Tween buffer into the tube containing MoFe protein and mix the solution gently (Note 12).
Incubate the sample for 30 min at room temperature.
Following incubation, centrifuge the sample at 14,000× g for 10 min.
Following centrifugation, carefully transfer the supernatant into a new clean 0.5 mL microcentrifuge tube.
Finally, label the MoFe protein in MOPS-(SPr)2V•-Tween buffer with the Red dye; the final concentration is 400 nM.
Prepare the 3.7 nm CdS QD solution (see Note 13).
Dilute the original stock of 3.7 nm CdS QD solution (in Milli-Q water) into QD buffer to a final concentration of 20,000 nM.
Prepare 16 clean PCR tubes and label them 1–16.
Add 10 µL of QD buffer to all the PCR tubes except for the first tube, labeled “1.”
Transfer 40 µL of 20,000 nM 3.7 nm CdS QD to tube 1.
Make a 3:1 dilution series of 3.7 nm CdS QD solution. Transfer 30 µL of 20,000 nM 3.7 nm CdS QD from tube 1 into tube 2, mix it thoroughly, and, using the same tip, transfer 30 µL from tube 2 into tube 3. Repeat this process until tube 16. After thoroughly mixing the sample in the final 16th tube, discard 30 µL from the 16th tube.
Add 10 µL of the 400 nM Red dye–labeled MoFe protein to all 16 tubes and mix the solutions in each individual tube gently with the pipette tip.
Follow the steps as described in C9–17.
Set the following experimental parameters: Highest concentration for the 3.7 nm CdS QD, 10,000 nM; Dilution, 3:1; Red dye–labeled MoFe protein concentration is maintained constant at 200 nM as described in Figure 5. After updating these parameters, click Add at the bottom of the window (Figure 5).
Figure 5. Screenshot of the MO.Control version 1.6.1 software depicting the information related to the dilution factor of the 3.7 nm CdS QD. Following this, click the Add button highlighted in the green box.
All experimental parameters, including the concentrations of the target (MoFe protein) and the ligand (3.7 nm CdS QD) are reflected appropriately in the next screen, as depicted in Figure 6. Now, start the experiment by clicking Start measurement at the bottom of the screen, indicated in the green rectangular box (Figure 6). Note that the emission maximum of the Red dye is approximately 660 nm.
Figure 6. Screenshot of the MO.Control version 1.6.1 software showing the final experimental parameters for the MoFe protein and 3.7 nm CdS QD MST experiment. After ensuring all parameters are correct, click the Start measurement button at the bottom of the window to begin the MST experiment.
At the end of the experiment, open the lid, gently remove the sample tray, and discard the capillaries. Analyze the dissociation constant (Kd) between the MoFe protein and 3.7 nm CdS QD using the PALMIST software (Note 14).
Data analysis
The MST data analysis has been performed using the PALMIST software, which is freely available and can be downloaded from the following link: https://www.utsouthwestern.edu/research/core-facilities/mbr/software/. For more information on data analysis, refer to Scheuermann et al. [15].
Open the PALMIST software, click File, and then click the Read .moc File (Figure 7).
Figure 7. Screenshot of the PALMIST version 1.5.8 software showing the location of the Read. moc File under the File tab
Open the saved MST experiment data (.moc) from its original location.
After opening the data, select T-Jump at the bottom-right corner of the PALMIST software (Figure 8).
Check Normalize Fluorescence at the top-right of the window (Figure 8).
Click Fit (Hill equation) to determine the dissociation constant (Kd) (Figure 8).
Figure 8. Screenshot of the PALMIST version 1.5.8 software displaying the location of the T-Jump, Normalize Fluorescence, and Fit (highlighted in individual green boxes) and processed data along with the Kd value. The top panel shows the change in fluorescence of all the capillaries across the length of the MST experiment. The upper part of the lower panel shows the experimental fit of the data while the bottom part shows the residuals, displaying the quality of the data fitting in relation to the experimental data.
Validation of protocol
This protocol has been validated in Pellows et al. [13]. High affinity electrostatic interactions support the formation of CdS Quantum Dot:Nitrogenase MoFe protein complexes. Nano Letters (Figure 2 panels A and B, Figures 3 and 4).
General notes and troubleshooting
General notes
Zeba spin desalting columns are ideal for desalting unwanted buffer constituents prior to the MST experiments. These columns are highly efficient in recovering most of the protein, especially when working under very low volumes.
Make sure to mark the side of the column where the resin is positioned upward. Always place the column in the Eppendorf tube with the mark facing outward when placing the tube in the centrifuge in all the subsequent steps wherever centrifugation is necessary. Otherwise, the desalting efficiency could be reduced.
Although other assay kits are currently available, the BCA kit is highly recommended when working with metalloproteins as it is compatible with several reducing agents.
When both BCA reagents are mixed, the solution initially appears to be turbid; however, upon gently mixing, the turbidity should disappear resulting in a clear green solution.
Make sure you have enough volume of WR to carry out one set of standards and at least two unknown concentrations of the target protein. If you are using the test tube method, 2 mL of the WR is required per sample. For smaller volumes, up to 200 µL is required per sample.
Caution: Be careful while breaking the glass seal of the vial. Broken glass may cause injuries. After the usage, carefully discard the broken glass into a glass disposal box.
If Nanodrop is not available, a microplate reader can be used with the same protocol. Use a compatible microplate instead of 0.2 mL Eppendorf tubes.
If neither Nanodrop nor the microplate reader is available, readings can be measured by either a colorimeter or spectrophotometer. When using a colorimeter/spectrophotometer, larger volumes are required depending on the cuvette’s size and volume.
Repeat the BCA assay at least three times and make sure to have consistent values from the three replicates.
This experiment is mandatory to determine the binding affinity of the target protein to the Red dye and, once determined, this experiment does not need to be performed as long as using the same target protein for the MST experiments.
It is sometimes necessary to “double dip” the capillary end in the capillary wax to ensure sealing. Check that both ends of each capillary are sealed before removing them from the mBraun.
Using a sub-stoichiometric concentration of the Red dye relative to the target protein is recommended, as any excess unbound Red dye can interfere with fluorescence changes during MST experiments and significantly alter the Kd values during data analysis.
It is critical that the 3.7 nm CdS QD dilution in QD buffer is performed just prior to mixing with the MoFe protein and MST data collection. Also, prepare all buffers with the (SPr)2V• on the day of the experiments to maintain optimal oxygen scavenging.
A lower dissociation constant (Kd) implies higher binding affinities, whereas higher Kd values indicate weaker interactions.
Troubleshooting
Since the MoFe protein and the 3.7 nm CdS QD are extremely oxygen-sensitive, it is important to handle these samples without interference from atmospheric oxygen. Always transfer these samples in air-tight Wheaton crimp vials sealed with rubber stopper and aluminum crimp seals before moving the samples into the mBraun glove box.
For detecting binding affinities between molecules having weaker interactions, it is recommended to increase the LED intensity to 100% as long as photobleaching is not significant.
Certain non-biological samples may have less stability under certain solvent conditions. It is recommended to optimize their stabilities under different solvent conditions before performing MST experiments.
During MST experiments, if you observe aggregation behavior during data collection, try to optimize and improve the solvent conditions. Also, try to minimize the time between sample preparation and data collection to avoid aggregation during data collection.
Careful pipetting is a must, given the small volumes being used for making serial dilutions. Avoid small drops on the sides of tubes during serial dilution transfers since small errors can have a big impact on the results.
Acknowledgments
This work has been funded by the U.S. Department of Energy Office of Basic Energy Sciences, Division of Chemical Sciences, Geosciences, and Biosciences, Physical Biosciences and Solar Photochemistry Programs. This work was authored in part by the Alliance for Sustainable Energy, LLC, the manager and operator of the National Renewable Energy Laboratory for the U.S. Department of Energy (DOE) under Contract No. DEAC36-08GO28308. The U.S. Government and the publisher, by accepting the article for publication, acknowledge that the U.S. Government retains a nonexclusive, paid-up, irrevocable, worldwide license to publish or reproduce the published form of this work or allow others to do so for U.S. Government purposes. This protocol has been validated in Pellows et al. [13], published in ACS Nano Letters.
Competing interests
The authors declare no competing interests.
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Chaudhary, Y. S., Woolerton, T. W., Allen, C. S., Warner, J. H., Pierce, E., Ragsdale, S. W. and Armstrong, F. A. (2012). Visible light-driven CO2reduction by enzyme coupled CdS nanocrystals. Chem Commun. 48(1): 58–60.
Hamby, H., Li, B., Shinopoulos, K. E., Keller, H. R., Elliott, S. J. and Dukovic, G. (2019). Light-driven carbon−carbon bond formation via CO2reduction catalyzed by complexes of CdS nanorods and a 2-oxoacid oxidoreductase. Proc Natl Acad Sci USA. 117(1): 135–140.
Utterback, J. K., Ruzicka, J. L., Keller, H. R., Pellows, L. M. and Dukovic, G. (2020). Electron Transfer from Semiconductor Nanocrystals to Redox Enzymes. Annu Rev Phys Chem. 71(1): 335–359.
Brown, K. A., Harris, D. F., Wilker, M. B., Rasmussen, A., Khadka, N., Hamby, H., Keable, S., Dukovic, G., Peters, J. W., Seefeldt, L. C., et al. (2016). Light-driven dinitrogen reduction catalyzed by a CdS:nitrogenase MoFe protein biohybrid. Science. 352(6284): 448–450.
Brown, K. A., Ruzicka, J., Kallas, H., Chica, B., Mulder, D. W., Peters, J. W., Seefeldt, L. C., Dukovic, G. and King, P. W. (2020). Excitation-Rate Determines Product Stoichiometry in Photochemical Ammonia Production by CdS Quantum Dot-Nitrogenase MoFe Protein Complexes. ACS Catal. 10(19): 11147–11152.
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Article Information
Publication history
Received: Mar 27, 2024
Accepted: Jun 24, 2024
Available online: Jul 18, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biophysics > Bioengineering > Nanomaterials
Biochemistry > Protein > Fluorescence
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5,042 | https://bio-protocol.org/en/bpdetail?id=5042&type=0 | # Bio-Protocol Content
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Peer-reviewed
Generation of Multicellular 3D Liver Organoids From Induced Pluripotent Stem Cells as a Tool for Modelling Liver Diseases
SM Setjie W. Maepa
MM Mohlopheni J. Marakalala
HN Hlumani Ndlovu
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5042 Views: 692
Reviewed by: Komuraiah MyakalaHsih-Yin Tan Anonymous reviewer(s)
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Abstract
The liver is an essential organ that is involved in the metabolism, synthesis, and secretion of serum proteins and detoxification of xenobiotic compounds and alcohol. Studies on liver diseases have largely relied on cancer-derived cell lines that have proven to be inferior due to the lack of drug-metabolising enzymes. Primary human hepatocytes are considered the gold-standard for evaluating drug metabolism. However, several factors such as lack of donors, high cost of cells, and loss of polarity of the cells have limited their widescale adoption and utility. Stem cells have emerged as an alternative source for liver cells that could be utilised for studying liver diseases, developmental biology, toxicology testing, and regenerative medicine. In this article, we describe in detail an optimised protocol for the generation of multicellular 3D liver organoids composed of hepatocytes, stellate cells, and Kupffer cells as a tractable robust model of the liver.
Key features
• Optimising a protocol for generating multicellular 3D liver organoids from induced pluripotent stem cells.
Keywords: Stem cells Liver Hepatocytes Stellate cells Kupffer cells 3D organoids Drug-induced liver injury
Graphical overview
Background
The liver is a vital organ that is involved in the metabolism, synthesis, and production of serum proteins and bile acid and the detoxification of xenobiotics [1]. Earlier liver models relied heavily on cancer-derived cell lines, which have been proven to be inferior due to the suboptimal activity of drug-metabolising enzymes [2,3]. Isolated primary human hepatocytes (PHH) are considered to be the “gold standard” for drug metabolism and toxicity screening [4,5]. However, these cells display a rapid decline in the phenotypic function when cultured in traditional two-dimensional monolayer cell cultures; also, there is a scarcity of donors [5]. To overcome these limitations of using PHHs, researchers have developed various approaches including genetic modification of the cells and three-dimensional cultures combined with tissue engineering and media compositions [1,6,7]. Additionally, advances in cell culture systems offer a great opportunity to generate patient-specific hepatocytes using 2D cell-based [2] and 3D organoid platforms [1,8].
The liver cells can be obtained by direct differentiation of iPSCs [7,9] and tissue-derived stem cells [1,6,10] by exploiting inductive and repressive signals essential for liver ontogenic development. The 3D cultured cells display better physiologic and metabolic features of the native liver tissue in comparison with 2D cell-based cultures [1,9]. However, the vast majority of the reported methods predominantly differentiate cells into hepatic epithelial lineage only, thus lacking essential supportive cellular lineages such as profibrotic hepatic stellate cells (hematopoietic stem cells; HSCs) and inflammatory cells (Kupffer cells; KCs). Hence, they lack the capacity to model inflammatory diseases [1,11]. To circumvent these challenges, other researchers developed co-culture cell models based on mixing both the epithelial cells and supportive lineages from iPSCs [12,13]. However, these methods suffer from artefactual inflammation and fibrosis resulting from the difficulty in choosing the appropriate culture medium and extracellular matrix in which multiple cell lineages can be co-maintained [12,13].
A recent study by Ouchi and colleagues described a protocol for the generation of a multicellular human liver organoid (HLO) composed of hepatocyte-like cells, hepatic stellate–like cells, and Kupffer-like cells from iPSCs [9]. The cells aggregated to form a 3D liver organoid that was used to model steatohepatitis after the addition of free fatty acids to the culture media [9]. In this paper, we describe a method for generating multicellular 3D liver organoids (Figure 1) adapted from the protocols published by Ouchi and colleagues [9] and Mun and colleagues [8]. We further describe in detail how to generate these organoids and demonstrate their utility as a tool to study downstream assays for liver disease models.
Materials and reagents
Biological materials
Schistosome eggs antigen (SEA) (Schistosome Biological Supply Center, Theodor Bilharz Research Institute)
Reagents
A83-01 (R&D Systems, catalog number: R2939)
Activin A (R&D Systems, catalog number: 338-AL)
Accutase (Sigma-Aldrich, catalog number: A6964)
CellTiter-GLO® cell viability assay (Promega, catalog number: PRG9681)
CHIR99021 (Sigma-Aldrich, catalog number: SML1046)
Advanced DMEM (Gibco, catalog number: 12634-010)
B27 supplement (Gibco, catalog number: A1486701)
Bone morphogenetic protein 4 (BMP-4) (R&D Systems, catalog number: 314-13P)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A8531)
Dexamethasone (Sigma-Aldrich, catalog number: D4902)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D2650)
Dulbecco’s phosphate buffered saline (DPBS) (Gibco, catalog number: 14190-094)
Essential 8 basal medium (Gibco, catalog number: A1517001)
Epidermal growth factor (R&D Systems, catalog number: 236-EG)
Ethylenediaminetetraacetic acid (EDTA) (Invitrogen, catalog number: 15575-020)
Fibroblast growth factor 4 (R&D Systems, catalog number: 325-4F)
Fibroblast growth factor 10 (R&D Systems, catalog number: F8924)
Geltrex (Life Technologies, catalog number: A1413302)
HCl (Merck, catalog number: 100317)
Hepatocyte culture medium (Lonza, catalog number: CC-3198)
Hepatocyte growth factor (Sigma-Aldrich, catalog number: H9661)
HEPES (Gibco, catalog number: 15630080)
ImProm-II reverse transcriptase (Promega, catalog number: PRA3800)
Knockout serum replacer (KSR) (Gibco, catalog number: 10828028)
N2 supplement (Gibco, catalog number: A1370701)
Oncostatin M (R&D Systems, catalog number: 295-OM)
P450-Glo CYP3A4 assay with Luciferin-IPA (Promega, catalog number: PRV9002)
Penicillin-streptomycin (Gibco, catalog number: 15140-12221)
PowerUPTM SYBRTM Green Master Mix (Applied Biosystems, catalog number: A25741)
Retinoic acid (RA) (Sigma-Aldrich, catalog number: R2625)
Rho kinase (ROCK) inhibitor (Y-27632) (Sigma-Aldrich, catalog number: Y0503)
RPMI-1640, GlutaMAX supplement (Gibco, catalog number: 61870-036)
TRIzolTM reagent (Thermo Fisher Scientific, catalog number: 15596026)
TryLETM Express enzyme (Thermo Fisher Scientific, catalog number: 12604013)
Vascular endothelial growth factor (R&D Systems, catalog number: 293-VE)
Vitronectin (Gibco, catalog number: A27940)
Antibody list for organoid phenotyping (Table 1)
Table 1. List of antibodies used to phenotype human liver organoids
Phenotype Antibody Concentration Fluorophore Host Vendor Dilution
Epithelial cells EpCam 0.2 mg/mL BV421 Mouse BioLegend 1:80
Stellate cells CD166 5 µL (0.06 µg) PE-ALCAM Mouse eBioscience 1:160
Kupffer cells CD68 5 µL (0.05 µg) PE-Cy7 Mouse BioLegend 1:80
Antiretroviral and anti-tuberculosis drugs (Table 2)
Table 2. List of anti-retroviral and anti-tuberculosis drugs used to study liver injury
Drug Manufacturer Catalogue number Final concentration
Efavirenz (EFV) Sigma-Aldrich 025M478 7.39 mM
Tenofovir (TDF) Sigma-Aldrich SML1795 34.2 mM
Lamivudine (3TC) Sigma-Aldrich L1295 27.91mM
Rifampicin (RIF) Sigma-Aldrich R3501 28.42 mM
Isoniazid (INH) Sigma-Aldrich MKCF2223 20.04 mM
Primer sequences for RT-qPCR (Table 3)
Table 3. Primer sequences for stage-specific markers for liver organoids
Human stem cell markers
Gene name Forward primer sequence (5'-3') Reverse primer sequence (5'-3')
Oct-4 CAGGAGATATGCAAAGCAGAAAC GGCACTGCAGGAACAAATT
Nanog AGCCTAATCAGCGAGGTTTC CAGAGCAAGACTCCGTTTCA
Sox2 GCTACAGCATGATGCAGGACCA TCTGCGAGCTGGTCATGGAGTT
Definitive endoderm (DE) markers
Sox17 CGCACGGAATTTGAACAGTA GGATCAGGGACCTGTCACAC
GSC GAGGAGAAAGTGGAGGTCTGGTT CTCTGATGAGGACCGCTTCTG
Immature human liver organoids markers
CYP3A7 GAAACACAGATCCCCCTGAA TCAGGCTCCACTTACGGTCT
Mature human liver organoids markers
HNF4α CATGGCCAAGATTGACAACCT TTCCCATATGTTCCTGCATCAG
ALB TTG GCA CAA TGA AGT GGG TA AAA GGC AAT CAA CAC CAA GG
A1AT CCACCGCCATCTTCTTCCTGCCTGA GAGCTTCAGGGGTGCCTCCTCTGTG
CYP3A4 TGTGCCTGAGAACACCAGAG GTGGTGGAAATAGTCCCGTG
GADPDH CCATCTTCCAGGAGCGAG GCAGGAGGCATTGCTGAT
Solutions
Vitronectin (see Recipes)
0.5 mM ultrapure EDTA (see Recipes)
Activin A (see recipe)
Bone morphogenetic protein 4 (see Recipes)
Fibroblast growth factor 4 (see Recipes)
0.1% (m/v) bovine serum albumin (see Recipes)
CHIR99021 (see Recipes)
Retinoic acid (see Recipes)
Hepatocyte growth factor (see Recipes)
Dexamethasone (see Recipes)
Oncostatin M (see Recipes)
Fibroblast growth factor 10 (see Recipes)
Epidermal growth factor (see Recipes)
A83-01 (see Recipes)
Rho kinase protein (ROCK) inhibitor (Y-27632) (see Recipes)
Cell culture media
hiPSCs maintenance media: complete essential 8 medium (see Recipes)
Definitive endoderm induction (days 1–3) media: Complete RPMI medium (see Recipes)
Foregut spheroid (FGS) induction (days 4–6) media: Complete Advanced DMEM/F2 (see Recipes)
Human liver organoids (HLO) formation (days 7–10) media 1: Advanced DMEM/F12 + retinoic acid (see Recipes)
Hepatocyte maturation (days 11–25) media: Hepatocyte culture medium (HCM) (see Recipes)
Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR)
First strand CDNA synthesis cocktail 1 (see Recipes)
cDNA synthesis master mix 2 (see Recipes)
qRT-PCR reagents mix (see Recipes)
Recipes
Vitronectin solution
Reagents Final concentration Amount
Vitronectin 9 µg/mL 0.9 mg/mL
DPBS 1× 990 mL
Total n/a 1,000 mL
0.5 mM ultrapure EDTA
Reagents Final concentration Amount
Ethylenediaminetetraacetic acid 0.5 mM 10 mL
DPBS 1× 9,990 mL
Total n/a 10,000 mL
Store at room temperature (RT) and use on the day of preparation.
Activin A solution
Reagents Final concentration Amount
Activin A 100 µg/mL 10 µg
HCl 4 mM 100 mL
Total n/a 100 mL
Prepare 1,000 ng/mL working solution in RPMI-1640 + GlutaMAX and store at -20 °C until needed for the experiment.
Bone morphogenetic protein 4 (BMP-4) solution
Reagents Final concentration Amount
BMP-4 100 µg/mL 10 µg
0.1% BSA in 4 mM HCl n/a 100 mL
Total n/a 100 mL
Prepare 1,000 ng/mL working solution in RPMI-1640 + GlutaMAX and store at -20 °C until needed for experiment.
Fibroblast growth factor 4 (FGF-4) solution
Reagents Final concentration Amount
FGF-4 100 µg/mL 25 µg
0.1% BSA in PBS (Recipe 6) n/a 100 mL
Total n/a 100 mL
Prepare 1,000 ng/mL working solution in Advanced DMEM/F12 and store at -20 °C until needed for the experiment.
0.1% (m/v) bovine serum albumin (BSA) in PBS
Reagents Final concentration Amount
Bovine serum albumin (BSA) 0.1% (m/v) 0.1 g
H2O n/a 100 mL
Total n/a 100 mL
CHIR99021 solution
Reagents Final concentration Amount
CHIR99021 5 mg/mL 5 mg
DMSO n/a 1,000 mL
Total n/a 1,000 mL
Prepare 1 mM working solution in advanced DMEM/F12 and store at -20 °C until needed for experiments.
Retinoic acid (RA) solution
Reagents Final concentration Amount
Retinoic acid 50 mg/mL 50 mg
DMSO n/a 1,000 mL
Total n/a 1,000 mL
Prepare 1 mM working solution in advanced DMEM/F12 and store at -20 °C until needed for experiments.
Hepatocyte growth factor solution
Reagents Final concentration Amount
Hepatocyte growth factor (HGF) 100 mg/mL 5 mg
0.1% BSA in PBS (Recipe 6) n/a 50 mL
Total n/a 50 mL
Prepare 1,000 ng/mL working solution in hepatocyte culture medium (HCM) and store at -20 °C until needed for the experiment.
Dexamethasone (Dex) solution
Reagents Final concentration Amount
Dexamethasone (Dex) 10 mM 5 mg
DMSO n/a 1,000 mL
Total n/a 1,000 mL
Prepare 1,000 µM working solution in HCM and store at -20 °C until needed for the experiment.
Oncostatin M (OSM) solution
Reagents Final concentration Amount
Oncostatin M (OSM) 100 mg/mL 10 µg
0.1% BSA in PBS (Recipe 6) n/a 100 mL
Total n/a 100 mL
Prepare 1,000 ng/mL working solution in HCM and store at -20 °C until needed for the experiment.
Fibroblast growth factor 10 (FGF-10) solution
Reagents Final concentration Amount
Fibroblast growth factor 10 (FGF-10) 250 mg/mL 25 µg
0.1% BSA in PBS (Recipe 6) n/a 100 mL
Total n/a 100 mL
Prepare 1,000 ng/mL working solution in advanced DMEM/F12 and store at -20 °C until needed for experiments.
Epidermal growth factor (EGF) solution
Reagents Final concentration Amount
Epidermal growth factor 500 mg/mL 200 µg/mL
0.1% (m/v) BSA in PBS n/a 400 mL
Total n/a 400 mL
Prepare 1,000 ng/mL working solution in advanced DMEM/F12 and store at -20 °C until needed for experiments.
A 83-01 solution
Reagents Final concentration Amounts
A 83-01 10 mg/mL 10 mg
DMSO n/a 1,000 mL
Total n/a 1,000 mL
Prepare 1 mM working solution in advanced DMEM/F12 and store at -20 °C until needed for experiments.
Rho kinase protein (ROCK) inhibitor (Y-27632) solution
Reagents Final concentration Amount
Y-27632 10 mM 5 mg
DMSO n/a 1,000 mL
Total n/a 1,000 mL
Store at -20 °C for up to six months.
Cell culture media
hiPSCs maintenance media: complete essential 8 medium
Reagents Final concentration Amount
E8M 1× 485 mL
E8 supplement 1× 10 mL
P/S 1× 5 mL
Total n/a 500 mL
Alternatives: other hiPSC culture media can be used such as mTeSR1, StemFlex, and TeSR-E8. Swirl the bottle to mix contents properly, label with name and date prepared, filter, and store at 4 °C for up to 14 days. Alternatively, 50 mL aliquots can be made and stored at -20 °C.
Definitive endoderm induction (days 1–3) media: Complete RPMI medium
Regents Final concentration Amount
RPMI-1640 1× 482 mL
P/S 1× 5 mL
HEPES 25 mM 12.5 mL
Activin A 100 ng/mL -
BMP-4 50 ng/mL -
KSR 1× -
Total n/a 500 mL
Amount of Activin A, BMP4, and KSR are not shown on the Table as we prepare them fresh every day and they are dependent on the number of wells and volumes you are working with.
Swirl the bottle to mix contents thoroughly, label with name and date prepared, filter, and store at 4 °C for up to 14 days. Alternatively, 50 mL aliquots can be made and stored at -20 °C. Day 1: RPMI media supplemented with 100 ng/mL Activin A and 50 ng/mL BMP-4. Day 2: RPMI media supplemented with 100 ng/mL Activin A and 0.2% KSR. Day 3: RPMI media supplemented with 100 ng/mL Activin A and 2% KSR without BMP4 (see Notes 1–3).
Foregut spheroid (FGS) induction (days 4–6) media: Complete Advanced DMEM/F2
Reagents Final concentration Amount
Advanced DMEM/F12 1× 475 mL
B27 supplement 1× 5 mL
N2 supplement 1× 10 mL
GlutaMAX 1× 5 mL
P/S 1× 5 mL
FGF-4 - -
CHIR99021 - -
Total n/a 500 mL
Amounts of FGF-4 and CHIR99021 are not shown on the Table as we prepare them fresh every day and they are dependent on the number of wells and volumes you are working with. Swirl the bottle to mix contents thoroughly, label with name and date prepared, filter, and store at 4 °C for up to 14 days. Alternatively, 50 mL aliquots can be made and stored at -20 °C. Days 4–6 media: advanced DMEM/F12 supplemented with 3 µM CHIR99021 and 500 ng/mL FGF-4 (see Notes 1–3).
Human liver organoids (HLO) formation (days 7–10) media 1: Advanced DMEM/F12 + retinoic acid
Reagents Final concentration Amount
Advanced DMEM/F12 1× 475 mL
B27 supplement 1× 5 mL
N2 supplement 1× 10 mL
GlutaMAX 1× 5 mL
P/S 1× 5 mL
Retinoic acid 2 µM 1 mL
Total n/a 500 mL
Place the media in the dark or cover with foil.
Hepatocyte maturation (days 11–25) media: Hepatocyte culture medium (HCM)
Reagent Final concentration Amount
HCM basal medium 1× 500 mL
HGF - -
OSM - -
Dex - -
Total n/a 500 mL
Amounts for HGF, OSM, and Dex are not shown on the Table as we prepare them fresh every day in small quantities depending on the number of wells and volumes you are working with. Swirl the bottle to mix contents thoroughly, label with name and date prepared, filter, and store at 4 °C for up to 14 days. Alternatively, 50 mL aliquots can be made and stored at -20 °C. Days 11–25 media: supplement HCM medium with 10 ng/mL HGF, 0.1 µM Dex, and 20 ng/mL OSM.
Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR)
First-strand CDNA synthesis cocktail 1
Reagents Final concentration Amount
mRNA 2 µg -
Oligo dT primer n/a 1 µL
ddH2O n/a -
Total n/a 9 µL
Note: The amount of mRNA will depend on the concentration of RNA per sample; the volume of water will be calculated based on the concentration value of RNA.
cDNA synthesis master mix 2
Reagents Amount
5 first synthesis strand buffer 5 µL
dNTPs mix 1 µL
RNase inhibitor 1 µL
MgCl2 2 µL
ImPromp II reverse transcriptase 1 µL
ddH2O 6 µL
Total 16 µL
qRT-PCR reagents mix
Reagents Final concentration Amount
SYBR Green PCR Master Mix n/a 6.25 µL
Forward primer (10 µM) 10 µM 0.5 µL
Reverse primer (10 µM) 10 µM 0.5 µL
ddH2O n/a 4.25 µL
cDNA n/a 1 µL
Total n/a 12.5 µL
Laboratory supplies
Nunclon Sphera multi-well plates (Thermo Fisher Scientific, catalog number: 174930)
Sterile 15 mL Falcon tubes (Nest Biotechnology, catalog number: 602072)
Sterile 50 mL Falcon tubes (Nest Biotechnology, catalog number: 601001)
Equipment
Flow Cytometer (BD Biosciences, model: BD LSR Fortessa)
Biosafety cabinet suitable for cell culture (Nuaire)
Centrifuge (United Scientific, model: Orto Alresa)
Microplate Luminometer (Promega, model: GLOMAX 96)
Heat block (Acorn Scientific, model: Stuart)
Spectrophotometer (Thermo Fisher Scientific, model: NanoDropTM 2000)
Phase contrast/inverted microscope (Leica Microsystems, model: Leica DM IL LED)
Thermal Cycler (Thermo Fisher Scientific, model: QuantStudio 3 RT-PCR Systems)
Shellab CO2 Water Jacket HEPA Filter Incubator (United Scientific, model: SC05W-2)
Vortex machine (Labnet International)
Software and datasets
GraphPad Prism (GraphPad Software Inc.; version 5)
FlowJo software (Treestar; version 10)
Procedure
Coating plates with matrix
Vitronectin (VTN) solution
Thaw the stock vial of vitronectin (VTN) solution at RT or overnight at 4 °C.
Prepare 60 µL aliquots of 1 mL VTN and store at -80 °C until needed for the experiment.
For coating 6-well tissue culture plates or 35 mm dishes, dilute VTN (1:100 dilution, e.g., 60 µL in 6,000 µL in DBPS. Gently swirl the bottle to allow the solution to mix thoroughly.
Immediately dispense 1 mL of the VTN solution to as many wells of the 6-well plate as required, rock the plate to allow even distribution of the VTN solution across the surface of the plate, and incubate for 1 h at RT (see Note 1).
Geltrex
Thaw a 5 mL stock bottle of Geltrex overnight (16–20 h) on ice.
Working on ice and using chilled pipettes tips, mix well the contents without creating bubbles and aliquot into 1.5 mL cryotubes.
Label with name and date of preparation and store at -20 °C until needed for an experiment.
hiPSCs culture and organoid generation
Resuscitation of hiPSCs
Remove iPSCs from liquid nitrogen storage and thaw quickly in a 37 °C water bath.
Carefully add 1 mL of E8 + ROCK solution dropwise to the cryovial and transfer cells to a sterile 15 mL tube containing 8 mL of warm E8 + ROCK solution.
Pellet the cells by centrifugation at 398× g for 3 min at 4 °C.
Aspirate the supernatant and gently resuspend the pellet in 1 mL of warm E8 + ROCK solution (mix thoroughly by pipetting up and down). Transfer the cell suspension to one well of the pre-coated plate.
Agitate the plate gently within the tissue culture hood to ensure an even distribution of cells throughout the well and incubate cells at 37 °C and 5% CO2 overnight (see Note 5).
Check cell attachment under phase contrast microscope (see Note 6).
Change media daily by removing 95% of old medium using aspirator pipette. Cells require passaging upon 70%–80% confluency (see Note 7).
Passaging
Aspirate the old media and rinse the cells with 2 mL of DBPS.
Add 1 mL of 0.5 mM EDTA solution to the well to be passaged and rock the plate to cover the entire surface of the well.
Incubate at RT for 4–8 min and observe cells under a phase-contrast microscope.
Aspirate the 0.5 mM EDTA solution by tilting the plate forward slightly to collect the EDTA solution at the bottom edge of the well.
Immediately add 2 mL of complete E8 medium to the well and pipette the media to dislodge cell clusters (see Note 8).
Use the 2 mL of medium to gently wash the cells from the plate by pipetting the medium around the well approximately three times using a 5/10 mL serological pipette (see Note 6).
Dilute the cell suspension with complete E8 medium in a 15/50 mL tube at an appropriate cell density (in accordance with your desired split ratio, see Note 9).
Maintenance
Aspirate complete E8M from the hiPSCs and add 1 mL of Accutase solution per well to a 6-well plate. Place the plate in the incubator for 3–5 min, checking if cells detach. The cells are ready when they easily detach with gentle pipetting. Add 5 mL of complete E8 medium to a 15 mL Falcon tube.
Remove the plate from the incubator, add 1 mL of complete E8 medium, and lift the cells by gentle pipetting. Add this cell suspension in Accutase solution to the 5 mL complete E8 medium in the 15 mL Falcon tube. Gently pipette up and down twice to rinse off Accutase on the cells with complete E8 medium.
Centrifuge the cells at 398× g for 3 min at 4 °C.
Measure viable cells using Trypan blue exclusion method and calculate the number of viable cells/mL in the cell suspension.
For each cell line, optimal seeding cell density will need to be determined to achieve 90% confluency on day one.
For our hiPSCs cell line (N-N), 1.5 × 106 cells/well in a 6-well plate was ideal.
Aspirate the medium from the pellet and add 1 mL of complete E8M supplemented with 10 µM ROCK inhibitor solution to the VTN-coated 6-well plate. The ROCK inhibitor increases cell viability of the single cells in suspension. Add 1 mL of the cell suspension and the complete E8M supplemented with ROCK inhibitor solution and rock the plate back and forth and sideways to ensure even distribution of cells. Place the plate back in the incubator for 24 h.
At Day 0, warm complete E8M at RT. Confirm that the plated cells are healthy and growing at a uniformly high density. Aspirate the old medium and replenish the cells with warm complete E8M without ROCK inhibitor solution.
Place the plate back in the incubator for 24 h.
Step-by-step liver organoid method
Definitive endoderm (DE) induction (days 1–3)
To ensure a successful DE induction of hiPSCs, it is critical to start with high-quality hiPSCs. The cells should be at 80%–90% confluency with no sign of differentiation. Before beginning, warm complete E8 medium, Accutase, and base medium at RT.
Day 1: Cells should have reached 80%–90% confluency. Too low cell density will result in poor differentiation and cell death after Activin A solution treatment. Therefore, before beginning, titrate the cell number to ensure 80%–90% confluency of cells. It is possible to wait for an extra 24 h until the cells have reached the desired density.
Warm day 1 media at 37 °C. As Activin A solution treatment will result in cell death, prewarming the media at 37 °C increases the survival of cells. Aspirate complete E8M and add 2 mL/well of a 6-well plate of day 1 media and place the plate back in the incubator for 24 h.
Day 2: Check under the microscope if there still are layers of cells attached to the plate. If satisfied, aspirate floating dead cells and day 1 medium and replace with 2 mL/well for a 6-well plate of the warm day 2 media. Place the plate back in the incubator for 24 h. However, if there is little to no layer of cells attached, do not continue with the differentiation process and restart the protocol.
Day 3: Warm day 3 media at 37 °C. Aspirate Day 2 medium and replace with 2 mL/well for a 6-well plate of day 3 medium. Place the plate back in the incubator for 24 h.
Day 4: Endoderm cells should be confluent after differentiation (see Figure 1B). At this stage of the protocol, RT-qPCR and immunofluorescence can be performed to confirm gene-specific stage markers and cell localisation markers. Cells should express SRY-box transcription factor 17 (SOX17), Goosecoid (GSC), and Forkhead box protein A2 (FOXA2), and 85%–95% of the cells should co-stain double-positive for SOX17 and FOXA2.
Warm freshly prepared days 4–6 media at 37 °C. Each day, aspirate the media and replace with 2 mL/well of a 6-well plate of the prewarmed days 4–6 media. Owing to the number of cells present in the well, the media will turn yellow. Confirm the presence of 3D structures forming from the monolayer culture, including attached and floating spheroids (see Figure 1B).
Figure 1. Schematic of liver organoid differentiation protocol. (A) Schematic overview of the differentiation method for liver organoids. (B) Phase-contrast images of iPSCs-HLO depicting change in cell morphology. Scale bar = 100 µm. (C) Reverse transcription qPCR analysis of represented genes related to pluripotency or undifferentiated state and (D) definitive endoderm (DE) state and hepatic function. Undifferentiated iPSCs (iPSCs, n = 3), DE (n = 3), posterior foregut spheroid (foregut, n = 3), and HLO (n = 3). Data are presented as mean ± SEM (n = 3) and analysed by Student t-tests and one-way analysis of variance (ANOVA) using Dunnett’s multiple comparison as a post-hoc test. p < 0.05*, p < 0.001**, and p < 0.0001***.
HLO formation: enzymatic dissociation
Warm base medium, Accutase, and freshly made HLO formation medium at RT. In this step, collect the day 6 medium and floating spheroids into a 15 mL Falcon tube containing 5 mL of base medium. Add 2 mL of Accutase solution to each well and place the plate in the incubator for 3 min. Detach cells and pipette up and down four times. Aspirate and add cells–Accutase suspension to 5 mL of base medium.
Centrifuge cells at 398× g for 3 min at 4 °C. Completely aspirate the supernatant so it does not interfere with Geltrex.
Remove Geltrex from 4 °C and keep it on ice in the biosafety cabinet. Using chilled 1,000 µL pipette tips, use a P1000 pipette to mix the Geltrex thoroughly without introducing bubbles and add 1 mL of Geltrex to the pellet of cells in the 15 mL Falcon tube. Gently mix the contents by pipetting up and down several times to ensure even distribution of cells in the suspension. Keep the tube on ice.
For plating, use 24-well Nunclon Sphera multi-well plates. To each well of 24-well plate, add 40 µL (two 20 µL) drops of Geltrex–cell suspension at the centre of the plate forming a dome-shape.
Gently place the plate back in the incubator for 15–20 min to solidify Geltrex.
Add 500 µL of day 7 medium to each well of a 24-well plate and place it back into the incubator. There will be different sizes of 3D structures. Change medium every two days.
On day 9, 48 h later, warm the freshly made HLO formation medium at RT. Gently, aspirate old medium without disturbing the Geltrex drops and add 500 µL of fresh HLO formation medium per well of a 24-well plate.
Mechanical dissociation: cryopreservation of FGS
Alternative to HLO enzymatic dissociation step, we cryopreserve the FGS cells post mechanical dissociation of cells. Longer culture periods often introduce potential contaminations. This step is necessary for resting the FGS and is optional.
Use day 6 medium in each well and pipette up and down forcefully (5×) using a P1000 pipette.
Collect cell clusters and transfer into a 15 mL Falcon tube.
Centrifuge cells at 398× g for 3 min at 4 °C.
Aspirate supernatant and resuspend pellet in 1 mL of cryopreservation medium (30% basal advanced DMEM/F12, 60% FBS, and 10% DMSO).
Carry the cryopreservation tubes in a box containing cotton cloth dampened with isopropanol.
Store at -80 °C for two weeks or in liquid nitrogen for long-term storage.
FGS resuscitation & formation medium 2
FGS resuscitation & formation 2 consists of advanced DMEM/F12 + 5 factors (5 µM A83-01 solution, 20 ng/mL FGF-10 solution, 3 µM CHIR99021 solution, 20 ng/mL EGF solution, and 50 ng/mL VEGF solution).
Thaw FGS cells in organoid formation medium 2 supplemented with Y27632 solution (10 µM).
Centrifuge cells at 398× g for 3 min at 4 °C.
Completely aspirate the supernatant so it does not interfere with Geltrex.
Remove the Geltrex from 4 °C and keep it on ice in the biosafety cabinet. Using chilled 1,000 µL pipette tips, use a P1000 pipette to mix the Geltrex thoroughly without introducing bubbles and add 1 mL of Geltrex to the pellet of cells in the 15 mL Falcon tube. Gently mix the contents by pipetting up and down several times to ensure an even distribution of cells in the suspension. Keep the tube on ice.
For plating, see enzymatic dissociation section above.
On day 9, 48 h later, warm the freshly made HLO formation medium for 30 min at RT. Gently, aspirate old medium without disturbing the Geltrex drops and add 500 µL of fresh HLO formation medium per well of a 24-well plate every two for four days (see Note 11).
HLO maturation
On day 11, switch to HCM medium, which is hepatocyte-specific medium. Warm HCM to RT, aspirate HLO formation medium, and add 500 µL of HCM to each well of a 24-well plate.
Every 3 days, exchange the media by aspirating media in the well and adding 500 µL of fresh HCM.
Day 18: On this day, do not aspirate the media but pipette the media up and down to dislodge the cell drop. Add 1 mL of cold DBPS to dissociate the Geltrex until no pieces are visible. Place 250 µm cell strainer on a 15 mL tube and collect cells from each well and sieve through the cell strainer. This step ensures the removal of extra mesenchymal-like larger aggregates of cells. Also, this step is necessary where organoids will need to be individually sorted for downstream applications and sorted into 96- or 384-well plates. Warm fresh HCM at RT.
Add 4 mL of DBPS to the 15 mL Falcon tube with organoids and centrifuge at 398× g for 3 min at 4 °C.
Aspirate the supernatant, add HCM with 10% Geltrex to pelleted organoids, and pipette up and down to mix the cells. The cells will still be supported by the Geltrex matrix in suspension. Add 40 µL of Geltrex–cell suspension to each well of a new 24-well plate and place the plate back in the incubator.
Day 21: On this day, organoids can start to be used for downstream assays such as functional assays (see Figure 2A–2C) and staining. Confirm if the organoids are healthy and growing (see Figure 1B and 1D). No aspirations are needed for organoids not chosen for downstream assays; add 500 µL of fresh HCM and place the plate back in the incubator.
Perform quantitative reverse transcriptase polymerase chain reaction (qRT-PCR) for hepatic gene markers (Figure 1C–1D), enzyme linked immunosorbent assay (ELISA) to detect cytokines (Figure 4), and flow cytometry (Figure 2) to analyse the organoids.
Figure 2. Plot showing the percentage of EpCAM+, CD166+, and CD68+ cells. Human liver organoids (HLO) were examined for the co-expression of EpCAM+, CD166+, and CD68+ cells. Briefly, HLO were rinsed with DBPS, followed by dissociation into single organoids using the TrypLE express solution. Subsequently, dissociated HLO were incubated with FACS buffer solution containing EpCAM (1:80), CD166 (1:160), and CD68 (1:80) antibodies. The y-axis represents forward scatter (FSC-A), where an increased signal indicates the frequency of cells in HLO. The x-axis indicates the GFP fluorescence. The black lines are included to guide the eye to distinguish the fluorescence frequencies between EpCAM- cells and EpCAM+, CD166+, and CD68+ stained cells present in our HLO. (A) Percentage of EpCAM+ stained cells. (B) Percentage of CD166+ stained cells. (C) CD68+ stained cells of the whole organoids cell population after 21 days of HLO culture. EpCAM+ cells (60.4%), CD166+ (11.2%), and CD68+ (5.96%), respectively.
Total RNA isolation and RT-qPCR
Total RNA isolation
First, isolate the organoids from Geltrex using 1 mL of TrypLETM Express Enzyme solution in the 24-well plate and collect them from each well into a 15 mL Falcon tube.
Subsequently, rinse the organoids twice with 3 mL of DBPS followed by centrifugation at 398× g for 3 min at 4 °C until a thin layer of Geltrex remains.
Add 1 mL of TRIzolTM reagent to the pelleted organoids to extract total RNA following the manufacturer’s instructions.
Quantify extracted RNA using the ND-1000 Nanodrop spectrometer and store RNA at -80 °C until needed for further experiments.
Complementary DNA (cDNA) synthesis
For first-strand cDNA synthesis, reverse transcribe the total RNA using the ImProm-II reverse transcriptase following manufacturer's instructions, as shown in Recipes 1 and 2 of the Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR).
Mix 2 µg of template RNA, 1 µL Oligo dT, and ddH2O to a final volume of 9 µL. Heat the mixture for 10 min at 70 °C in a thermal block cycler with a heated lid to denature any secondary structure in the RNA and to minimise evaporation of solutions. Place the mixture on ice for 5 min.
Prepare 16 µL of the second cocktail [see Recipe 2 of Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR)] by adding and mixing reagents outlined in Recipe 1 of Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR), followed by centrifugation.
Incubate the mixture at 25 °C for 5 min to anneal oligo DT primers to the template cDNA, followed by 2 h of incubation at 42 °C.
Inactivate reverse transcriptase by heating the incubated mixture for 10 min at 70 °C, followed by stopping reaction and placing the tubes on ice. Synthesised cDNA may be diluted and stored at -20 °C until needed for further experiments.
qRT-PCR
Perform qRT-PCR in triplicates [see Recipe 3 of Quantitative reverse transcriptase polymerase chain reaction (qRT-PCR)] using PowerUpTM SYBRTM Green Master Mix and amplify synthesised cDNA on a thermal cycler with heated lid.
Calculate relative mRNA gene expression of 2-∆∆Ct method with GAPDH as a normalisation control. All primers (Table 3) were purchased from Integrated DNA Technologies (IDT).
Flow cytometry analysis
Optional: Before starting, coat the tubes and pipette tips with 1% BSA to reduce the loss of organoids that adhere to the sides of tips and tubes.
Collect organoids from wells of the 24-well plate into 15 mL Falcon tubes. If you want to process only one well, then collect from one well only. Add 6 mL of cold DBPS and gently pipette up and down to wash the Geltrex from the organoids.
Centrifuge the Falcon tube(s) at 398× g for 3 min at 4 °C. There will be a pellet of organoids at the bottom of the tube, a small layer of Geltrex in the middle, and media layered on top. Gently aspirate the media without disturbing the pellet and Geltrex layer.
Wash for a second time with 6 mL of DBPS and centrifuge the cells at 398× g for 3 min at 4°C.
After aspirating the DBPS, add 1 mL of TrypLETM solution and incubate for 10 min at RT to dissociate the organoids into single cells. After 10 min of incubation, add 4 mL of DBPS and centrifuge at 398× g for 3 min at 4 °C.
Use DBPS to wash out TrypLETM solution and centrifuge the cells again.
Aspirate the supernatant and resuspend cells in 100 µL of antibody + FACS buffer solution.
Incubate for 30 min at RT in the dark or cover the tubes with foil.
Add 100 µL of FACS buffer and centrifuge the cells at 398× g for 3 min at 4 °C.
Aspirate the supernatant and resuspend the pellet in 200 µL of FACS buffer.
Transfer the entire 200 µL of resuspended pellet into FACS tubes.
Acquire the samples on the BD LSR Fortessa flow cytometer (Figure 2).
Modelling liver injury
Antiretroviral (ARV), anti-tuberculosis drugs-induced hepatotoxicity
Perform steps E1–E5.
Aspirate the supernatant and resuspend cells in a mix of antiretroviral drugs or a mix of anti-retroviral drugs and anti-tuberculosis drugs.
Incubate the organoids with the drugs for 24 h at 37 °C in a 5% CO2 incubator.
After 24 h, take images of the organoids using a phase-contrast microscope (Figure 3A).
Collect the supernatants and analyse cytokine expression using ELISA (Figure 4).
Figure 3. Drug treatment of human liver organoids (HLO), day 23. HLO were rinsed with DBPS, followed by dissociation into single organoids using the TrypLE express solution. Subsequently, dissociated HLO were embedded in Geltrex and cultivated in 96-well sphere U-bottom plates for 24 h. HLO were further incubated with media only (UN), ART, Rifampicin (50 µM), and 1% Triton X-100 (TX-100) to determine cell viability using 3D GLO cell viability assay and CYP3A4 activity using the CYP3A4 GLO assay. (A) HLO phase contrast images of different treatment groups: untreated (UN), ART, rifampicin (RIF), TX-100 (Triton X-100). (B) Cell viability percentages. (C) CYP3A4 enzyme activity in untreated HLO, ART, and ART+TB treated HLO and 3D HepaRG model post 72 h of treatment. Data are presented as mean ± SEM (n = 3) and were analysed by one-way analysis of variance (ANOVA) using Dunnett’s multiple comparison as a post-hoc test. Ns: non-significant, p < 0.05*, p < 0.001**, and p < 0.0001***.
Figure 4. Inflammatory response after drug and pathogen treatment in human liver organoids (HLO). HLO were rinsed with DBPS, followed by dissociation into single organoids using the TrypLE express solution. Subsequently, dissociated HLO were embedded in Geltrex and cultivated in 96-well sphere U-bottom plates for 24 h. HLO were incubated with media only, ART & A+TB for 24 h, and SEA for 7 days. After each incubation time or treatment period, supernatants were aspirated and aliquoted onto the wells of a corresponding 96-well plate. Supernatants were used to carry out ELISA assay using BD Bioscience ELISA kit according manufacturer’s protocol. (A) Schematic diagram showing treatment duration for ART, A+TB- and SEA-induced liver injury. (B) Levels of proinflammatory mediators IL-6, (C) IL-1β, and (D) IL-4 in ART, A+TB post 24 h, and SEA post 7 days compared to UN counterpart. (D) Levels of the pathological mediator (E) TNF-α and (F) the anti-inflammatory mediator IL-10 compared to UN group. Data are presented as mean ± SEM (n = 7) and analysed by one-way analysis of variance (ANOVA) using Dunnett’s multiple comparison as a post-hoc test. p > 0.05ns, p < 0.05*, p < 0.001**, and p < 0.0001***.
Induction of inflammation with Schistosome eggs antigen (SEA)
Follow steps F1a–e described in the induction of hepatotoxicity with drugs.
Stimulate the organoids with 20 µg/mL of SEA and incubate the organoids for seven days at 37 °C in a 5% CO2 incubator.
After seven days, take the images using a phase contrast microscope (Figure 3A).
Collect the supernatants and analyse cytokine expression using the enzyme linked immunosorbent assay (ELISA) (Figure 4).
Cell viability determination using CellTiter-GLO® cell viability assay
Treat organoids with 0.05% DMSO, 1% Triton X-100, and a combination of antiretroviral drugs (ARVs) or rifampicin (RIF) for 24 h in a 5% CO2 incubator.
Remove the supernatant from each well. Add 100 µL of CellTiter-GLO® reagent to each well and 100 µL of hepatocyte culture media (HCM).
Mix the contents vigorously for 5 min to induce cell lysis.
Incubate the plate for 25 min at RT to stabilise the luminescent signal.
Read the plate using the GLOMAXTM 96 Microplate Luminometer (Figure 3B).
Cytochrome P450 Assay
Treat organoids or 3D HepaRG cells (control cell line) with troglitazone (TGZ) or rifampicin (RIF) for 24, 48, and 72 h. Vehicle control is 0.05% DMSO.
Remove the supernatants from each well. Wash cells twice with ice-cold PBS.
Add 50 µL of 50 µM P450-GloTM CYP3A4 (5 mM luciferin-BE) dissolved either in advanced DMEM or full HCM to each well.
Incubate the mixture at 37 °C in a 5% CO2 incubator for 4 h.
Collect the supernatants and transfer 25 µL of the supernatant to white-walled clear-bottom plates. Add 25 µL of luciferin detection reagent (reconstitution buffer with luciferase detection reagent).
Incubate the plate at 37 °C in a 5% CO2 incubator for 30 min.
Read the plate using the GLOMAXTM 96 Microplate Luminometer (Figure 4C).
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Mun et al. [8]. Generation of expandable human pluripotent stem cell derived hepatocyte-like liver organoids. Journal of Hepatology. 2019; 71:970–985. See Figure 2A.
Ouchi et al. [9]. Modeling Steatohepatitis in Humans with Pluripotent Stem Cell-Derived Organoids. Cell Metabolism. 2019; 30: 374–384.e6. See Figure 1A–D.
The robustness and reproducibility of our protocol can be evidenced from the above research articles.
General notes and troubleshooting
General notes
Store all media at 4 °C. Aliquot and store at -20 °C if required.
The growth factors (e.g., Activin, BMP-4, FGF-4, CHIR99021, Dex, HGF, RA, and OSM) should be added fresh every day.
Complete essential 8 medium, RPMI-1640 medium, advanced DMEM medium, and all growth factors require filtering before use. HCM does not require filtering.
Plates that have been coated with Vitronectin or Geltrex should be covered with parafilm and stored at 4 °C for up to one week. They should be clearly labelled with the date they were coated. Discard any plates not used within one week.
When plating cells, agitate the plate gently within the tissue culture hood to ensure even distribution of cells across the well, as colonies tend to settle in the centre of the plate affecting cell replating and differentiation.
After 24 h of plating iPSCs, check for cell attachment. If cell attachment is good, change medium to 2 mL of E8 media. If there are more cells floating than attached, then top up with 1 mL of freshly made E8 + ROCK solution.
Cells should be passaged when they have reached 70% confluency and are well compacted and the colonies have well-defined edges. Cells may also require passaging if levels of differentiation exceed that of iPSC or colonies start to look overgrown or unhealthy.
Split cells in the ratio of 1:4 to 1:6 (e.g., transferring all colonies from four or six wells).
When washing cells with 2 mL of E8 media after aspirating 0.5 mM EDTA solution, ensure to pipette the media up and down as this should dislodge cell clusters without dislodging any differentiated cells. Do not over pipette because it may result in single cells rather than cell clusters.
Forty-eight hours post FGS resuscitation, replace the resuscitation & organoid medium with organoid formation medium (supplemented with RA) every two days for a period of four days. Proceed with the protocol to HLO maturation.
Cryopreserved FGS-formed organoids get ready for downstream assays on day 23 instead of 21.
Acknowledgments
We would like to thank Dr Janine Scholefield from the Council for Scientific and Industrial Research (CSIR) in South Africa for generously gifting us with the induced pluripotent stem cell line. This study was funded by the South African Medical Research Council (SA-MRC) Self-Initiated Research grant and the National Research Foundation Competitive Support for Unrated Researchers (CSUR) grant (grant number 116260) awarded to Associate Professor Hlumani Ndlovu. This protocol was adapted from protocols developed by Mun et al. [8] and Ouchi et al. [9].
Competing interests
There are no competing interests to disclose.
Ethical considerations
No ethical considerations to disclose.
References
Huch, M., Gehart, H., van Boxtel, R., Hamer, K., Blokzijl, F., Verstegen, M. M., Ellis, E., van Wenum, M., Fuchs, S. A., de Ligt, J., et al. (2015). Long-Term Culture of Genome-Stable Bipotent Stem Cells from Adult Human Liver. Cell. 160: 299–312.
Zeilinger, K., Freyer, N., Damm, G., Seehofer, D. and Knöspel, F. (2016). Cell sources for in vitro human liver cell culture models. Exp Biol Med. 241(15): 1684–1698.
Yokoyama, Y., Sasaki, Y., Terasaki, N., Kawataki, T., Takekawa, K., Iwase, Y., Shimizu, T., Sanoh, S. and Ohta, S. (2018). Comparison of Drug Metabolism and Its Related Hepatotoxic Effects in HepaRG, Cryopreserved Human Hepatocytes, and HepG2 Cell Cultures. Biol Pharm Bull. 41(5): 722–732.
Godoy, P., Hewitt, N. J., Albrecht, U., Andersen, M. E., Ansari, N., Bhattacharya, S., Bode, J. G., Bolleyn, J., Borner, C., Böttger, J., et al. (2013). Recent advances in 2D and 3D in vitro systems using primary hepatocytes, alternative hepatocyte sources and non-parenchymal liver cells and their use in investigating mechanisms of hepatotoxicity, cell signaling and ADME. Arch Toxicol. 87(8): 1315–1530.
Pfeiffer, E., Kegel, V., Zeilinger, K., Hengstler, J. G., Nüssler, A. K., Seehofer, D. and Damm, G. (2014). Featured Article: Isolation, characterization, and cultivation of human hepatocytes and non-parenchymal liver cells. Exp Biol Med. 240(5): 645–656.
Broutier, L., Andersson-Rolf, A., Hindley, C. J., Boj, S. F., Clevers, H., Koo, B. K. and Huch, M. (2016). Culture and establishment of self-renewing human and mouse adult liver and pancreas 3D organoids and their genetic manipulation. Nat Protoc. 11(9): 1724–1743.
Siller, R., Greenhough, S., Naumovska, E. and Sullivan, G. J. (2015). Small-Molecule-Driven Hepatocyte Differentiation of Human Pluripotent Stem Cells. Stem Cell Rep. 4(5): 939–952.
Mun, S. J., Ryu, J. S., Lee, M. O., Son, Y. S., Oh, S. J., Cho, H. S., Son, M. Y., Kim, D. S., Kim, S. J., Yoo, H. J., et al. (2019). Generation of expandable human pluripotent stem cell-derived hepatocyte-like liver organoids. J Hepatol. 71(5): 970–985.
Ouchi, R., Togo, S., Kimura, M., Shinozawa, T., Koido, M., Koike, H., Thompson, W., Karns, R. A., Mayhew, C. N., McGrath, P. S., et al. (2019). Modeling Steatohepatitis in Humans with Pluripotent Stem Cell-Derived Organoids. Cell Metab. 30(2): 374–384.e6.
Garnier, D., Li, R., Delbos, F., Fourrier, A., Collet, C., Guguen-Guillouzo, C., Chesné, C. and Nguyen, T. H. (2018). Expansion of human primary hepatocytes in vitro through their amplification as liver progenitors in a 3D organoid system. Sci Rep. 8(1): 8222.
Kruitwagen, H. S., Oosterhoff, L. A., Vernooij, I. G., Schrall, I. M., van Wolferen, M. E., Bannink, F., Roesch, C., van Uden, L., Molenaar, M. R., Helms, J. B., et al. (2017). Long-Term Adult Feline Liver Organoid Cultures for Disease Modeling of Hepatic Steatosis. Stem Cell Rep. 8(4): 822–830.
Takebe, T., Sekine, K., Enomura, M., Koike, H., Kimura, M., Ogaeri, T., Zhang, R. R., Ueno, Y., Zheng, Y. W., Koike, N., et al. (2013). Vascularized and functional human liver from an iPSC-derived organ bud transplant. Nature. 499(7459): 481–484.
Nie, Y. Z., Zheng, Y. W., Ogawa, M., Miyagi, E. and Taniguchi, H. (2018). Human liver organoids generated with single donor-derived multiple cells rescue mice from acute liver failure. Stem Cell Res Ther. 9(1): 5.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Workflow for establishment, expansion, and differentiation of iPSCs-HLOs.
Figure S2. Phase-contrast images representative of iPSCs-HLO at different days of the differentiation protocol.
Table S1. Drugs and antigen treatment groups.
Article Information
Publication history
Received: Dec 5, 2023
Accepted: Apr 30, 2024
Available online: Jul 18, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Detection and Quantification of Programmed Cell Death in Chlamydomonas reinhardtii: The Example of S-Nitrosoglutathione
LL Lou Lambert
AD Antoine Danon
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5043 Views: 413
Reviewed by: Luis Alberto Sánchez VargasAdrian Pascal NievergeltShuhei Ota
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Original Research Article:
The authors used this protocol in Plant Physiology Apr 2024
Abstract
Chlamydomonas (Chlamydomonas reinhardtii) is a unicellular model alga that has been shown to undergo programmed cell death (PCD) that can be triggered in response to different stresses. We have recently shown that Chlamydomonas is particularly well suited to the study and quantification of PCD. We have shown for the first time that S-nitrosoglutathione (GSNO), a nitric oxide (NO) donor, is able to induce PCD and can be used as a study system in Chlamydomonas. In this article, we provide a simple and robust protocol for quantifying GSNO-induced PCD, which can be adapted to any other treatment. We explain how to detect NO production in the cell following GSNO treatment. We show how PCD can be identified simply by analyzing the degradation profile of genomic DNA. We also provide an easy and reproducible cell death quantification protocol, which makes it possible to follow the course of PCD over time and highlight very fine differences in the number of affected cells between different samples.
Key features
• Use of S-nitrosoglutathione (GSNO) as a means to study programmed cell death (PCD) in Chlamydomonas.
• Discrimination of PCD vs. necrosis.
• In vivo determination of NO production in the cell.
• A simple, robust protocol for PCD quantification.
Keywords: Chlamydomonas reinhardtii Programmed cell death S-nitrosoglutathione Nitric oxide DNA fragmentation Evans blue staining
Graphical overview
Background
Programmed cell death (PCD) is a crucial process identified in animals in 1972 [1], which plays a role in a wide range of biological processes. Understanding targeted disappearance of cells whose presence is no longer desired is straightforward in multicellular organisms. However, the concept of PCD is more difficult to apply to a unicellular organism, since the death of the cell corresponds to the death of the organism. In recent years, however, it has been shown that PCD does exist in unicellular organisms such as Chlamydomonas, where it was found to be an altruistic mechanism that allows the survival of the population [2,3]. Nitric oxide (NO) has been shown in several studies to be an important molecule for PCD in Chlamydomonas [4,5]; this is why we used the main source of NO in the cell, S-nitrosoglutathione (GSNO), as a system for studying PCD in Chlamydomonas [3]. Several specific criteria can be used to discriminate programmed cell death from necrosis; one of the simplest is to analyze the DNA degradation profile during death [6]. If PCD occurs, DNA migrates as multiples of 180 bp in gel electrophoresis, resulting in a DNA ladder; in the case of necrosis, continuous DNA degradation represented by a smear will be observed [7]. We describe how to analyze the DNA degradation profile in Chlamydomonas, as well as a simple and robust method for calculating the percentage of dead cells in a population. In the case of GSNO-induced PCD, it is important to be able to quantify NO in the cell after treatment; here, we explain how to implement a method for doing so in vivo, using a fluorescent probe. Our protocol outlines the steps to detect and quantify PCD in Chlamydomonas using Evans blue staining, whether you are using GSNO as an inducer or any other treatment.
Materials and reagents
Biological material
We used Chlamydomonas reinhardtii D66 (CC-4425) [8] and CLiP library (CC-4533) [9] strains, but any strain could be used
Reagents
Note: Unless specified otherwise, the reagent can be stored at room temperature.
TAP (tris-acetate-phosphate) solution (Life Technologies, catalog number: T8050)
S-nitrosoglutathione (GSNO) produced in our laboratory but also available for purchase (Sigma, catalog number: N4148), stored at -20 °C
Evans blue powder (Sigma, catalog number: E2129)
Phenol:chloroform:isoamyl (25:24:1) (Sigma, catalog number: 77617), stored at 4 °C
Ethanol 100% (Sigma, catalog number: 32205-M)
Trizma® base (Sigma, catalog number: 93350)
Hydrochloric acid (HCl) (Sigma, catalog number: 258148)
Sodium chloride (NaCl) (Sigma, catalog number: S7653)
Ethylenediaminetetraacetic acid tetrasodium (EDTA) (Sigma, catalog number: E6511)
Sodium dodecyl sulfate (SDS) 20% (Sigma, catalog number: 05030)
Sodium acetate (Sigma, catalog number: S2889)
RNase A, 10 mg/mL (Thermo Scientific, catalog number: EN0531), stored at -20 °C
Agarose (Sigma, catalog number: A9539)
Tris borate EDTA (TBE) buffer (Sigma, catalog number: T4415)
Ethidium bromide solution (Sigma, catalog number: 46067)
GeneRuler 100 bp Plus DNA ladder (Thermo Scientific, catalog number: SM0321), store at 4 °C
DAF-FM diacetate (4-amino-5-methylamino-2’,7’-difluorescein diacetate) (Invitrogen, catalog number: D-23844), store at -20 °C, protected from light
Solutions
50 mM GSNO solution (see Recipes)
DNA extraction buffer (see Recipes)
Sodium acetate 3.3 M (see Recipes)
TBE 0.5× solution (see Recipes)
Evans blue solution (see Recipes)
Recipes
50 mM GSNO solution
Weigh 20 mg of GSNO and dissolve it in 800 µL of TAP medium [10] in a 1.5 mL tube.
Verify the concentration of the 100-fold diluted solution using the molar extinction coefficient of GSNO (922 M-1·cm-1 at 335 nm) [11].
Note: It is better to prepare fresh GSNO for each experiment to avoid its oxidation.
DNA extraction buffer (for 500 mL)
Reagent Final concentration Amount
Tris-HCl (1 M, pH 7.5) 200 mM 100 mL
NaCl (1 M) 250 mM 125 mL
EDTA (0.5 M, pH 8) 25 mM 25 mL
SDS (20%) 0.5% 12.5 mL
MilliQ water n/a 262.5 mL
Total n/a 500 mL
Note: It is recommended to autoclave extraction buffer (add SDS after autoclaving) before proceeding to DNA extraction.
Sodium acetate 3.3 M (for 100 mL)
Reagent Final concentration Amount
Sodium acetate 3.3 M 27 g
MilliQ water n/a 100 mL
Total n/a 100 mL
Dissolve using a magnetic stirrer and stir bar.
Filter sterilize (0.22 µm).
TBE 0.5× solution
Reagent Final concentration Amount
TBE (10×) 0.5× 100 mL
MilliQ water n/a 1.9 L
Total n/a 2 L
Evans blue solution
Reagent Final concentration Amount
Evans blue powder 0.5 % 0.5 g
TAP n/a 100 mL
Total n/a 100 mL
Filter sterilize (0.22 µm).
Laboratory supplies
Spectrophotometer semi-micro cuvette (Biosigma, catalog number: BSA002)
24-well plates (Evergreen Labware, Medical Caplugs, catalog number: 222-8044-01F)
Surgical tape (MicroporeTM, catalog number: 1530-0)
Counting chamber Neubauer (Blaubrand, catalog number: BR717805)
Equipment
Spectrophotometer (Implen Nanophotometer)
Microplate reader (ClarioStar Plus, BMG LABTECH)
Orbitron Rotary shaker (Infors)
Olympus BX43 microscope
Thermomixer (Eppendorf, catalog number: 5382000015)
Vortex (Dutsher Vortex Genie 2, catalog number: 079008)
Electrophoresis tanks (Embitec® runOneTM, model: EP-2000)
Camera (Q-IMAGING Micropublisher 3.3 RTV)
NanoDrop One (Thermo Scientific)
GelDoc Go Imaging System (Bio-Rad)
Software and datasets
Image Lab v6.1 for the GelDoc Go Imaging System (Bio-Rad)
Smart control data analysis for ClarioStar Plus (MARS, BMG LabTech)
Procedure
Cell culture
Cultivate Chlamydomonas cells in TAP liquid media at 25 °C, under continuous light (50 µmol photons/m/s) with shaking at 120 rpm until you obtain a concentration of 4 × 106 to 6 × 106 cell/mL.
Transfer 1 mL of culture to as many wells as required in a 24-well plate.
Add 20 µL of a 50 mM fresh solution of GSNO per well.
Close the 24-well plate with Micropore surgical tape and place it at 25 °C under continuous light (50 µmol photons/m/s) with shaking at 120 rpm.
NO detection after GSNO treatment
Add the DAF-FM diacetate to your sample at a final concentration of 5 µM (e.g., 1 µL of a 5 mM solution in a final volume of 1 mL) and place the cells under low light (e.g., 10 µmol photons/m/s) at 25 °C under agitation (120 rpm) for 30 min.
Centrifuge cells at 2,300× g for 3 min, remove the supernatant, and replace it with the same volume of TAP media. Repeat twice.
After a 15 min incubation period, add GSNO to your sample at a final concentration of 1 mM (e.g., 20 µL of a 50 mM solution in a final volume of 1 mL).
NO can be detected as soon as 30 min after GSNO treatment in a plate reader, using wavelengths corresponding to fluorescein (excitation 483 ± 14 nm and emission at 530 ± 30 nm) (Figure 1).
Figure 1. Example of nitric oxide (NO) quantification after S-nitrosoglutathione (GSNO) treatment. DAF-FM fluorescence was measured every 15 min for 4 h in a sample treated with GSNO (1 mM) and in the untreated control. Values represent the average of four biological replicates; error bars indicate ± SEM.
DNA degradation profile analysis
Eight hours after the beginning of the GSNO treatment, transfer the contents of two wells (2 mL) of the 24-well plate into a 2 mL Eppendorf tube.
Harvest the cells by centrifuging the tubes at 2,300× g for 5 min.
Remove the supernatant and dissolve the pellet in 600 µL of extraction buffer by placing it for 10 min at 37 °C under 1,400 rpm agitation in a thermomixer.
Centrifuge at 17,000× g for 3 min to pellet the debris.
Transfer 400 µL of the supernatant into a new 1.5 mL Eppendorf tube.
Add 500 µL of phenol:chloroform:isoamyl solution.
Mix the solution by vortexing for at least 30 s.
Centrifuge at 16,000× g for 5 min.
Carefully collect 300 µL of the upper phase and transfer it to a new 1.5 mL Eppendorf tube.
Add 750 µL of 100% ethanol and 45 µL of sodium acetate 3.3 M, mix gently turning the tube over a few times, and leave in ice for 30 min.
Centrifuge at 17,000× g for 30 min at 4 °C to pellet the DNA.
Remove all supernatant and rinse the pellet with 500 µL of ethanol 70%. Centrifuge at 17,000× g for 5 min at 4 °C.
Remove all supernatant and dry the pellet by placing the tube upside down on absorbent paper for 10 min.
Note: No residues of ethanol should remain at this stage. If needed, the tubes can be dried for a longer time.
Add 100 µL of water and dissolve the pellet by shaking at 1,400 rpm for 5 min at 50 °C in a thermomixer.
Note: The pellet can be hard to dissolve. If needed, use a pipette tip and increase agitation time.
DNA concentration (typically 200–500 ng/µL) and A260nm/A280nm ratio (expressing protein contamination, typically between 1.8 and 2) are estimated spectrophotometrically using a NanoDrop.
Digest 10 µg of DNA with 1 µL of RNase A for 15 min at 37 °C.
Load 10 µg of your DNA samples and 4.5 µL of GeneRuler 100 bp Plus DNA ladder on a 2% agarose gel.
Check the DNA degradation profile after 20 min of migration at 100V and estimate the size of the DNA fragments, using GelDoc Go and Image Lab (Figure 2).
Figure 2. Analysis of DNA degradation profile in untreated cells compared with cells treated with S-nitrosoglutathione (GSNO) 0.5 mM and 1 mM, 8 h after treatment. For each condition, two independent samples are shown. The size of the marker bands is expressed in base pairs.
Death quantification procedure
At different times after GSNO treatment (we recommend 4, 8, and 24 h), take 20 µL of culture from each well and add 8 µL of Evans blue 0.5%.
Observe the cells under a microscope (at 200× or 400× magnification) using a slide with sufficient space to avoid crushing the cells (e.g., Counting chamber Neubauer) and take a representative photo of each sample.
Determine the percentage of dead cells over a population of at least 100 cells by counting the number of blue (dead cells) and living cells (green cells) (Figure 3A). The percentage of dead cells is equal to the number of dead cells divided by the total number of cells, multiplied by 100 (Figure 3B).
Figure 3. Cell death quantification using Evans blue staining. A. Living cells remain green, while dead cells appear blue after treatment with Evans blue. Scale bar represents 50 µm. B. Percentage of dead cells at different times after S-nitrosoglutathione (GSNO) treatment compared with control. Values represent the average of six biological replicates; error bars indicate ± SEM. Adapted from Lambert et al. [12].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Lambert et al. [12]. Type II metacaspase mediates light-dependent programmed cell death in Chlamydomonas reinhardtii. Plant Physiology (Figures 1, 2–6).
In this article, the percentage of death was assessed in different Chlamydomonas populations, using between four and six biological replicates and a Student's t-test to highlight significant differences. In this way, we were able to reveal quite fine statistical differences between the different samples tested (e.g., Figure 1A).
Acknowledgments
This work was supported in part by the CNRS (MITI, ADAPT-VIVANT), Sorbonne Université (iBio initiative). We would like to thank the authors of the publication from which this Bio-protocol article was inspired: Lambert, L., de Carpentier, F., André, P., Marchand, C. H. and Danon, A. (2024). Type II metacaspase mediates light-dependent programmed cell death in Chlamydomonas reinhardtii. Plant Physiol. 194(4): 2648–2662 [12].
Competing interests
The authors declare no conflicts of interest.
References
Kerr, J. F., Wyllie, A. H. and Currie, A. R. (1972). Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer. 26(4): 239–257.
de Carpentier, F., Lemaire, S. D. and Danon, A. (2019). When Unity Is Strength: The Strategies Used by Chlamydomonas to Survive Environmental Stresses. Cells. 8(11): 1307.
Lambert, L., de Carpentier, F., André, P., Marchand, C. H. and Danon, A. (2024). Type II metacaspase mediates light-dependent programmed cell death in Chlamydomonas reinhardtii. Plant Physiol. 194(4): 2648–2662.
Chang, H. L., Hsu, Y. T., Kang, C. Y. and Lee, T. M. (2013). Nitric Oxide Down-Regulation of Carotenoid Synthesis and PSII Activity in Relation to Very High Light-Induced Singlet Oxygen Production and Oxidative Stress in Chlamydomonas reinhardtii. Plant Cell Physiol. 54(8): 1296–1315.
Yordanova, Z. P., Woltering, E. J., Kapchina-Toteva, V. M. and Iakimova, E. T. (2013). Mastoparan-induced programmed cell death in the unicellular alga Chlamydomonas reinhardtii. Ann Bot. 111(2): 191–205.
Danon, A., Delorme, V., Mailhac, N. and Gallois, P. (2000). Plant programmed cell death: A common way to die. Plant Physiol Biochem. 38(9): 647–655.
Vavilala, S. L., Gawde, K. K., Sinha, M. and D’Souza, J. S. (2015). Programmed cell death is induced by hydrogen peroxide but not by excessive ionic stress of sodium chloride in the unicellular green alga Chlamydomonas reinhardtii. Eur J Phycol. 50(4): 422–438.
Schnell, R. A. and Lefebvre, P. A. (1993). Isolation of the Chlamydomonas regulatory gene NIT2 by transposon tagging. Genetics. 134(3): 737–747.
Li, X., Patena, W., Fauser, F., Jinkerson, R. E., Saroussi, S., Meyer, M. T., Ivanova, N., Robertson, J. M., Yue, R., Zhang, R., et al. (2019). A genome-wide algal mutant library and functional screen identifies genes required for eukaryotic photosynthesis. Nat Genet. 51(4): 627–635.
Gorman, D. S. and Levine, R. P. (1965). Cytochrome f and plastocyanin: their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardi. Proc Natl Acad Sci USA. 54(6): 1665–1669.
Broniowska, K. A., Diers, A. R. and Hogg, N. (2013). S-Nitrosoglutathione. Biochim Biophys Acta BBA - Gen Subj. 1830(5): 3173–3181.
Article Information
Publication history
Received: May 13, 2024
Accepted: Jul 2, 2024
Available online: Jul 18, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Plant Science > Plant physiology > Abiotic stress
Cell Biology > Cell signaling > Stress response
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5,044 | https://bio-protocol.org/en/bpdetail?id=5044&type=0 | # Bio-Protocol Content
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Peer-reviewed
Construction of ThermoMaze
AR Aryeh Rothstein
MV Mihály Vöröslakos
YZ Yunchang Zhang
KM Kathryn McClain
RH Roman Huszár
GB György Buzsáki
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5044 Views: 423
Reviewed by: Oneil Girish BhalalaRaquel Tonello Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in eLIFE Nov 2023
Abstract
Physiological changes during awake immobility–related brain states remain one of the great unexplored behavioral states. Controlling periods of awake immobility is challenging because restraining the animal is stressful and is accompanied by altered physiological states. Here, we describe the ThermoMaze, a behavioral paradigm that allows for the collection of large amounts of physiological data while the animal rests at distinct experimenter-determined locations. We found that the paradigm generated long periods of immobility and did not alter the brain temperature. We combined the ThermoMaze with electrophysiology recordings in the CA1 region of the hippocampus and found a location-specific distribution of sharp-wave ripple events. We describe the construction of the ThermoMaze with the intention that it helps enable large-scale data recordings on immobility-related brain states.
Key features
• Controlling periods of awake immobility in rodents.
• Electronic-friendly analog of the Morris water maze.
Keywords: Rodent behavior Electrophysiology Thermoregulation Sharp-wave ripples
Background
Neural oscillations operate as the language of the brain, underlying neuronal processes such as cognitive capabilities [1]. Recent advances (e.g., optogenetics) have enabled researchers to explore in detail the anatomy and functioning of the brain irrespective of behavior [2]. However, utilization of behavioral paradigms is critical for understanding the relationship between behavioral states and ongoing brain activity [3]. In basic neuroscience studies, behavioral paradigms should utilize natural stimuli and investigate behaviors that are part of the animal’s ethogram [4].
One ubiquitous rodent behavioral task is the Morris water maze (MWM), in which a rodent (generally a rat or mouse) needs to swim through a pool to find a platform to stand on [5]. The periods of immobility on the platform provide important neural information, as periods of wakeful rest (similar to periods of immobility) have been linked to initial consolidation processes with increased replay in humans [6]. Despite this and its abundant use, the MWM is a stressful environment for both mice [7] and rats [8] and, as such, does not serve as a natural environment for these rodents. Furthermore, the aqueous environment of the maze makes electrical recordings difficult, if not impossible, hindering the ability of scientists to dissect occurring physiological changes.
Here, we describe the construction of an electronic-friendly analog of the MWM, the ThermoMaze. The paradigm was validated via electrophysiological measures, but its use extends any electronics-based data collection modality, including but not limited to freely moving 2-photon imaging, wide-field calcium imaging, and fiber photometry. Utilizing an abrasively cold environment with specific hotspots, the paradigm capitalizes on thermotaxis and thermoregulatory behaviors of mice to generate long periods of immobility during the behavioral task, enabling a deeper and richer understanding of the immediate oscillatory changes underlying memory.
Materials and reagents
Note: Information on vendors, model numbers, and links to purchase equipment is provided when possible. The list of materials used here can also be accessed at https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/BOM_ThermoMaze_v03.txt
Peltier elements (Amazon, catalog number: TEC1-12706, https://www.amazon.com/GeeBat-TEC1-12706-Thermoelectric-Heatsink-Cooling/dp/B01IT8SAZG/)
Aluminum water cooling block (Amazon, catalog number: B08JKP6HYC, https://www.amazon.com/uxcell-Aluminum-Heatsink-Computer-Graphics/dp/B08JKP6HYC/)
Submersible water pump (Amazon, catalog number: B085NQ5VVJ, https://www.amazon.com/LEDGLE-Submersible-Ultra-Quiet-Dual-Purpose-Hydroponics/dp/B085NQ5VVJ/)
UV-resistant cast acrylic 1/8” thickness, white (McMaster, catalog number: 8505K743, https://www.mcmaster.com/8505K743)
Clear soft PVC plastic tubing for air and water, 5/16" ID, 7/16" OD (McMaster, catalog number: 5233K59, https://www.mcmaster.com/5233K59/)
Thermally conductive epoxy (MG Chemicals, catalog number: 8349TFM-25ML, https://www.amazon.com/MG-Chemicals-8349TFM-Thermally-Condcutive/dp/B08Z73HH23/)
Wood epoxy
3D-printed frame (https://github.com/misiVoroslakos/3D_printed_designs/tree/main/ThermoMaze)
K-type thermocouple (OMEGA, catalog number: 5TC-TT-K-40-36, https://www.newark.com/omega/5tc-tt-k-40-36/thermocouple-wire-type-k-40awg/dp/30AC8682)
Thermistor (Mouser, catalog number: 954-223FU3122-07U015, https://www.mouser.com/ProductDetail/Semitec/223Fu3122-07U015/?qs=raqtESnDWsAF5g197HBRcQ%3D%3D)
Crushed ice
Refreezable ice packs
Bucket for water
FLIR C5 compact thermal imaging camera (Mouser, catalog number: 685-89401-0202, https://www.mouser.com/ProductDetail/Teledyne-FLIR/89401-0202?qs=vmHwEFxEFR8KCgYmUURHLQ%3D%3D)
Equipment
Video camera (Basler, model: ace 2 a2A2590-60ucBAS)
Lens (Basler, 16 mm, model: C23-1616-2M)
GPIO cable (Basler, model: GP-I/O Cable 6p/open, 10 meters)
Tripod mount (Basler, catalog number: 2200000314)
E36102A benchtop current generator (Keysight Technologies, catalog number: E36102A/0E3/902, https://www.mouser.com/ProductDetail/Keysight/E36102A-0E3-902?qs=YCa%2FAAYMW03ip5pRT6eiig%3D%3D)
Microcontroller circuit board (Arduino, model: Mega 2560 Rev3, https://store-usa.arduino.cc/collections/boards-modules/products/arduino-mega-2560-rev3?_pos=2&_fid=04e83805a&_ss=c)
8-Channel relay module (Amazon, catalog number: 101-70-102, https://www.amazon.com/SainSmart-101-70-102-8-Channel-Relay-Module/dp/B0057OC5WK/ref=asc_df_B0057OC5WK/?tag=hyprod-20&linkCode=df0&hvadid=312070784062&hvpos=&hvnetw=g&hvrand=14266163912783471503&hvpone=&hvptwo=&hvqmt=&hvdev=c&hvdvcmdl=&hvlocint=&hvlocphy=9067609&hvtargid=pla-348881239402&th=1)
Microcontroller circuit board (Arduino, model: Uno)
Red LED (Amazon, catalog number: ED_P05_R_100Pcs, https://www.amazon.com/EDGELEC-100pcs-Resistors-Included-Emitting/dp/B077XDYTTP)
Resistor (470 Ohm, catalog number: ED_P05_R_100Pcs, https://www.amazon.com/EDGELEC-100pcs-Resistors-Included-Emitting/dp/B077XDYTTP)
Cables for Arduino (ELEGOO, model: EL-CP-004, https://www.amazon.com/Elegoo-EL-CP-004-Multicolored-Breadboard-arduino/dp/B01EV70C78)
Banana plugs with micro grabbers (Pomona, catalog number: EM5053-12-0#)
Handheld thermometer (Omega, model: HH800, https://www.omega.com/en-us/test-inspection/handheld-meters/temperature-and-humidity-and-dew-point-meters/p/HH800)
Software and datasets
Arduino IDE (version 1.8.19, open source). This was used to control the submerged DC pumps and to run the behavioral paradigm
Intan RHD2000 USB Interface Board software was used to test the electrical collections and collect data
Pylon Viewer (Basler AG, Ahrenberg, Germany; version 6.2.0, available at https://www2.baslerweb.com/en/downloads/software-downloads/#version=6.2.0) was used to control the video camera from the computer.
Windows built-in video recorder was used to record infrared (IR) video from the FLIR camera
MATLAB (version R2021A) was used to analyze data using custom scripts
Key links:
Important information on using 3D printing files can be found at: https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/README.md
The Arduino code for running the behavioral paradigm can be found here: https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/ThermoMaze_control/Cooling_box_v04_relays_20220410.ino
The file for 3D printing the floor can be found at: https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/3D_printed_design/cooling_box_v03_platform.stl
Procedure
Calibrating Peltier elements and determining driving voltage
To determine the ideal driving voltage, grab a single Peltier device, the power supply of the Peltier, two banana plugs with micro grabbers, a thermocouple, and a handheld thermometer.
The Peltier device temperature gradients are chronically stable; so, an assessment of the driving voltage at the experiment's outset would enable consistent temperature drops [9]. The manufacturer recommends using a constant current application to drive the Peltier devices; we adjusted voltage as our power source could generate the necessary constant current based on our constant voltage. We found that the constant current was 0.975A.
Critical: For power sources that cannot generate a current of that magnitude, it is prudent to check for and utilize a constant driving current and not voltage.
Connect the banana plug with the micro grabber and turn on the power supply of the Peltier.
Ensure that the experimenter can manipulate the voltage and that the intensity of the current will automatically be adjusted. If your power source does not have the capability to generate a sufficiently high current based on the voltage set, use variable currents and run the following steps adjusting current instead of adjusting voltage.
Set the voltage to 1 V.
Rest the thermocouple on the Peltier device.
Record the initial temperature using the handheld thermometer. The readout should indicate a value around 25 °C (i.e., room temperature).
Connect the micro grabber tips to the cables on the Peltier device.
Use the power supply to drive a current across the Peltier device for a couple of seconds and record the lowest temperature the Peltier achieves.
Remove the micro grabbers from the Peltier cables (breaking the circuit/disconnecting the Peltier element from the power supply), let the Peltier element warm up, and record the highest temperature achieved.
Once back at room temperature, increase the voltage by 0.2 V.
Repeat steps A7–A11 until you identify the target temperature for the study.
We found that stimulating between 2.4 and 3.4V without any heat removal led to our ideal hotspot temperature of 25 °C (Table 1).
Note: Peltier elements have a built-in temperature gradient (∆T), and so, by switching polarities, the Peltier element will switch from heating to cooling. We found passive cooling via flowing ice water was sufficient to drop the floor temperature to 10 °C; so, we only needed to stimulate the Peltier element to form the hotspot. We identified that ∆T = 10 °C for the Peltier; by flipping the polarity when stimulating, we generated the 35 °C hotspot.
Caution: Each Peltier element has its own ∆T, so it is imperative to check the specifications and run this type of trial when starting a project with these devices.
Table 1. Trial data for determining driving voltage.
Each trial involved measuring the room temperature, the coldest temperature, and the temperature 1 min after the current was stopped (all temperature values measured in °C). Current (in amperes; A) was autogenerated by the power source of the Peltier based on the input voltage (measured in volts; V) of the current generator. Note: Due to the built-in temperature gradient in the Peltier elements, the difference in temperature is what we focused on, and so it is equally effective to investigate the highest temperature recorded depending on how the element is stimulated.
Voltage (V) Current (A) Initial temperature (°C) Coldest temperature (°C) Stable temperature after 1 min (°C)
1.0V 0.300A 22.1°C 19.0°C 22.7°C
1.2V 0.350A 22.1°C 17.9°C 22.6°C
1.4V 0.405A 22.0°C 17.3°C 23.0°C
1.6V 0.480A 22.5°C 17.0°C 23.1°C
1.8V 0.535A 22.2°C 16.3°C 23.4°C
2.0V 0.600A 22.2°C 15.5°C 23.2°C
2.2V 0.650A 22.2°C 15.5°C 23.4°C
2.4V 0.700A 22.4°C 15.3°C 24.5°C
2.6V 0.775A 22.2°C 14.8°C 24.5°C
2.8V 0.820A 22.5°C 14.5°C 24.9°C
3.0V 0.890A 22.5°C 14.3°C 25.2°C
3.2V 0.950A 22.1°C 13.1°C 24.2°C
3.4V 1.000A 22.0°C 12.7°C 24.8°C
3.6V 1.080A 22.2°C 12.4°C 25.4°C
3.8V 1.130A 22.4°C 12.2°C 26.0°C
4.0V 1.200A 22.2°C 12.1°C 25.3°C
4.2V 1.250A 22.4°C 12.0°C 26.5°C
4.4V 1.300A 22.7°C 12.1°C 26.7°C
Once the trial is complete (i.e., once the driving voltage is identified), proceed to connect each Peltier element to a cooling block.
Constructing the floor of the maze
Take a Peltier device and connect it to the 3D floor printout using hot glue.
Once the glue has hardened, place wood epoxy to fill in any gaps between the Peltier and frame.
After the Peltier is secured to the frame, flip the frame over and ensure that the semi-circle cutouts in the frame (see Figure 1B) align with the locations of the cables on the Peltier.
Critical: The Peltier elements will be connected to the power source via banana plugs. It is critical to ensure proper orientation with the cutouts in the frame (Figure 1B) so that all banana plugs will connect to the Peltier while maintaining the levelness of the floor. Failure to properly align the plugs and the cutouts will cause the Peltier to partially pop up out of the frame, creating an unlevel floor.
Once the alignment above is achieved, take one cooling block and orient it such that the flat portion faces the Peltier element.
Place the silver (heat-resistant) epoxy on the flat portion of the cooling block.
Rest the Peltier element on the epoxy until the epoxy hardens; then, cement the Peltier element to the cooling block.
Repeat this process for each of the 25 Peltier devices.
Connecting the floor to the water tank
Once all of the Peltier elements are connected to the cooling blocks and placed in the floor printout, begin working on connecting the water tank to the cooling blocks.
Take the end of a piece of PVC tubing and begin by cutting the pipe into small segments.
Connect the end of one piece of pipe to a submerging DC pump and the other end to the inflow nozzle of element 1.
Note: The number of the Peltier element used is based on proximity to the water tank, with 1 being the element closest and 5 the element farthest from the water tank.
After securing the tubing, grab another piece of pipe and secure one end to the outflow nozzle of 1 to the inflow nozzle of 4.
Using separate pieces of pipe, connect the outflow of 4 to the inflow of 2, the outflow of 2 to the inflow of 5, and the outflow of 5 to the inflow of 3.
Figure 1. Schematic of the paradigm setup. A) Overview of the maze, video, and IR cameras, LED, and water tank. The dashed box demonstrates the composition of each Peltier element once on the maze floor. B) Connections between the Arduino, relay switches, and Peltier elements. Arrows point to examples of the semicircles cut out of the floor printout to create space for the Peltier’s cables. Red dot: cathode; Blue dot: anode. C) Connections between the water tank and Peltier elements. Yellow squares: water pumps (one pump for each five elements). Dashed lines show the direction of water flow. Reprinted from Vöröslakos et al. [10] with permission.
Finally, take a piece of pipe and connect one end to the outflow of 3 and place the other end into the water tank (see Figure 2D).
Repeat this process for the remaining cooling block elements and submerging DC pumps, ensuring that a single pump circulates the water through one row of Peltier elements.
Critical: After completing all connections between the water cooler and the five Peltier elements, check that the water temperature arriving at each Peltier element is kept cool at the temperature of the water tank. There should be no significant difference—less than 1 °C—between the temperature in the tank water and the water cooling each of the five Peltier elements.
Note: Our analysis demonstrated that each of the five Peltier elements was efficiently cooled to the tank water temperature.
Once connections between the floor and water tank are complete, proceed to construct the final part of the ThermoMaze: the walls of the maze.
Figure 2. Overview of the connections to run the paradigm. A) Schematic showing the connections between the Arduino mega, the relay module, the hotspots in the ThermoMaze, and the power source of the Peltier elements. B) Photograph of ThermoMaze with all Peltier elements attached to a 3D-printed frame (bottom view). One row of water coolers (n = 5) is also attached to Peltier elements. C) Photograph of the bottom view of the ThermoMaze showing 25 water coolers without tubing attached. D) Demonstration of how the Peltier elements in one row are connected so the water circulates through all the elements. Peltier element 1 is closest to the pump and 5 is closest to the thermometer when determining the tubing connections between Peltier elements (steps C3–6). This example demonstrates that passive cooling suffices to drop the surface temperature of the Peltier elements to the desired value, as the ice-cold water flowing through the tubes and between the water coolers and Peltier elements (turned off) passively reduced the surface temperature of the Peltier element to 0.8 °C. The temperature is measured by a K-type thermocouple attached to the surface of the last Peltier element in a row. Figure is reprinted from Vöröslakos et al. [10] with permission.
Constructing the box
To construct the walls, take the cast acrylic and cut out four identical walls measuring 20 cm × 20 cm × 40 cm (width × length × height).
Using hot glue, secure the first wall to the floor.
Note: Hot glue was used so that the walls could be angled outward, providing a large opening on top for the camera to easily visualize the entirety of the maze.
After securing that wall in place, take the second wall and secure both to the floor of the maze and to the first wall using hot glue.
Repeat this process with the remaining two walls to complete the construction of the ThermoMaze box.
Connecting the Peltier elements to the power supply of the Peltier
Open the Arduino code for the behavioral paradigm.
Connect the Arduino mega board to the computer and transfer the behavioral code to the board.
Connect the Arduino board to a relay module.
Connect each of the four Peltier elements that will serve as hotspots to each of the four relay elements in the module.
Connect the positive side of the variable voltage source to each of the four relay modules, and the negative side of the variable voltage source to each of the four Peltier elements that will be hotspots.
Note: To enable the single port (on the variable voltage source) to connect the four other parts (either the relay spots or the hotspots), stack banana plugs so that all four plugs can access a single spot.
Hotspot customization
All aspects of the hotspots (location, order, duration) are customizable and should be modified to the experiment's needs.
Each Peltier element is individually connected to the relay module, allowing any of the 25 Peltier elements to serve as a hotspot, and providing the experimenter with the option to have anywhere from 1 to 25 hotspots. To adjust the hotspot location, alter the connections between the relay module and Peltier elements such that the code will activate the newly chosen hotspots (in Figure 2A, moving the colored connections to different elements will shift the hotspots to the new locations).
To adjust the order of the hotspots, edit the Arduino code (https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/ThermoMaze_control/Cooling_box_v04_relays_20220410.ino) and reorder when each relay and TTL signal are high (see lines 4–16 and 69–120 of the code).
Increasing the duration of each hotspot allows for sleep at said spot. We found that turning hotspots on for periods of 20 min was sufficient for triggering sleep [10]. To alter the duration of the hotspot, change the number in line 27 of the above code (can be compared to https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/ThermoMaze_control/Cooling_box_v04_relays_20220410_sleep.ino).
Final touches in constructing the paradigm
Tape an Arduino-compatible LED roughly halfway up one of the walls of the maze (Figure 1A). In our construction, the LED was taped approximately 20 cm up the wall and roughly centered on the wall.
Critical: Ensure the LED is high enough on the wall that the animal cannot reach it.
Notes:
The LED serves two purposes: 1) the pulsing of the LED enables you to sync the video to the electrophysiology signals, so that should any frames drop, the LED will provide clear timing between the signals as to when the frames dropped; and 2) the LED serves as a distal cue.
In placing the LED, the coordinates do not need to be measured precisely. Rather, by eye the LED should be roughly halfway up and in the center of the wall. Ensure that the LED is blinking and visualized on both the IR and video cameras.
Secure both the IR camera and the video camera above the maze.
Calibrate the IR camera.
On the IR camera, click on settings, and select the feature that enables you to change the temperature gradient.
With the gradient adjustable, press lock on the upper bound while the lower bound remains adjustable.
Pause point: Ensure that the upper bound is locked and the lower bound is adjustable. If both bounds are adjustable, the pre-existing range will not change, but the bounds on the range will change in unison. If both are locked, then no changes will occur.
Set the lower temperature bound to a value well below the lowest expected floor temperature (for our purposes, the lower bound was set to 4 °C).
Lock the lower bound and unlock the upper bound so that it is adjustable.
Pause point: Ensure the lower bound is locked and the upper bound is adjustable. See step G3c above for further explanation.
Set the upper temperature bound to a value well above the highest expected floor temperature (for our purposes, the upper bound was set to 40 °C).
Lock the upper bound.
Critical: Ensure that both bounds are locked at the desired values before proceeding. With the bounds set on the IR camera, confirm that it is connected to the computer.
On the computer, open the Windows built-in video recorder and check that the IR camera captures the entire field.
The IR camera is now properly calibrated.
Calibrate the video camera.
To begin calibrating the video camera, ensure that the LED is turned on (i.e., blinking).
On the computer, open the Intan RHD USB board interface software.
Check that the LED pulse signal is visible.
Once you confirm the LED pulses are picked up by the Intan RHD USB board interface software, open the pylon viewer software.
Confirm that the entirety of the maze is clearly displayed on the screen.
The video camera is now properly calibrated.
Prepare the pulley system for freely moving recordings.
To enable extracellular electrophysiological recordings while the animal is in the maze, set up a pulley system.
The system should have a long, wire-thin cable with a connector to a head stage on the end.
Once the implanted probe is connected to the head stage during an experimental session, adjust the slack in the long wire-thin cable. This cable should not be too loose that it folds over itself and affects the mouse's movement, and not so tight that it pulls up on the mouse’s head.
Once the maze is constructed and you finish all connections, check for any electromagnetic interference (EMI)-induced noise.
Critical: Checks for EMI-induced noise should be conducted upon completing construction (to ensure no faulty connections).
To check for EMI noise, place an implanted animal inside the ThermoMaze and run the “Cooling” sub-session protocol.
If line noise (50/60 Hz) appears on all recording channels regardless of the location of the animal, then the ground connection of the submerging pump(s) is improper.
Note: Tap water includes impurities, such as dissolved sodium, calcium, and magnesium salts; thus, it is an excellent conductor of electricity.
If line noise (50/60 Hz) appears on all recording channels when the animal is approaching an active Peltier, then the ground connection of that Peltier is improper.
Note: Peltier effect produces a temperature difference between two sides of a Peltier device when a current is flowing. Improper grounding of Peltier device(s) can induce spatially restricted line noise.
Potential solutions to remedy any EMI-induced noise identified:
Ensure that all wires are tightly screwed into their location.
Avoid ground loops and make sure that all electronic devices (Peltier devices, submerging pumps, Arduino, relays, and recording system) share a common ground.
Note: Use a multimeter to test the continuity of the ground lines.
For any noise stemming from the submerging pumps, remove all pumps from the water tank. Place one pump inside the water tank and observe the brain signal for potential line noise. If there is no electrical noise, repeat these steps with the remaining submerging pumps.
Critical: Replace any submerging pump(s) that induces electrical noise. If noise still persists, consider using deionized pure water instead of tap water.
Check for any moveable appliances (which could be the source of the noise); remove any and all of these appliances from the recording room.
Try moving the wires from the relay system around to see if there is a position that removes the noise.
For any noise stemming from the long wire-thin cable, try to move it around. Once you find a position that keeps the signal noise-free, rest the cable on a nearby platform or clip it to a nearby surface.
Wrap some of the cables in aluminum foil, which can help to re-ground the signal. For this option, continue to test different groupings of wires until you find a place where the aluminum foil wrapped around the wires removes the EMI noise.
If none of these solutions remedy the EMI-induced noise, check all cables for signs of damage or fraying. Replace the cables and wires if necessary.
Running the paradigm (Figure 3A)
Figure 3. Overview of the paradigm. A) Five sub-sessions constitute a daily recording session: (i) rest epoch in the home cage, (ii) pre-cooling exploration epoch (Pre), (iii) cooling, (iv) post-cooling exploration epoch (Post), and (v) another rest in the home cage. B) Schematic of temperature landscape changes when the animal is in the ThermoMaze (top) and example animal trajectory (below). During cooling, one Peltier element always provides a warm spot for the animal (four Peltier elements in the four corners were used in this experiment). Each Peltier element was turned on for 5 min in a sequential order (1–2–3–4) and the sequence was repeated four times. C) Session-averaged duration of immobility (speed ≤ 2.5 cm/s) that the animal spent at each location in the ThermoMaze. Color code: temporal duration of immobility(s); white lines divide the individual Peltier elements; n = 17 sessions in 7 mice. Adapted from Vöröslakos et al. [10] with permission.
Pre-maze home cage baseline recording (2 h; Figure 3Ai)
Begin by placing the mouse (small rodent) in a home cage in the same room as the maze.
Note: The animal has access to food or water throughout the duration of the home-cage experimental session.
Connect the mouse to the recording software and check for any EMI-induced noise. Follow procedure step G7 to remedy any EMI-induced noise that should arise.
Once the signal is free of EMI-induced noise, start the recording.
To avoid any olfactory cues from affecting behavior, the experimenter should either leave or remain in the recording room for the entire 2 h.
After 2 h, stop the recording and keep the mouse in the home cage as you set up the maze.
Pre-behavior in maze baseline (Pre-cooling; 10 min; Figure 3Aii)
Fill the tank halfway with 25 °C water.
Critical: Ensure that all five submerged pumps are completely covered. If they are not completely submerged, their vibrations can generate EMI noise and serve as a noxious auditory stimulus to the rodent.
Ensure the LED light is securely taped to the wall of the maze and is blinking.
Take the mouse from its home cage and place it in the maze, ensuring that the rodent either remains connected to or is reconnected to the recording software once in the maze. If the mouse is anxious after placement in the maze and ensuring connections to the recording software, allow it a bit of time to calm down before proceeding to the next step.
Turn on the pumps to begin cycling the water on the floor of the maze.
Critical: Place a finger onto the suction point of each of the five submerged pumps and check that each is working properly (you should feel the suction pulling on your finger). If there are any suction issues, detach the tube from the pump (allowing water to flow through the pump) and reattach them to remedy the issue.
For those using tap water, ensure there is no EMI-induced noise in the signal. If noise is present in the signal, it is likely due to one of the pumps. Remedy the issue by gently resting the problematic pump on a neighboring tube (light enough that it does not obstruct flow) to remove the effect of the pump vibrating against the floor of the tank. Using distilled water is unlikely to cause any issues as it is free of conductive ions.
Critical: Start the electrophysiological recording, then the IR and regular video recordings. The electrophysiological recording must be started prior to the video recordings to ensure that all video frames are timestamped to a data point in the electrical recording, so that if there are any issues with the video recordings (e.g., dropped frames), their points can be identified on the electrical data.
After 10 min, stop the IR and regular video recording and then the electrophysiological recording.
Critical: Stop the video recordings while still collecting electrophysiological data (see reasoning in step H2h).
Once all recordings are stopped, turn off the pumps.
Run the paradigm (Cooling; 80 min; Figure 3Aiii, Video 1)
With the pumps stopped (and the water no longer circulating), fill the tank with crushed ice to lower the temperature to 10 °C using the handheld thermometer to continuously monitor the temperature. Make sure to stir the ice around the tank before checking temperature to ensure the water temperature is uniform.
Once at 10 °C, place two reusable ice packs (“ice blocks”) to stabilize the temperature at 10 °C.
Note: The ice blocks will keep the temperature stabilized for the duration of the cooling portion of the experiment (80 min) in standard laboratory conditions (room temperature kept at 21 ).
Once the water temperature is stable, ensure the LED is still blinking, check for any EMI-induced noise, and remedy any issues (see steps F6 and F7).
Critical: Start the electrophysiological recording and, subsequently, begin the IR and regular video recording. See step G2h for the importance of starting electrical recordings prior to video recordings.
Once the electrophysiological and video recordings have begun, ensure the driving voltage is properly set on the Peltier driving power supply, begin circulating the water, and start the behavioral paradigm software protocol.
Critical: Ensure that the pumps and the behavioral paradigm are working properly. To check the pumps and solve any issues, see step G2f. To ensure the paradigm is working properly, pull up the Windows built-in video recording software on the computer. The entire floor should be at the temperature of the circulating water, except for one hotspot, which should be at 35 °C. Also, check that the animal finds and remains immobile on the hotspot.
As the paradigm is running, the experimenter should remain in the room and sporadically check all of the maze parameters (i.e., check for any EMI-induced noise, water temperature, and IR video).
If any EMI-induced noise is identified, follow step F7 to resolve the issue.
If the water temperature exceeds 13.5 °C, add some more crushed ice in small increments, stirring the water (with your finger) and then checking the temperature after each increment. Do not let the water temperature fall below 9 °C.
When checking the video, ensure the floor is at the water temperature (10 °C) except for the hotspot (which should be at 35 °C), that the animal is finding and remaining immobile on the hotspot, and that you see the hotspot switch locations.
Video 1. Mouse engaging in the ThermoMaze behavioral task. Adapted from Vöröslakos et al. [10] with permission.
Post-behavior in-maze baseline (Post-cooling; 10 min; Figure 3Aiv)
After 80 min, the behavioral paradigm should be complete. Immediately stop circulating the water (to help the floor start to warm up for the animal), then stop the IR and regular video recordings, and finally stop the electrophysiology recording (see Procedure G2h for the importance of stopping order).
Once all of the recordings are stopped, remove the ice blocks from the tank and pour out some of the tank water into a bucket (making room for adding warmer water).
Add warm/hot water to the tank to raise the water temperature back to 25 °C.
Once the water is at 25 °C, immediately begin circulating the water to make the rodent more comfortable.
Once the pumps are turned on, follow steps H2h–k.
Post-maze home cage baseline recording (2 h; Figure 3Av)
With all of the recordings stopped, take the rodent out of the maze and place it back in the home cage where the pre-maze home cage baseline recording occurred.
Note: The animal has access to food and water during the duration of the home-cage experimental session.
Connect the mouse to the recording software and check for any EMI-induced noise. Follow step F7 to remedy any EMI-induced noise that should arise.
Once the signal is free of EMI-induced noise, start the recording.
To avoid any olfactory cues from affecting behavior, the experimenter should either leave or remain in the recording room for the entire 2 h.
After 2 h of recording, the experimental session is now complete.
Data analysis
Detailed description of data analysis can be found in the Quantification and statistical analysis section of Vöröslakos et al. [10]. To perform the analysis, expertise using MATLAB is necessary.
Validation of protocol
The preparations and data analysis are exactly as described previously [10] and are reprinted below for convenience. All experiments were approved by the Institutional Animal Care and Use Committee at New York University Langone Medical Center. Animals were handled daily and accommodated to the experimenter and the ThermoMaze before surgery and electrophysiological recordings. Mice (adult female n = 8, mean = 22 g; male n = 5, mean = 26 g) were kept in a vivarium on a 12/12 h light/dark cycle and housed two per cage before surgery and individually after it. Atropine (0.05 mg/kg, s.c.) was administered after isoflurane anesthesia induction to reduce saliva production. Body temperature was kept between 36 and 37 °C via a temperature controller (TCAT-LV; Physitemp, Clifton, NJ). Stages of anesthesia were maintained by confirming the lack of a nociceptive reflex. The skin of the head was shaved, and the surface of the skull was cleaned by hydrogen peroxide (2%). A custom 3D-printed baseplate ([11]; Form2 printer, FormLabs, Sommerville, MA) was attached to the skull using C&B Metabond dental cement (Parkell, Edgewood, NY). The craniotomy site was marked and a stainless-steel ground screw was placed above the cerebellum. Silicon probe attached to a metal Microdrive [12] was implanted into the dorsal CA1 of the hippocampus (Bregma: -2 mm AP, -1.5 mm ML). For surgeries testing brain temperature changes, tungsten wires were implanted in place of the silicone probe, and a thermistor was placed in the contralateral dorsal CA1 (Bregma: -2 mm AP, +1.5 mm ML; Figure 4A). A protective copper mesh cap was built around the probe. Animals received ketoprofen (5.2 mg/kg, s.c.) at the end of the surgery and on each of the following two days and were given at least five days to recover prior to experiments. The electrophysiology data was digitized at 20,000 samples/s using an RHD2000 recording system (Intan technologies, Los Angeles, CA). All data analyses occurred using custom codes in MATLAB.
Hippocampal temperature unaffected
The next concern to address is validating the paradigm-assessed brain temperature. Homeostatic thermoregulation is critical to ensure proper physiological functioning. Mice brain temperatures operate within a 4 °C range (35.5–39.5 °C), with fluctuations arising based on behavioral state [13]. Temperatures below this range can affect the time course of action potentials and neurotransmitter release [14].
Rodents rely on environmental temperature for thermoregulation. Since this paradigm capitalizes on rodent thermotaxis and thermoregulatory mechanisms, we needed to ensure that the experimenter-generated abrasive environmental temperature (10 °C on non-hotspots) did not cause temperature deviations outside of the physiologic range. To assuage this concern, we implanted mice (n = 2) with thermistors at the same coordinates contralaterally to the recording probes (see Figure 4A) and ran the paradigm collecting information on brain temperature. Tungsten wires were placed in place of silicon probes to monitor electrical activity. The hippocampal temperature remained within the thermoregulatory zone, albeit with a shift toward the zone’s upper bounds. This finding confirms and reaffirms the dependency of brain temperature on behavioral state [13,15,16] and suggests tightly regulated mechanisms for homeostatic temperature maintenance, which are independent of the ambient temperature [10].
Figure 4. Effects of environmental temperature on brain temperature. A) Schematic of the thermistor and tungsten wire implantation for trials comparing brain temperature changes during cooling and room temperature sessions. B) Probability mass function of brain temperature during cooling (blue) and room temperature (orange) sub-sessions. The data came from 10 behavioral trials using two mice. The graph demonstrates that in both cooling and ambient temperature conditions, the hippocampal temperature stayed within a physiological range. C) Median brain temperature during both cooling (blue) and ambient (orange) conditions. There is no significant difference in brain temperature when cooling vs. ambient temperature conditions (Kolmogorov-Smirnov test). D) Linear regression demonstrating a lack of correlation between brain temperature and environmental temperature (R = 0.03, p = 0.384). Adapted from Vöröslakos et al. [10] with permission.
Paradigm generates rapid and uniform temperature changes
Initial validation of the protocol began with confirmation that the code for generating hotspots (https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/ThermoMaze_control/Cooling_box_v04_relays_20220410.ino) led to rapid and uniform temperature changes in the floor. To accomplish this, we ran an emulated behavioral session without an animal. A thermocouple was placed in each of the four hotspots, the water was cooled to 10 °C (experimental conditions), and the behavioral portion of the paradigm was run (step H3). We found that cooling of the floor was uniform, and the transition between heating spots occurred within 30 s (i.e., after each 5 min period, there were 30 s as the prior hotspot cooled down and the new hotspot warmed up to 35 °C; Figure 5).
Figure 5. Temperature changes during the animal-free emulated behavioral sessions. A) Temperature changes in each hotspot measured via thermocouples. Each color corresponds to a Peltier element. Results demonstrate the floor temperature dropped to the water tank temperature within 15 s of the pumps turning on, except for the hotspot, which increased to 35 °C. The transition in hotspot location (both the heating of the new and cooling of the old hotspots) occurred within 30 s. B) Same as A), but temperatures were measured via IR camera. Adapted from Vöröslakos et al. [10] with permission.
Electrophysiological and behavioral changes during cooling
Initial investigations revolved around assessing the effects of the maze in shaping behavior, learning that the paradigm posed little issue to the mice, as it took only a few practice sessions before they readily found the hotspot (median = 3 practice sessions). Subsequently, we looked at how the cooling shaped behavior. As theorized, the cold environment promoted immobility (Figure 6), as the mice spent a smaller proportion of time moving during cooling compared to pre- and post-cooling (cooling: 23 ± 12%, pre-cooling: 40 ± 19%, post-cooling: 34 ± 16%, mean ± SD, movement defined as speed > 2.5 cm/s). Conversely, the mice spent a greater percentage of time immobile during cooling compared to pre- and post-cooling (cooling: 76.74 ± 12.41%, pre-cooling: 59 ± 19%, post-cooling: 66 ± 16%, mean ± SD, immobility defined as speed < 2.5 cm/s).
Changing hotspot locations triggered exploration (Figure 6E–H). Video 1 shows the thermotaxis behavior of a mouse in the ThermoMaze (infra-red image is overlaid on the raw video, the second half of the video is 10 times faster than real time). The mice hastily transitioned from immobility to movement (median = 12.28 s, n = 20 sessions across 7 mice; transition is defined as the time taken from the hotspot shutting off for the mice to increase their speed from 0 to 2.5 cm/s), promptly abandoning their location (median = 12.99 s, n = 20 sessions across 7 mice), and finding the new hotspot (median = 23.45 s, n = 20 sessions across 7 mice).
Neuronal firing during sharp wave ripples demonstrates place-selectivity at experimenter-designated locations
Brain states underlying behaviors can dichotomously split into “preparative” or “consummatory” [17], where the preparative class relates to ongoing appetitive behaviors that culminate in consummatory behavior. Consummatory behaviors are sometimes termed non-voluntary or non-conscious states and can be identified via electrophysiological monitoring of various brain states [18]. Of particular note, sharp wave ripples (SPW-Rs) in the hippocampus are a hallmark of consummatory behaviors [19]. SPW-Rs are critical to spatial memory processes [20–23]. They are hypothesized to participate in sleep consolidation processes by transmitting spiking information from the hippocampus to the neocortex as temporally compressed packets of the spiking that occurred while awake [24]. SPW-Rs most frequently fire during non-rapid eye movement (NREM) sleep or periods of awake immobility, both of which characterize the major behavioral state of the mice during cooling. Given that SPW-Rs readily appear during NREM sleep and awake immobility and that they are critical to consolidating spatial information, our electrophysiological assessment focused on neuronal firing within SPW-Rs. We found that pyramidal (excitatory) neurons demonstrated greater place-specific firing compared to interneurons (inhibitory), and both classes showed a higher place-specific firing compared to controls (one-sided Wilcoxon rank sum test, p < 0.001; Figure 7).
Figure 6. Cooling generated periods of immobility in the hotspots. A) Comparison between the pre-cooling, cooling, and post-cooling of the proportion of time mice spent in any of the corners (hotspots) of the maze. Median, Kruskal–Wallis test: H = 19.69, d.f. = 2, p = 5.29 × 10-5. The proportion of time spent in any corner during cooling was significantly greater than during either pre-cooling (p = 0.0004) or post-cooling (p = 0.0003). The proportion of time spent in any corner during pre- and post-cooling did not significantly differ (p = 0.9996). Dots (females) and diamonds (males) between the boxes represent the individual sessions, and the same color represents sessions from the same animal. B) Immobility duration map of an example session in which the animal was in the ThermoMaze under 25 °C (room temperature) condition. Immobility spatial distribution demonstrates that, during ambient conditions, the SPW-R are centered around the mouse’s preferred corner. C) Same as B) but during a cooling session. Distribution demonstrates that during cooling, the immobility is focused on the hotspots. D) Immobility durations within an 80-min period of free exploration of the ThermoMaze either under room temperature or during the cooling sub-session in two groups of mice (room temperature n = 3; cooling n = 20; p = 0.49, one-sided Wilcoxon rank sum tests). E) Cumulative distribution of animal speed in the ThermoMaze during three sub-sessions from 7 mice). Median, Kruskal–Wallis test: H = 139304.10, d.f. = 2, p < 0.001. F) Animal’s distance from the previously heated Peltier element site. G) Speed of the animal centered around warm spot transitions. H) Animal’s distance from the target warm spot as a function of time (red curve: median; time 0 = onset of heating). Modified from Vöröslakos et al. [10] with permission.
Figure 7. Cooling demonstrated place-specific firing during sharp wave ripples (SPW-Rs). A) Heat map showing SPW-R counts during an example session in which the animal was in the ThermoMaze under 25 °C room temperature condition. The SPW-R spatial distribution demonstrates that, during ambient conditions, the SPW-R is centered around the mouse’s preferred corner. B) Heat map showing SPW-R counts during an example cooling session. The map demonstrates that during cooling, the SPW-R firing was restricted to the hotspots. C) Total SPW-R counts within an 80-min period of free exploration of the ThermoMaze either under room temperature or during the cooling sub-session in two groups of mice (room temperature n = 3; cooling n = 20; p = 0.62, one-sided Wilcoxon rank sum test). D) Same plots as in C) but for the degree to which their spatial distributions deviate from a uniform distribution (p = 0.04, one-sided Wilcoxon rank sum test). Modified from Vöröslakos et al. [10] with permission.
Discussion
We developed the ThermoMaze, a new small rodent behavioral paradigm that is a non-aqueous Morris water maze analog. The paradigm forces the animal into long periods of immobility at experimenter-chosen locations, allowing electrophysiological analysis of the immediate changes in neural activity. The paradigm relies on the use of innate thermotaxis behaviors, leading the rodent to find and stay immobile in a heated area while the ambient temperature remains cold. Utilization of natural thermotaxis mechanisms enhances the ethological validity of the findings, an important feature in designing behavioral paradigms for animals [4].
One potential issue that arose is that the current design of the ThermoMaze is relatively small (20 cm × 20 cm). Traditional open fields for mice measure 40 cm × 40 cm. Slight modifications to the 3D floor design (https://github.com/misiVoroslakos/3D_printed_designs/blob/main/ThermoMaze/3D_printed_design/cooling_box_v03_platform.stl) and to the cast acrylic cut will enable the maze to be scaled to bigger dimensions. Additionally, the current size leads the animal to follow the walls to each of the four hotspots (located at the corners) and rarely visit the central portion of the maze. We found that altering hotspot location leads the mice to find the new hotspot away from a corner, providing better spatial coverage of the maze [10]. Given small rodents' aversion to large, open spaces, this will pose a problem irrespective of maze size.
The reliance on innate behavioral mechanisms led the rodents to rapidly learn the task (i.e., finding the hotspot), requiring only three practice sessions on average before an animal was completely trained. Additionally, the paradigm does not alter brain temperatures, signifying that the electrophysiological data represents naturally occurring changes and not some brain temperature–dependent changes. The place-specific firing patterns of neurons within sharp wave ripples, both inhibitory (interneurons) and excitatory (pyramidal cells), indicate that many consolidation processes occur in the periods of immobility immediately following a behavior. Therefore, utilization of this paradigm will enable researchers to gather large data sets on processes that have yet to be thoroughly investigated.
To facilitate widespread use of the paradigm, we provide detailed descriptions of the construction process, along with links for purchasing all necessary equipment. We further include links to the 3D-printable elements and relevant codes, accessible on the Buzsaki Lab GitHub (https://github.com/misiVoroslakos/3D_printed_designs/tree/main/ThermoMaze). We hope that this helps to foster the use of the ThermoMaze, a non-aqueous analog of the Morris water maze.
Acknowledgments
Supported by MH122391, and U19NS107616. This protocol was used in ‘ThermoMaze: A behavioral paradigm for readout of immobility-related brain events’ [10]. We thank Yiyao Zhang, Anna Maslarova and Leeor Alon for their help with different aspects related to the experiments.
Competing interests
There are no conflicts of interest or competing interest.
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Article Information
Publication history
Received: Apr 19, 2024
Accepted: Jun 27, 2024
Available online: Jul 24, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,045 | https://bio-protocol.org/en/bpdetail?id=5045&type=0 | # Bio-Protocol Content
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Reconstitution of the Melibiose Permease of Salmonella enterica serovar Typhimurium (MelBSt) into Lipid Nanodiscs
PH Parameswaran Hariharan
LG Lan Guan
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5045 Views: 426
Reviewed by: Chiara AmbrogioIstvan Stadler Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Feb 2024
Abstract
Membrane proteins play critical roles in cell physiology and pathology. The conventional way to study membrane proteins at protein levels is to use optimal detergents to extract proteins from membranes. Identification of the optimal detergent is tedious, and in some cases, the protein functions are compromised. While this detergent-based approach has produced meaningful results in membrane protein research, a lipid environment should be more suitable to recapture the protein’s native folding and functions. This protocol describes how to prepare amphipathic membrane scaffold-proteins (MSPs)-based nanodiscs of a cation-coupled melibiose symporter of Salmonella enterica serovar Typhimurium (MelBSt), a member of the major facilitator superfamily. MSPs generate nano-assemblies containing membrane proteins surrounded by a patch of native lipids to better preserve their native conformations and functions. This protocol requires purified membrane protein in detergents, purified MSPs in solution, and detergent-destabilized phospholipids. The mixture of all three components at specific ratios is incubated in the presence of Bio-Beads SM-2 resins, which absorb all detergent molecules, allowing the membrane protein to associate with lipids surrounded by the MSPs. By reconstituting the purified membrane proteins back into their native-like lipid environment, these nanodisc-like particles can be directly used in cryo-EM single-particle analysis for structure determination and other biophysical analyses. It is noted that nanodiscs may potentially limit the dynamics of membrane proteins due to suboptimal nanodisc size compared to the native lipid bilayer.
Key features
• This protocol was built based on the method originally developed by Sligar et al. [1] and modified for a specific major facilitator superfamily transporter
• This protocol is robust and reproducible
• Lipid nanodiscs can increase membrane protein stability, and reconstituted transporters in lipid nanodiscs can regain function if their function is compromised using detergents
• The reconstituted lipids nanodisc can be used for cryo-EM single-particle analysis
Keywords: MelBSt Membrane scaffold proteins Major facilitator superfamily Membrane proteins Nanodiscs Cryo-EM single-particle analysis Reconstitution with lipids
Graphical overview
Figure 1. Overview of the protocol to prepare MelBSt lipid nanodiscs. Section A illustrates the preparation of detergent-destabilized lipids. Section C-1 shows the steps for mixing the detergent-stabilized lipids with purified MelBSt and then with purified membrane scaffold protein MSP1E3D1. Section C-2 illustrates the formation of lipid nanodiscs upon incubation with detergent-adsorbing Bio-Beads SM2. Section C-3 illustrates the Ni+-agarose resin affinity chromatography to isolate the reconstituted MelBSt nanodiscs. DDM, n-Dodecyl-β-D-maltopyranoside; UDM, n-Undecyl-β-D-maltopyranoside.
Background
Membrane proteins and phospholipids are two major essential components of cell membranes. Lipid-embedded membrane proteins play critical roles in supplying nutrients and extruding unwanted or toxic substances by providing specific paths that allow polar or charged molecules across the cell membrane at a rate that effectively meets cellular needs [2]. The functions of membrane proteins are carried out in the lipid environment, and their activities are modulated or regulated by lipids [3]. The interactions between membrane proteins and phospholipids, including cholesterol, are dynamic and complicated, and some membrane proteins have binding site(s) to tightly interact with specific lipids [4,5]. The roles of lipids on membrane proteins vary from stabilization and oligomerization to modulating conformational dynamics [6,7]. This is a poorly characterized research area due to technical challenges stemming from intrinsic hydrophobicity. The conventional way to conduct the study of membrane proteins at protein levels is to use optimal detergents to extract proteins from the membranes for their function analyses. Identification of the optimal detergent is tedious and often challenging. Accordingly, varied methods have been developed to overcome this problem [8–10]; one is to reconstitute the protein back to a lipid environment after it has been purified, such as proteoliposomes and lipid nanodiscs [1,11–13]. The membrane scaffold protein (MSP)-based lipid nanodiscs [1] contain a lipid-bilayer core around membrane proteins, which retain and provide a more native-like environment than detergent micelles. In most cases, those protein nanodiscs contain a single copy of membrane protein per particle. The nanodisc-like samples have successfully facilitated cryo-EM single-particle reconstruction and membrane biophysical analyses. Reconstitution into the native-like environments allows some membrane proteins to regain function. For example, a protein that lost its function when solubilized in the detergents can bind ligands again after being reconstituted into proteoliposomes or lipid nanodiscs [5,14–16].
Compared to proteoliposomes, the nanodisc form lacks membrane-sidedness, and the membrane proteins are exposed to bulk solvents at both hydrophilic surfaces. This also means that the nanodisc form cannot be used to test the transport activity of the reconstituted transporters since no boundary exists. It is also noted that nanodiscs may potentially limit the dynamics of membrane proteins if their size is suboptimal.
In this protocol, we describe with great detail the reconstitution of a cation-coupled melibiose symporter of Salmonella enterica serovar Typhimurium (MelBSt) into nanodisc form (Figure 1), which has been validated by cryo-EM single-particle analysis and biophysical analysis [17,18]. MelBSt is a member of the major superfamily of transporters with 12 transmembrane helices embedded in the membrane [19]. This family of proteins is dominated by a hydrophobic domain with limited hydrophilic surfaces. Maintaining the protein's stability and function after extraction from native membranes is often challenging. MelBSt catalyzes the symport of a galactopyranoside with either H+, Li+, or Na+, and it is a unique model system for studying cation-coupled transport mechanisms [17,20–22]. This nanodisc protocol, developed for studying this protein, is robust and reproducible, and was originally developed by Sligar [1] with a modified lipids preparation procedure as described [23].
Materials and reagents
Reagents
E. coli polar lipids extract (Avanti, catalog number: 100500P)
n-Dodecyl-β-D-maltopyranoside (DDM) powder (Anatrace, catalog number: D310)
n-Undecyl-β-D-maltopyranoside (UDM) (Anatrace, catalog number: U300)
Bio-Beads SM-2 adsorbents (Bio-Rad, catalog number: 1523920)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
Sodium chloride (NaCl) (Fisher bioreagents, catalog number: S271)
Tris base (RPI, catalog number: T60040)
Concentrated HCl (Fisher Chemical, catalog number: A144-212)
Imidazole (Acros Organics, catalog number: 301870010)
Glycerol (Fisher Bioreagents, catalog number: BP229)
Methanol (Fisher Chemical, catalog number: A452SK-4)
Ethanol (Fisher bioreagents, catalog number: BP28184)
MSP1E3D1∆His at 260 μM (purified by nickel-based affinity chromatography using INDIGO Ni-Agarose resin and processed by TEV protease digestion; alternative source, Cube Biotech, catalog number: 26162 and concentrated by centrifugal concentrator Vivaspin 20, 5,000 MWCO)
His-tagged MelBSt (purified by cobalt-based affinity chromatography using TALON® Metal Affinity Resin)
Liquid nitrogen (Airgas)
Solutions
0.001% Triton X100 bath liquid (see Recipes)
7.5% DDM (see Recipes)
MelBSt dialysis buffer (see Recipes)
TBS (see Recipes)
TBS + 5 mM imidazole buffer (column binding buffer) (see Recipes)
TBS + 25 mM imidazole buffer (column wash buffer) (see Recipes)
TBS + 300 mM imidazole buffer (column elution buffer) (see Recipes)
1 M Tris-HCl (see Recipes)
1 M imidazole (see Recipes)
Recipes
0.001% (v/v) Triton X100 bath liquid
Reagent Final concentration Amount
100% Triton X-100 0.001% 0.05 mL
Milli-Q H2O* n/a To 5 L
*Milli-Q H2O is prepared using Milli-Q IQ7000 model equipment with resistivity 18.2 MΩ·cm at 25 °C and total organic carbon (TOC) ≤ 5 ppb.
7.5% DDM (w/v)
Reagent Final concentration Amount
DDM powder 7.5% 0.75 g
Milli-Q H2O n/a Dissolved in 10 mL
*Freeze at -20 °C.
MelBSt dialysis buffer
Reagent Final concentration Amount
NaCl (4 M in milli-Q H2O) 150 mM 37.5 mL
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
Glycerol (100%) 10% (v/v) 100 mL
DDM (10%) (w/v) 0.01% (w/v) 1 mL
Milli-Q H2O n/a To 1 L
TBS (nanodiscs collection buffer)
Reagent Final concentration Amount
NaCl (4 M in milli-Q H2O) 150 mM 37.5 mL
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
Milli-Q H2O n/a To 1 L
TBS buffer with 5 mM imidazole (column binding buffer)
Reagent Final concentration Amount
NaCl (4 M in milli-Q H2O) 150 mM 37.5 mL
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
Imidazole (1 M, pH 7.5) 5 mM 5 mL
Milli Q H2O n/a To 1 L
TBS buffer with 25 mM imidazole (column washing buffer)
Reagent Final concentration Amount
NaCl (4 M in milli-Q H2O) 150 mM 37.5 mL
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
Imidazole (1 M, pH 7.5) 25 mM 25 mL
Milli Q H2O n/a To 1 L
TBS buffer with 300 mM imidazole (column eluting buffer)
Reagent Final concentration Amount
NaCl (4 M in milli-Q H2O) 150 mM 37.5 mL
Tris-HCl (1 M, pH 7.5) 20 mM 20 mL
Imidazole (1 M, pH 7.5) 300 mM 300 mL
Milli Q H2O n/a To 1 L
Note: All solutions are stored at 4 °C.
1 M Tris-HCl
1 M Tris base in milli-Q H2O was adjusted to pH 7.5, using concentrated HCl.
Reagent Final concentration Amount
Tris base 1 M 121.1 g
Milli-Q H2O Dissolve to 900 mL
Concentrated HCl Milli-Q H2O Adjust to pH 7.5
1 M Imidazole
1 M Imidazole in milli-Q H2O was adjusted to pH 7.5, using concentrated HCl.
Reagent Final concentration Amount
Imidazole 1 M 68.08 g
Milli-Q H2O Dissolve to 900 mL
Concentrated HCl Milli-Q H2O Adjust to pH 7.5 and fill to 1 L
Laboratory supplies
20 mL glass vials (DWK Life Sciences, catalog number: W224589)
Plastic conical tubes, 50 mL (Thermo Scientific, catalog number: 339653)
NuncTM serological pipettes, 5 mL (Thermo Scientific, catalog number: 170355)
1.5 mL Eppendorf tubes (FisherBrand, catalog number: 05-408-129)
2 mL Eppendorf tubes (FisherBrand, catalog number: 05-408-138)
Stir bars (FisherBrand, catalog number:14-513-82)
Transfer pipettes (VWR, catalog number: 16001-194)
Slider-A-Lyzer dialysis cassettes, 10 k MWCO (Thermo Scientific, catalog number: 66810)
Centrifugal concentrator Vivaspin 2, 50,000 MWCO PES (Sartorius, catalog number: VS0231)
Centrifugal concentrator Vivaspin 20, 5,000 MWCO PES (Sartorius, catalog number: VS2012)
INDIGO Ni-agarose resin, 50% slurry (Cube Biotech, catalog number: 75105)
TALON® metal affinity resin (Takara, catalog number: 635503)
Poly-Prep chromatography column (Bio-Rad, catalog number: 731-1550)
Econo-Column® chromatography column, 2.5 × 10 cm (Bio-Rad, catalog number: 737-2512)
Econo-Column funnel, 250 mL reservoir (Bio-Rad, catalog number: 731-0003)
SDS-12% gel in 1.5 mm thickness (prepared according to the instruction manual of Mini-PROTEAN Tetra Vertical Electrophoresis kit)
Equipment
Branson ultrasonic cleaner (Branson, model: 2510)
Glass dewar flask for liquid nitrogen (FisherBrand, catalog number: 10-196-7)
Magnetic stirrer (Fisher Scientific, model: Isotemp)
Ultracentrifuge (Beckman Coulter, model: L-100XP Ultracentrifuge)
Eppendorf benchtop centrifuge (Eppendorf, model: 5417R)
Vortex mixer (Fisher Scientific, model: Analog Vortex Mixer, catalog number: 02215365)
Pipette controller (DrummondTM, model: Portable Pipet-AidTM XP)
Mini-PROTEAN Tetra Vertical Electrophoresis kit (Bio-Rad, catalog number: 165-8027FC)
NGC Quest FPLC system (Bio-Rad, model: NGC Quest 10 Plus, catalog number: 7880003)
SuperdexTM 200 Increase small-scale S columns, 10 × 300 mm (Cytiva, catalog number: 28990944)
Spectrophotometer (Thermo Fisher, model: Genesys 10-S UV-Vis)
Procedure
Prepare 5 mL fresh lipid mixture at 40 mg/mL in 7.5% DDM solution
Weigh 200 mg of E. coli polar lipids extract powder using a pre-weighed 50 mL plastic conical tube.
Add 4.5 mL of 7.5% DDM (see Recipe 2) to the dry lipid.
Vortex the mixture for 2–3 min.
Sonicator setup: Fill the sonicator with ice-cold 0.001% Triton X100 bath liquid (see Recipe 1) and adjust the water level to achieve the maximum strength of sonication.
Place the tube into the sonicator tank and sonicate the suspension for 1 min. During sonicating, pipette the suspension up and down using a 5 mL plastic serological pipette on a portable pipette controller. Use the same pipette until the end of step A7.
Cool on ice for 5 min.
Repeat steps A4–5 until completely dissolved. It might take 5–7 cycles. An optional 1 min vortex can also be applied at the beginning of each cycle.
Note: Avoid overheating. Shipping freezer packs can be used during intervals to cool the bath liquid. Maintain ice-cold bath liquid to the optimal level and top it off if required.
Measure the volume of the suspension using the same 5 mL pipette and adjust the volume to 5 mL using 7.5% DDM solution to obtain 40 mg/mL.
Conduct the freeze-thaw-sonicate cycle 3–4 times until the lipid mixture is translucent.
Freeze the 5 mL lipid samples in the same tube by plunging and rotating the tube into a glass dewar flask containing liquid nitrogen; hold the tube at an angle to provide more surface area to create a thin layer of frozen lipids around the wall of the tube. The purpose of the thin layer is not only to make thawing faster in the next step but also to make sure the entire suspension is completely frozen (Figure 2A).
Thaw at room temperature until the lipid sample becomes liquid. Typically, this may take approximately 5–10 min with shaking; allow a brief warming period (~5 s) by hand.
Sonicate the liquid sample for 30 s.
Cool on ice for 5 min before the next freeze-thaw-sonicate cycle.
Keep the translucent lipid mixture at room temperature for the next step (Figure 2B).
Aliquot the excess lipid mixture by 0.5 mL into 1.5 mL Eppendorf tubes, flash freeze in liquid nitrogen, and store at -80 °C.
Figure 2. Lipids preparation. A. Frozen detergent-stabilized lipid suspension on the wall of the 50 mL tube. B. Processed pale-yellowish translucent lipid suspension after freeze-thaw-sonicate cycles.
Note: When using the frozen lipids from -80 °C, thaw them at room temperature, sonicate for 30 s, and leave the sample at room temperature for immediate use. The lipids can be frozen again with liquid nitrogen and used 2–3 times.
Bio-Beads preparation
Note: Prepare the Bio-Beads before setting up the nanodisc reconstitution.
Pack the ~20–25 g of Bio-Beads in a Bio-Rad glass Econo-Column® cartography column (2.5 × 10 cm, volume ~50 mL) fitted with a 250 mL solution reservoir funnel.
Wash once with 10 volumes (~200–250 mL) of methanol and let it drain completely.
Wash once with 10 volumes (~200–250 mL) of ethanol and let it drain completely.
Wash 10 times with 10 volumes (2–2.5 L) of milli-Q water.
Remove the funnel and transfer the prepared Bio-Beads in milli-Q water into a 50 mL plastic conical tube and store at 4 °C. The prepared Bio-Beads samples in milli-Q water can be stored for 4–6 months.
Reconstitution of nanodiscs
Required materials:
40 mg/mL E. coli lipids in 7.5% DDM (see Recipe 2).
MelBSt dialysis buffer (see Recipe 3).
Purified His-tagged MelBSt in the defined MelB dialysis buffer.
Purified and TEV-treated membrane scaffold protein MSP1E3D1∆His (alternate source: Cube Biotech, catalog number: 26162).
20 mL glass vial with cap.
Mixing. Prepare a suspension containing 6.65 mM E. coli lipids and 1 mg/mL His-tagged MelBSt, corresponding to a ratio of His-tagged MelBSt to lipids of 1:350 mole/mole.
In a 20 mL glass vial, take 1 mL of His-tagged MelBSt at 1 mg/mL concentration (~19 μM).
Add 133 μL of 50 mM lipids and mix using pipettes with no vortex.
Incubate the mixture for 10 min at room temperature.
Add 365 μL of 260 μM MSP1E3D1∆His to the MelBSt/lipid mixture, bringing the final concentration to 95 μM and a ratio of MSP1E3D1∆His to MelBSt of 5:1.
Incubate the mixture (Figure 3A) at room temperature with intermittent mixing for 30 min.
Note: The mixture should be clear at the end of the 30 min incubation, indicating no protein precipitation (see General note 1 and 2).
Figure 3. Reconstitution procedure of MelBSt nanodiscs. A. Mixing the purified MelBSt and MSP1E3D1∆His protein with the detergent-destabilized phospholipids at 1:5:350 ratios, respectively. B. Collecting the slightly turbid suspension containing MelBSt-reconstituted lipid nanodiscs and the empty nanodisc without MelBSt.
Detergent removal by incubation with Bio-Beads (1 mg of MelBSt may need ~ 800 mg beads).
Weigh the prepared Bio-Beads using a pre-weighed 2 mL microfuge tube after removing the water using a transfer pipette.
Wash the Bio-Beads with 1 mL of TBS buffer (see Recipe 4) by mixing and removing the buffer three times.
Add 2/3 of treated Bio-Beads (~500 mg) to the lipids/protein mixture (Figure 3B).
Incubate at 4 °C for 2 h with mild stirring.
Add the remaining Bio-Beads (~300 mg).
Continue the incubation overnight at 4 °C under mild stirring/mixing.
Note: The nanodiscs should be formed after removing the detergents with the Bio-Beads, and the suspension should be cloudy, indicative of nanodiscs (Figure 3B).
Transfer the nanodiscs-containing slightly turbid suspension into a fresh 1.5 mL Eppendorf tube using a transfer pipette (or gel-loading tips) (Figure 3B).
Centrifuge at 20,000× g for 10 min at 4 °C to remove the traces of contaminating Bio-Beads.
Transfer the nanodiscs-containing supernatant into a fresh 2 mL microfuge tube.
Note: At this step, the nanodiscs may contain one or two molecules of MelBSt per disc, or they may be empty discs with no MelBSt. The nanodisc samples are in the TBS buffer after completely removing the detergents.
Note: In the cryo-EM single-particle analysis, we observed the majority of loaded nanodisc containing one molecule of MelBSt. Although its heterogeneity interferes with determining protein concentrations, SDS-PAGE analysis can analyze the protein species and their stoichiometry.
Isolation of MelBSt-reconstituted nanodiscs by affinity chromatography.
Note: The empty nanodiscs with no His-tag and the His-tagged MelBSt nanodiscs can be separated by affinity chromatography using Indigo Ni-agarose resin (Figure 4A). The empty nanodiscs are in the void, and the His-tagged MelBSt nanodiscs are adsorbed by the Ni-NTA beads. We usually conduct this separation using a gravity column at room temperature. However, the following steps can be performed at 4 °C and/or using pre-packed affinity chromatography columns on an FPLC system.
Pack 2 mL of 50% INDIGO Ni-agarose resin to a 1 mL bed volume on a chromatography column.
Wash the column using 10 mL (10× volume) of milli-Q water.
Wash the column using 10 mL of the TBS buffer containing 5 mM imidazole (see Recipe 5).
Add 1 M of imidazole solution into the nanodisc mixture to a final concentration of ~5 mM.
Load the nanodiscs mixture onto the washed column.
Wash the column with 10 mL (10× volume) of 5 mM-imidazole TBS buffer (see Recipe 5).
Wash the column twice with 10 mL of 25-mM imidazole TBS buffer (see Recipe 6) to remove the empty nanodiscs without a His tag.
Elute the bound His-tagged MelBSt nanodiscs with 10 mL of 300-mM imidazole TBS buffer (see Recipe 7).
Immediately collect fractions at each 0.5 mL in 1.5 mL Eppendorf tubes.
Pool the protein-containing fractions (judged by 280 nm absorption using the elution buffer to blank).
Transfer all pooled solution into a 2 mL Vivaspin 2 centrifugal concentrator 50,000 MWCO PES and centrifuge at 3,000× g for 5 min at 4 °C. Mix the solution using a transfer pipette before starting another centrifugation at the same condition until the volume reduces to ~0.5 mL to obtain the protein concentration to ~3 mg/mL.
Dialyze MelBSt nanodiscs against the TBS buffer (see Recipe 4) overnight using a dialysis cassette with 10,000 MWCO.
Note: Estimation of protein concentration: using a spectrophotometer, measure A280 nm absorbance value, which is used in the following equation:
Protein concentration, C (M) = A280 nm/extinction coefficient () × light path-length (l),
= 135110 (https://web.expasy.org/protparam/) based on 1 MelBSt: 2 MSP1E3D1∆His per nanodisc.
l = 1 when using a 1 cm cuvette
Data analysis
SDS-12% PAGE. An aliquot of protein fraction (10 μL) from the Ni-affinity chromatography was analyzed using SDS-12% PAGE and stained with Coomassie blue (Figure 4B). The 54 kDa His-tagged MelBSt migrated to 37 kDa, and the 30 kDa His-tag-removed MSP1E3D1 migrated slightly above 25 kDa. The unbound fraction in the void shows a strong MSP1E3D1 band and weak MelBSt band, which indicates that most nanodiscs have no reconstituted MelBSt. The elution fractions indicate the great enrichment of MelBSt-containing nanodisc with a stochiometric ratio of 1:2 with MSP1E3D1.
Size exclusion chromatography. Load 0.5 mL of concentrated and dialyzed nanodiscs onto a Superdex 200 Increase (10 × 300 mm) column pre-equilibrated with TBS (see Recipe 4) on a Bio-Rad NGC Quest FPLC system. Figure 4C presents the chromatogram of the MelBSt nanodiscs. The chromatogram peak fractions collected using the BioFrac fraction collector can be directly used for downstream applications such as biochemical studies [15,18], cryo-EM single-particle analysis [17], etc., as described in the validation section.
Figure 4. Separation of the mixed nanodiscs sample by Ni-affinity chromatography. A. Illustration of the gravity flow chromatography using a column packed with INDIGO Ni-agarose resin. B. SDS-12%PAGE analysis. From each fraction, 10 μL were loaded and the gel was stained by Coomassie blue. C. Size exclusion chromatography shows the major peak of the reconstituted MelBSt in lipid nanodiscs is sharp and symmetric.
Validation of protocol
This protocol or parts of it have been used and validated in the following research article(s):
In the J Gen Physiol article [18], MelBSt nanodiscs were prepared to test the binding of MelB regulatory protein. In the J Mol Biol article [15], a few single-site mutations of MelBSt largely decreased protein stability, preventing the downstream biophysical analysis. Reconstitution into lipid nanodiscs allowed cation binding and substrate binding assays to be possible. Figure 5 shows different orientations of nanodiscs carrying MelBSt in complex with the nanobody725_4, NabFab, and anti-Fab Nb. In the eLife article [17], MelBSt was reconstituted into nanodiscs, which increased protein stability and single-particle picking.
Figure 5. 2D class averages. Representative data from the study was described in the eLife article [17]. The sample containing the wild-type MelBSt in lipid nanodiscs using MSP1E3D1∆His, the MelBSt-specific Nb725_4, NabFab, and anti-Fab Nb were used for imaging using Titan Krios TEM with a K3 detector of S2C2, Stanford, CA, as described [17]. The image processing, particle pick, and 2D classification were performed in CryoSPARC program. Upper row: side view of the nanodiscs carrying the MelBSt complex; each class contains 4,000–7,000 particles. Lower row: top view of nanodiscs carrying MelBSt complex; each class contains 1,000–4,000 particles. Bar size, 140 Å.
General notes and troubleshooting
General notes
Tips for reconstituting other membrane proteins:
An essential modification in Section C is to use a protein-specific buffer to maintain membrane protein stability at the mixing step. The most important component is to use the detergent optimal to the membrane protein of interest. The detailed detergent study in [24] contains useful information for guiding detergent selection.
In the mixing step of Section C, the concentration of the membrane proteins should be < 1 mg/mL.
Troubleshooting
Problem 1: Protein precipitation during the mixing step.
Possible cause: The quality of the protein sample is poor.
Solution: A) Perform all incubations on the ice. B) Optimize the membrane protein buffer and dilute the protein if necessary. C) Increase the ratio of MSPs with the membrane protein up to 10:1. D) Select a suitable MSP type according to the size and hydrodynamic radius of the membrane protein.
Problem 2: No membrane protein with nanodiscs.
Possible cause: Protein aggregations or incompatibility with the selected lipids mixture.
Solution: A) The protein may require a different lipid composition. Lipid selection is important. The lipid components of the prokaryotic and eukaryotic membranes are different. The E. coli polar lipids extract (Avanti, catalog number: 100500P) can be first selected to test for most bacterial membrane proteins. More information on lipids selection is available in [25]. However, a trial-and-error approach is often needed to identify optimal lipid composition.
Laboratory safety
Use proper personal protective equipment (PPE) and adhere to general cryogenic safety procedures while handling liquid nitrogen. Transfer liquid nitrogen in well-ventilated areas to prevent generating low-oxygen surroundings and causing breathing health hazards.
Review the MSDS provided by the manufacturer for all chemicals and materials.
Acknowledgments
We thank Dr. Guillermo Altenberg for providing the expression vectors of MSP1E3D1 and TEV-protease. We also thank Dr. Mariana Fiori for providing valuable feedback during the initial optimization of this protocol for MelBSt reconstitution into nanodiscs. This work was supported by the National Institutes of Health Grant 1R35GM153222-01 to L.G.
Competing interests
The authors declare no competing financial interests.
References
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Hariharan, P., Tikhonova, E., Medeiros-Silva, J., Jeucken, A., Bogdanov, M. V., Dowhan, W., Brouwers, J. F., Weingarth, M. and Guan, L. (2018). Structural and functional characterization of protein-lipid interactions of the Salmonella typhimurium melibiose transporter MelB. BMC Biol. 16(1): 85. https://doi.org/10.1186/s12915-018-0553-0.
Drew, D. and Boudker, O. (2024). Ion and lipid orchestration of sondary active transport. Nature 626(8001): 963–974. https://doi.org/10.1038/s41586-024-07062-3.
Dowhan, W., Mileykovskaya, E. and Bogdanov, M. (2004). Diversity and versatility of lipid-protein interactions revealed by molecular genetic approaches. Biochim Biophys Acta. 1666(1–2): 19–39. https://doi.org/10.1016/j.bbamem.2004.04.010.
Ryabichko, S., Ferreira, V. M., Vitrac, H., Kiyamova, R., Dowhan, W. and Bogdanov, M. (2020). Cardiolipin is required in vivo for the stability of bacterial translocon and optimal membrane protein translocation and insertion. Sci Rep. 10(1): 6296. https://doi.org/10.1038/s41598-020-63280-5.
Guan, L. (2023). The rapid developments of membrane protein structure biology over the last two decades. BMC Biol. 21(1): 300. https://doi.org/10.1186/s12915-023-01795-9.
Fiori, M. C., Jiang, Y., Zheng, W., Anzaldua, M., Borgnia, M. J., Altenberg, G. A. and Liang, H. (2017). Polymer Nanodiscs: Discoidal Amphiphilic Block Copolymer Membranes as a New Platform for Membrane Proteins. Sci Rep. 7(1): 15227. https://doi.org/10.1038/s41598-017-15151-9.
Lee, H. J., Lee, H. S., Youn, T., Byrne, B. and Chae, P. S. (2022). Impact of novel detergents on membrane protein studies. Chem. 8(4): 980–1013. https://doi.org/10.1016/j.chempr.2022.02.007.
Denisov, I. G., Grinkova, Y. V., Lazarides, A. A. and Sligar, S. G. (2004). Directed self-assembly of monodisperse phospholipid bilayer Nanodiscs with controlled size. J Am Chem Soc. 126(11): 3477–3487. https://doi.org/10.1021/ja0393574.
Levental, I. and Lyman, E. (2023). Regulation of membrane protein structure and function by their lipid nano-environment. Nat Rev Mol Cell Biol. 24(2): 107–122. https://doi.org/10.1038/s41580-022-00524-4.
Nath, A., Atkins, W. M. and Sligar, S. G. (2007). Applications of phospholipid bilayer nanodiscs in the study of membranes and membrane proteins. Biochemistry. 46(8): 2059–2069. https://doi.org/10.1021/bi602371n.
Amin, A., Hariharan, P., Chae, P. S. and Guan, L. (2015). Effect of Detergents on Galactoside Binding by Melibiose Permeases. Biochemistry. 54(38): 5849-5855. https://doi.org/10.1021/acs.biochem.5b00660.
Katsube, S., Liang, R., Amin, A., Hariharan, P. and Guan, L. (2022). Molecular Basis for the Cation Selectivity of Salmonella typhimurium Melibiose Permease. J Mol Biol. 434(12): 167598. https://doi.org/10.1016/j.jmb.2022.167598.
Zoghbi, M. E., Cooper, R. S. and Altenberg, G. A. (2016). The Lipid Bilayer Modulates the Structure and Function of an ATP-binding Cassette Exporter. J Biol Chem. 291(9): 4453–4461. https://doi.org/10.1074/jbc.M115.698498.
Hariharan, P., Shi, Y., Katsube, S., Willibal, K., Burrows, N. D., Mitchell, P., Bakhtiiari, A., Stanfield, S., Pardon, E., Kaback, H. R., et al. (2024). Mobile barrier mechanisms for Na(+)-coupled symport in an MFS sugar transporter. eLife 12. https://doi.org/10.7554/eLife.92462.
Hariharan, P. and Guan, L. (2021). Cooperative binding ensures the obligatory melibiose/Na+ cotransport in MelB. J Gen Physiol. 153(8). https://doi.org/10.1085/jgp.202012710.
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Guan, L. and Hariharan, P. (2021). X-ray crystallography reveals molecular recognition mechanism for sugar binding in a melibiose transporter MelB. Commun Biol. 4(1): 931. https://doi.org/10.1038/s42003-021-02462-x.
Hariharan, P., Bakhtiiari, A., Liang, R. and Guan, L. (2024). Distinct roles of the major binding residues in the cation-binding pocket of the melibiose transporter MelB. J Biol Chem: 107427. https://doi.org/10.1016/j.jbc.2024.107427.
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Article Information
Publication history
Received: May 10, 2024
Accepted: Jul 3, 2024
Available online: Jul 21, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Molecular Biology > Protein > Detection
Biophysics > Electron cryotomography
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A Social Stimulation Paradigm to Ameliorate Memory Deficit in Alzheimer's Disease
QR Qiaoyun Ren
SW Susu Wang
WX Wei Xie
AL An Liu
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5046 Views: 385
Reviewed by: Salim GasmiRachael E. Hokenson Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Feb 2024
Abstract
Alzheimer's disease (AD) poses a global health threat, progressively robbing patients of their memory and cognitive abilities. While it is recognized that meaningful social contact can alleviate the symptoms of dementia in AD patients, the precise mechanisms by which social stimulation mitigates AD symptoms remain poorly understood. We found that social interaction with novel mice, also known as novel social, simulated meaningful socializing. Therefore, we developed the multiple novel social (MNS) stimulation paradigm to train AD model mice and found that MNS effectively alleviated cognitive deficits in AD mice. This discovery not only opens up a new avenue for investigating the relationship between social stimulation and Alzheimer's disease but also lays the groundwork for delving into the underlying mechanisms, thereby providing crucial theoretical support for developing novel strategies to treat Alzheimer's disease.
Key features
• Designing a new social stimulation method to simulate meaningful social interactions in daily life.
• The MNS stimulation protocol spans 14 days, with one novel mouse introduced to the subject mice each day.
• The subjects were 2.5-month-old FAD4T mice, simulating patients with mild cognitive impairment (MCI).
• Results of behavioral tests confirm the efficacy of MNS in reducing cognitive deficits in the AD model.
Keywords: Alzheimer's disease Multiple novel social stimulation Cognition deficit Cognitive ability Social memory
Graphical overview
Background
Alzheimer’s disease (AD) is one of the most influential and common neurodegenerative diseases in the world. It also ranks as the fifth leading cause of mortality among elderly individuals worldwide [1,2]. Memory and cognitive decline are recognized as the most incapacitating aspects of Alzheimer's disease [3]. In recent years, there have been emerging drugs in clinical practice that show promise in alleviating the dementia symptoms associated with Alzheimer’s disease [4–6]. However, the precise effects and potential side effects of these medications on cognitive function and disease progression are still being closely monitored. Non-pharmacological therapies have also been increasingly recognized and valued. Non-pharmacological treatments, such as engaging in physical activities, cognitive exercises, social interactions, and managing vascular and metabolic risks, are integrated alongside drug therapies for Alzheimer's disease [2].
Despite the recognition of non-pharmacological interventions, there remains a research gap regarding the mechanisms underlying the benefits of social activities. Studies have shown that an enriched social network and frequent social contact can prevent dementia and increase patients' cognitive reserve [7–11]. Additionally, social isolation exacerbates anxiety and asymmetrical brain atrophy in Alzheimer's patients [12]. Conversely, complex social interactions can improve episodic memory, enhance brain reserve, and reduce the risk of dementia [13]. Although the benefits of social stimulation for Alzheimer's patients have been well established, the specific biological mechanisms remain unclear.
We found that a single novel social (SNS) stimulation with an unfamiliar mouse effectively activated the α-secretase activity in the ventral hippocampus of both wildtype (WT) and AD model mice and inhibited the amyloidogenic-cleavage pathway but was not enough to reduce the accumulation of Aβ in AD model mice [14]. Therefore, we implemented a 14-day multiple novel social (MNS) protocol to extend the intervention effect of this paradigm by increasing the number of social stimuli, allowing the subject mice to experience 14 novel social interactions. Given that the social memory of mice is transient, typically lasting only a few days [15], we cycled through seven different novel mice (designated N1–N7) for the first and second seven-day periods of the protocol. It should be noted that we also recommend using 14 different unfamiliar mice for the 14-day MNS protocol. As a control, group-housed (GH) subject mice were only able to interact with their familiar littermates.
After the MNS treatment, through the novel object recognition test and 3-chamber test, we found that the MNS stimulation paradigm significantly improved the cognitive ability and social memory of AD mice. Therefore, the MNS stimulation paradigm represents a promising method to alleviate dementia symptoms in Alzheimer’s disease.
Materials and reagents
Mice [all animals were housed in standard laboratory conditions with ad libitum access to food and water, as well as a 12/12 h light/dark cycle (light: 7:00 AM–7:00 PM), a temperature range of 22–26 °C, and a humidity level of 55%–60%. The mice used in this study were cared for in accordance with the authorized protocols of Southeast University, Nanjing, China)].
FAD4T mice [C57BL/6 genetic background, 10–12 weeks old. FAD4T mice co-expressed human APP with the Swedish (KM670/671NL) and India (V717F) variants, together with human mutant PS1 (M146L, L286V), driven by the mouse Thy1 promoter (Gempharmatech Co., Ltd, catalog number: T053302)]
Mice used for novel stimulation (N1–N7) and social test (S1–S2) (C57BL/6J genetic background, 10–12 weeks old, 22–25 g) (Gempharmatech Co., Ltd, catalog number: N000013, Jiangsu, China)
Nitrile laboratory gloves (Guangming, China)
Paper towels (Breeze, China)
75% Ethanol (Alladin, catalog number: A171299)
Equipment
Mouse breeding room with constant temperature system, ventilation system, and automatic light timing system
Home cage (380 mm × 180 mm × 170 mm, with a height under the partition bar inside the cage of 13 cm, which meets the requirements of the GB14925 national standard. It can accommodate 5–8 experimental mice weighing 20–30 g) (SHINVA, China)
Small electronic scale (127 mm × 106 mm × 19 mm) (Kubei, China)
Metal pen holder (golden, diameter 8 cm, height 10.3 cm, weight 140 g) (Wuling, China)
Timer (82 mm × 62 mm × 24 mm) (Anytime, model: XL-009B)
Spray bottle (500 mL) (LDPE, Sangon Biotech, catalog number: F505008)
Objects for novel object recognition (NOR) test (two bright-red metal cubes, 83 mm × 83 mm × 83 mm, were used as similar objects; one light-purple metal cylinder, 85 mm in diameter and 84 mm in height, was used as the novel object) (Aiziling, China)
Mouse behavioral testing room with constant temperature system, ventilation system, and automatic light timing system
Behavioral testing apparatus (Xinruan, China)
NOR apparatus
A square Plexiglas box (50 cm × 50 cm × 50 cm) with four 50 cm-high sidewalls and an open roof. In addition, two metal boxes with heavy objects inside were used as similar objects, which maintained consistency in size, color, and shape. Another metal box with a heavy object, similar in size but different in color and shape, was used as the novel object
3-chamber apparatus
A rectangular box made of Plexiglas (50 cm × 25 cm) with a detachable base, creating three distinct compartments divided by two partitions. Each partition contained a small door (10 cm × 5 cm) to facilitate mouse’s movement between the different chambers. In addition, there were two metal pen holders that were identical in size, shape, and color, used for restraining stranger mice.
Computer (Lenovo, model number: Tianyi 510pro-14IMB)
Camera (MOKOSE, model number: HDC10)
Software and datasets
Video tracking and analyzing software (Noldus, EthoVision XT 13 software)
Statistical analysis software (Prism 8.0.1, GraphPad, https://www.graphpad.com/)
Procedure
Pre-experimental preparation
Choose FAD4T mice that are 2.5 months old, ensuring that they exhibit standard body size and good health. Randomly allocate them to control and experimental groups, with each group maintaining 3–4 mice per cage.
Prepare seven cages for the novel mice that will be used for novel social stimulation. Each cage should contain about 3–5 mice. These novel mice should be from different parents, similar in age and body size to the FAD4T mice, and of the same sex. Place the cages at various locations in the breeding room to avoid olfactory similarity between the mice in each cage (these cages should be spaced at least three cage positions apart from each other).
Prepare 75% ethanol, paper towels, and metal pen holders. First, thoroughly clean the metal pen holder with 75% ethanol; then, carefully wipe it dry with a paper towel to ensure that no odor or impurities remain inside the pen holder.
To acclimate the FAD4T mice to the pen holders, place clean pen holders in their cages at 10:00 AM each day for 1 h, starting three days before the social stimulation is scheduled to begin.
Multiple novel social stimulation protocol
On the first day of social stimulation (Day 1), at 1:00 PM, place a clean pen holder in the cages of both the control and experimental FAD4T mice groups. Meanwhile, introduce the first wild-type novel mouse into the pen holder in the experimental FAD4T mice's cage, serving as the novel social stimulation “novel mouse” (N1). After 1 h, remove the N1 and pen holder, wash it with 75% ethanol, and then dry it with a paper towel.
On Day 2, repeat the procedure at 1:00 PM, introducing another wild-type novel mouse as the second social stimulation mouse (N2). After 1 h, remove the N2 and clean the pen holder with 75% ethanol, followed by drying it with a paper towel.
Continue this sequence and stimulate the subject mice with novel mice (N3–N7) daily until Day 7.
On Day 8, at 1:00 PM, start a new cycle of novel social stimulation starting from N1, which was used for novel stimulation on Day 1.
Repeat the novel social stimuli with N2–N7 from Day 9 to Day 14.
Post-stimulation assessment of cognitive ability and memory in FAD4T mice
Cognitive ability assessment (novel object recognition, NOR) (Figure 1)
Training phase
Individually place each mouse into the NOR apparatus, which contains two identical objects (A and A’). Give the mouse 10 min to explore these objects and utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map. Then, return it to the home cage.
Testing phase.
Twenty-four hours later, replace object A’ with a new, distinct object (B), which differs in both color and shape. Individually place each mouse into the arena and provide a 5 min period for it to freely explore. Utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map and return it to home cage.
Figure 1. Multiple novel social (MNS) stimulation rescues novel object recognition (NOR) impairment in FAD4T mice. A. Illustration of NOR test. B. Heat map of mouse movement in NOR test, with the training phase on top and the testing phase below. The FAD4T-MNS showed higher exploratory behavior toward the new object compared to the control group.
Social memory assessment (3-chamber test) (Figure 2)
In the first stage, place the test mouse in the central chamber, with two wired metal pen holders positioned in the bottom-left and top-right corners of the apparatus. Allow each mouse 10 min to freely explore. Utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map. Following habituation, gently return the test mouse to the central chamber, with both side doors open.
In the second stage (sociability test), place a stranger mouse (S1) inside one of the metal pen holders. Then, give the subject mouse another 10 min to explore the arena. Utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map.
In the third stage, return the test mouse to the central chamber for 10 min with the side doors closed. Utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map.
In the fourth stage (social memory test), introduce the second stranger mouse (S2) into the previously empty metal pen holder, and allow the subject mouse to explore the entire apparatus for 10 min. Utilize EthoVision XT 13 software to record and analyze the behavior of each mouse with a heat map.
Figure 2. Multiple novel social (MNS) stimulation rescues social memory in FAD4T mice. A. Illustration of the different stages of the 3-chamber test. B. Heat map of mouse movement in the 3-chamber test. The two groups of mice showed normal social abilities, and MNS stimulation increased the exploration time of FAD4T mice toward S2 mice.
Data analysis
Heatmap is indeed a very effective data visualization tool, especially in behavioral and neuroscience research, as it can help researchers quickly and intuitively understand the activity patterns of mice in complex environments. Through heatmaps, researchers can observe the activity of mice in specific experimental settings, such as their interest in certain objects or the time spent exploring certain areas. In behavioral research, heatmaps are usually generated by superimposing the mouse’s movement trajectory data on the floor plan of the experimental environment. The color depth of each point or area represents the duration of the mouse’s stay at that location, or the frequency of the mouse passing through that location. In this way, researchers can quickly identify the areas that mice prefer to explore (usually shown in dark red or warm tones) and the areas that are hardly visited at all (usually shown in blue or cold tones). Therefore, by observing the color and depth of the heatmap, we can understand the exploration time and tendency of mice for different objects or unfamiliar mice, and judge their cognitive and social memory levels.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Ren et al. [14]. Novel Social Stimulation Ameliorates Memory Deficit in Alzheimer's Disease Model through Activating α-Secretase. Journal of Neuroscience (Figure 5, panel A; Figure 6, panel F–L).
General notes and troubleshooting
After weaning (21 days after birth), the male and female mice are separated for breeding to prevent mating or pregnancy from affecting the experiment.
After the social stimulation program ends, the experimental mice and their cages are placed in the room where the behavioral tests are conducted to adapt to the new environment. Behavioral testing begins the following day.
The mouse breeding room and mouse behavioral testing room have a circadian rhythm with 12 h of light and 12 h of darkness (light: 7:00 AM–7:00 PM). Lighting intensity is around 100–200 lux, which is similar to the brightness of a typical office environment. The light quality should be white or cool-colored to avoid disrupting the mice’s circadian rhythms.
When moving mice from one location to another, use soft, padded forceps or gloves to avoid injuring the animals. Avoid sudden or jerky movements that can startle or distress the mice.
When conducting behavioral tests, such as novel object recognition or 3-chamber tests, ensure that the mice are placed gently into the testing apparatus and given time to acclimate before the experiment begins.
When observing the mice during the experiment, do so without causing them unnecessary stress. Avoid sudden movements or loud noises that can disrupt their behavior.
To minimize the stress and pressure on the mice during the stimulation or behavioral testing experiments, it is advisable to schedule these activities during periods of the day when the animals are known to exhibit lower corticosterone levels (8:00 AM–3:00 PM) [16,17]. This approach helps to ensure that the physiological responses of the mice to the experimental conditions are more representative of their baseline state and reduces the potential for stress-induced interference with the experimental outcomes.
If an animal demonstrates a pronounced bias or preference toward the right or left zone during the training phase or habituation phase, it is strongly advised to exclude it from the subsequent trials.
If during the novel object recognition phase, the total exploration time of the mouse for the objects does not exceed 8 s, the data for that mouse will be excluded.
The “stranger mice” in the 3-chamber test are not the same as the mice used for social stimulation. We use “stranger mice” and “novel mice” to distinguish between them.
In the 3-chamber test, the stages are conducted sequentially, one after the other. Each stage lasts 10 min, for a total of 40 consecutive minutes.
All the experiments in this research were completed in double-blind conditions.
Acknowledgments
This work was supported by grants from STI2030-Major Projects (2022ZD0205900, A.L.; 2021ZD0204000, W.X. and A.L.), Natural Science Foundation of China (NSFC 91632201, W.X.; NSFC 31970958, A.L.), Natural Science Foundation of Jiangsu Province (BK20211561, A.L.), Basic Research Project of Leading Technology of Jiangsu Province (BK20192004, A.L.), Shenzhen Science and Technology Innovation Foundation (2021Szvup028, A.L.), Guangdong Key Project (2018B030335001, W.X.). This protocol was adapted from the publication Ren et al. [14].
Competing interests
The authors declare that no competing interests exist.
Ethical considerations
All procedures involving mice were performed according to and approved by the Animal Care Committee at Southeast University, China.
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Ren, Q., Wang, S., Li, J., Cao, K., Zhuang, M., Wu, M., Geng, J., Jia, Z., Xie, W., Liu, A., et al. (2024). Novel social stimulation ameliorates memory deficit in Alzheimer's disease model through activating α-secretase. J Neurosci.: e1689232024.
Wu, X., Morishita, W., Beier, K. T., Heifets, B. D. and Malenka, R. C. (2021). 5-HT modulation of a medial septal circuit tunes social memory stability. Nature. 599(7883): 96–101.
Barriga, C., Martín, M. I., Tabla, R., Ortega, E. and Rodríguez, A. B. (2001). Circadian rhythm of melatonin, corticosterone and phagocytosis: effect of stress. J Pineal Res. 30(3): 180–187.
Gong, S., Miao, Y. L., Jiao, G. Z., Sun, M. J., Li, H., Lin, J., Luo, M. J. and Tan, J. H. (2015). Dynamics and Correlation of Serum Cortisol and Corticosterone under Different Physiological or Stressful Conditions in Mice. PLoS One. 10(2): e0117503.
Article Information
Publication history
Received: May 2, 2024
Accepted: Jul 3, 2024
Available online: Jul 19, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Neuroscience > Behavioral neuroscience > Cognition
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5,047 | https://bio-protocol.org/en/bpdetail?id=5047&type=0 | # Bio-Protocol Content
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Determination of Ligand-Target Interaction in vitro by Cellular Thermal Shift Assay and Isothermal Dose-response Fingerprint Assay
DD Danyu Du
SY Shengtao Yuan
JX Jing Xiong
Published: Vol 14, Iss 15, Aug 5, 2024
DOI: 10.21769/BioProtoc.5047 Views: 706
Reviewed by: Dipak Kumar PoriaSrajan KapoorGundeep Kaur
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Original Research Article:
The authors used this protocol in Oncogene Oct 2023
Abstract
The cellular thermal shift assay (CETSA) and isothermal dose-response fingerprint assay (ITDRF CETSA) have been introduced as powerful tools for investigating target engagement by measuring ligand-triggered thermodynamic stabilization of cellular target proteins. Yet, these techniques have rarely been used to evaluate the thermal stability of RNA-binding proteins (RBPs) when exposed to ligands. Here, we present an adjusted approach using CETSA and ITDRFCETSA to determine the interaction between enasidenib and RBM45. Our assay is sensitive and time-efficient and can potentially be adapted for studying the interactions of RBM45 protein with other potential candidates.
Key features
• This protocol builds upon the method developed by Molina et al. and extends its application to new protein classes, such as RBPs.
Keywords: Ligand–target interaction Cellular thermal shift assay (CETSA) Isothermal dose-response fingerprint assay (ITDRFCETSA) RNA binding motif protein 45
Graphical overview
Background
RNA-binding proteins (RBPs) contribute considerably to post-transcriptional regulation; thus, their dysregulation has been linked to a variety of illnesses, including malignancies [1]. Thereafter, targeting RBPs with specific ligands to perturb cancer development has attracted much attention [2]. Recent studies, including ours, have unraveled the role of RNA-binding motif protein 45 (RBM45) in promoting hepatocellular carcinoma (HCC) progression [3,4], highlighting its importance in cancer development. More crucially, our previous work, which combines a molecular docking approach as well as molecular and cellular techniques, identified enasidenib, a previously considered mutant isocitrate dehydrogenase 2 (mIDH2) inhibitor that binds to RBM45. Therefore, a better clarification of the binding affinity between enasidenib and RBM45 would be beneficial for the design of RBM45-targeted therapeutics.
Recently, several label-free techniques have been developed to determine ligand-target engagement using intact cells or cell lysates. These techniques include drug affinity responsive target stability (DARTS) [5], stability of proteins from rates of oxidation (SPROX) [6], and cellular thermal shift assay (CETSA) [7]. Among these, CETSA is more widely applicable since it can determine the specific ligand-induced target thermodynamic stabilization in biological settings such as whole cell lysates, intact cells, or even tissues [7]. Typically, the shift in thermal stability is assessed by quantifying the density of the remaining soluble target proteins using available antibodies via immunoblots analysis. As a result, thermal melt curves are generated, in which the target protein of interest, with or without ligands, is exposed to a panel of temperatures in the CETSA experiment. Alternatively, an isothermal dose-response curve is also generated, in which the target protein is subjected to increasing concentrations of compounds under a constant temperature (a checkpoint at which the unliganded target protein is degraded as determined by the CETSA experiment), referred to as the isothermal dose-response fingerprint assay (ITDRFCETSA) [8]. Undoubtedly, CETSA and ITDRFCETSA techniques offer the opportunity to investigate whether ligands engage their intended targets in complicated environments and determine at what concentration they exert their effects.
Moreover, the CETSA approach has been demonstrated to be effective in determining the thermal stability of enzymes such as dihydrofolate reductase (DHFR) and thymidylate synthase (TS) [7], kinases including cyclin-dependent kinases CDK4 and CDK6 [7], mitogen‐activated protein kinases p38α and ERK1/2 [8]), as well as membrane proteins like solute carriers (SLCs) [9], in the absence and presence of potential ligands. However, the CETSA technique has rarely been applied to assess the thermodynamic stabilization of RBPs when subjected to compounds. Furthermore, a cell-based CETSA decreases the sensitivity to the effects of low-affinity ligands due to drug dissociation from the target after cell lysis, making lysate-based CETSA a preferred choice [9]. Here, we report the modified cell lysate CETSA and ITDRFCETSA techniques for determining the interaction between enasidenib and RBM45. Our assay is sensitive and time saving and can potentially be adapted to investigate RBM45 protein with other candidate ligands.
Materials and reagents
Cell line
Human liver cancer cell line SK-HEP-1 was purchased from the Institute of Biochemistry and Cell Biology of the Chinese Academy of Sciences (Shanghai, China).
Materials
1.5 mL tubes (Corning Axygen, catalog number: MCT-150-C)
200 μL tubes (Corning Axygen, catalog number: PCR-02-C)
10 cm dishes (Corning Axygen, catalog number: 430167)
Reagents
MEM medium (Thermo Fisher Scientific, catalog number: 61100061)
Fetal bovine serum (FBS) (Royacel SERUM, catalog number: RYS-KF22-01)
Non-essential amino acids (100×) (Thermo Fisher Scientific, catalog number: 11140050)
Sodium pyruvate 100 mM solution (Thermo Fisher Scientific, catalog number: 11360070)
Penicillin-Streptomycin (100×) (New Cell & Molecular Biotech Co., Ltd., catalog number: C100C5)
1× phosphate buffered saline (PBS), pH 7.4 (Shanghai Zhong Qiao Xin Zhou Biotechnology Co., Ltd, catalog number: ZQ-1300)
0.25% trypsin-EDTA (New Cell & Molecular Biotech Co., Ltd., catalog number: C125C1)
RIPA lysis buffer (Beyotime Biotechnology, catalog number: P0013B)
ProtLytic protease inhibitor cocktail (EDTA-Free, 100× in DMSO) (New Cell & Molecular Biotech Co., Ltd., catalog number: P001)
Liquid nitrogen
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: 67-68-5)
Enasidenib (Jiangsu Aikon Biopharmaceutical R&D Co., Ltd., catalog number: 4409639)
Bicinchoninic acid (BCA) Protein Assay kit (enhanced) (Beyotime Biotechnology, catalog number: P0009)
Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) sample loading buffer, 5× (Beyotime Biotechnology, catalog number: P0015)
Rabbit-polyclonal-anti-RBM45 (ABclonal, catalog number: A13843)
Goat anti-Rabbit IgG (H+L) secondary antibody, HRP (Thermo Fisher Scientific, catalog number: 31460)
Enhanced chemiluminescence (ECL) detection reagent (Vazyme, catalog number: E411-03/04/05)
Equipment
Gene amplification instrument (Bioer Technology, model: G1000 GeneExplorer)
Metal Bath (Labnet)
Electrophoresis systems & transfer equipment (Bio-Rad)
Image analyzer (Tanon, model: 5200)
Refrigerated high-speed benchtop centrifuge (Thermo Fisher Scientific, model: 75005289)
Rotating incubator (Haimen Kylin-Bell Lab Instruments Co., Ltd., model: QB-228)
Software and datasets
ImageJ 1.46r
Microsoft Excel
GraphPad Prism 9.0.0
Procedure
Preparation of cell lysates for CETSA
Note: SK-HEP-1 cells were cultured in 10 cm dishes in MEM medium supplemented with 10% FBS, 1% non-essential amino acids, 1 mM sodium pyruvate, and 1% penicillin-streptomycin at 37 °C with 5% CO2 atmosphere. Cells were characterized by short tandem repeat profiling and tested for mycoplasma contamination upon receipt. Cells were split at a 1:3 ratio when they reached 80%–90% confluence.
Digest cells with 0.25% trypsin-EDTA when reaching the indicated confluence (suitable for two dishes, approximately 4 × 106 cells per dish). Transfer to two 1.5 mL tubes separately and pellet them by centrifugation (1,000× g for 5 min at room temperature).
Remove the supernatant, wash cells with cold PBS once, and collect cell pellets by centrifugation (1,000× g for 5 min at room temperature).
Resuspend cell pellets with 1 mL of RIPA lysis buffer containing protease inhibitor cocktail (1×) to each cell pellet.
Fast freeze the above-mentioned cells using liquid nitrogen and thaw (on ice). Repeat the freeze-thaw cycle three times.
Separate the soluble fractions (lysates) from the cell debris by centrifugation (20,000× g for 20 min at 4 °C) and determine the protein concentration using BCA assay kit (Protein concentration in cell lysates usually ranges between 0.1 and 2.0 mg/mL, depending on the abundance of protein expression).
Divide cell lysates evenly into two 1.5 mL tubes, incubate with enasidenib (final concentration: 30 μM) or an equivalent amount of DMSO, and rotate at room temperature for 1 h.
Divide each mixture into 100 μL aliquots in nine 200 μL tubes.
Heat enasidenib or DMSO-treated lysates at the indicated temperatures (40–70 °C) for 4 min using a G1000 GeneExplorer; then, cool the samples at room temperature for 3 min.
Note: The appropriate temperatures need to be adjusted according to the literature or pre-experiments. Here, specific settings were 40, 44, 48, 52, 57, 62, 65, 68, and 70 °C for 4 min, then switch to 25 °C for 3 min.
Collect the supernatants containing soluble fractions by centrifugation (20,000× g for 20 min at 4 °C).
Preparation of cell lysates for ITDRFCETSA
Note: For ITDRFCETSA, it is necessary to determine the temperature at which the unliganded protein starts to degrade, as determined by the CETSA experiment (i.e., the chosen temperature at which the majority of the unliganded protein is degraded and cannot be detected). The steps are nearly identical to those outlined in section A with the following exceptions:
After cell lysates are prepared following liquid nitrogen freeze-thaw and centrifugation, divide cell lysates evenly into four 1.5 mL tubes, incubate with enasidenib (final concentrations: 3, 10, and 30 μM) or DMSO, and rotate at room temperature for 1 h.
Transfer each mixture into 100 μL aliquots in four 200 μL tubes.
Heat various concentrations of enasidenib or DMSO-treated lysates on a G1000 GeneExplorer at the indicated temperature according to CETSA results (here, we chose 62 °C) for 4 min and then cool the samples at room temperature for 3 min.
Immunoblots of cell lysates for CETSA or ITDRFCETSA
Mix one volume of SDS-PAGE sample loading buffer 5× with four volumes of protein sample (here, we added 25 μL of loading buffer into 100 μL of cell lysates).
Heat and denature proteins in a metal bath at 99 °C for 5 min.
Separate the prepared protein samples (10–20 μg) on a 10% SDS-PAGE gel for immunoblots analysis. After transferring proteins from gels to the polyvinylidene fluoride (PVDF) membrane, block the membrane using a 5% milk solution. Incubate the membrane with a primary antibody (anti-RBM45) at a dilution of 1:3,000 overnight at 4 °C, followed by incubation with a secondary antibody (anti-rabbit-HRP) at a dilution of 1:5,000 for 1 h at room temperature. A chemiluminescent signal was generated with ECL detection reagent and captured using an Image analyzer.
Note: To precisely investigate whether enasidenib increases the thermal stabilization of RBM45 protein, the bands of RBM45 originated from enasidenib or DMSO-treated samples were put together upon exposure to maintain the same exposure conditions (with an exposure time ranging from 1 to 5 min).
Data analysis
For CETSA, densitometry analysis of the bands from immunoblots was performed with ImageJ. Specifically, the image background was subtracted (set the rolling ball radius to 50 pixels and choose the Light background option), regions of measurement were selected, and the grey signals of each lane were quantified. Then, the ratio of RBM45 at each indicated temperature was normalized to that at 40 °C (see Figure 1). The relative band intensity at different temperatures was plotted using GraphPad, and the thermal stability of RBM45 was measured by the temperature-dependent cellular thermal shift assay. Notably, the temperature corresponding to the protein that cannot be detected is selected to ITDRFCETSA, and the thermal stability of RBM45 was thus measured by the concentration-dependent cellular thermal shift assay at 62 °C.
Immunoblots and quantification images have been published in Oncogene [4]; for details, please refer to Figure 7e of the mentioned publication.
Figure 1. Quantitative analysis workflow for cellular thermal shift assay (CETSA)
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Du et al. [4]. RNA binding motif protein 45-mediated phosphorylation enhances protein stability of ASCT2 to promote hepatocellular carcinoma progression. Oncogene (Figure 7, panel e).
General notes and troubleshooting
In the case of CETSA, it is essential to make adjustments to the temperatures employed for a specific protein based on relevant literature or pre-experimental findings. To precisely investigate whether potential ligands enhance the thermal stabilization of the target protein, it is recommended to group together the target protein bands originating from both ligand- and vehicle-treated samples during exposure in order to maintain consistent exposure conditions and enable valid comparisons between different treatment groups.
Regarding ITDRFCETSA, the selected temperature should be modified based on the specific characteristics of the protein and ligand being studied. It is crucial to adapt the temperature based on the results obtained from CETSA experiments.
Acknowledgments
The study was supported by the National Natural Science Foundation of China (Nos. 82070883, 82273982, 81872892), the Natural Science Foundation of Jiangsu Province, China (BK20221525) and Scientific Research Foundation for high-level faculty, China Pharmaceutical University. This protocol has been previously described and validated in Oncogene [4].
Competing interests
The authors declare no competing interests.
References
Hashimoto, S. and Kishimoto, T. (2022). Roles of RNA-binding proteins in immune diseases and cancer. Semin Cancer Biol. 86: 310–324.
Kathman, S. G., Koo, S. J., Lindsey, G. L., Her, H. L., Blue, S. M., Li, H., Jaensch, S., Remsberg, J. R., Ahn, K., Yeo, G. W., et al. (2023). Remodeling oncogenic transcriptomes by small molecules targeting NONO. Nat Chem Biol. 19(7): 825–836.
Wang, C., Chen, Z., Yi, Y., Ding, Y., Xu, F., Kang, H., Lin, K., Shu, X., Zhong, Z., Zhang, Z., et al. (2023). RBM45 reprograms lipid metabolism promoting hepatocellular carcinoma via Rictor and ACSL1/ACSL4. Oncogene. 43(5): 328–340.
Du, D., Qin, M., Shi, L., Liu, C., Jiang, J., Liao, Z., Wang, H., Zhang, Z., Sun, L., Fan, H., et al. (2023). RNA binding motif protein 45-mediated phosphorylation enhances protein stability of ASCT2 to promote hepatocellular carcinoma progression. Oncogene. 42(42): 3127–3141.
Lomenick, B., Hao, R., Jonai, N., Chin, R. M., Aghajan, M., Warburton, S., Wang, J., Wu, R. P., Gomez, F., Loo, J. A., et al. (2009). Target identification using drug affinity responsive target stability (DARTS). Proc Natl Acad Sci USA. 106(51): 21984–21989.
Strickland, E. C., Geer, M. A., Tran, D. T., Adhikari, J., West, G. M., DeArmond, P. D., Xu, Y. and Fitzgerald, M. C. (2012). Thermodynamic analysis of protein-ligand binding interactions in complex biological mixtures using the stability of proteins from rates of oxidation. Nat Protoc. 8(1): 148–161.
Molina, D. M., Jafari, R., Ignatushchenko, M., Seki, T., Larsson, E. A., Dan, C., Sreekumar, L., Cao, Y. and Nordlund, P. (2013). Monitoring Drug Target Engagement in Cells and Tissues Using the Cellular Thermal Shift Assay. Science. 341(6141): 84–87.
Jafari, R., Almqvist, H., Axelsson, H., Ignatushchenko, M., Lundbäck, T., Nordlund, P. and Molina, D. M. (2014). The cellular thermal shift assay for evaluating drug target interactions in cells. Nat Protoc. 9(9): 2100–2122.
Hashimoto, M., Girardi, E., Eichner, R. and Superti-Furga, G. (2018). Detection of Chemical Engagement of Solute Carrier Proteins by a Cellular Thermal Shift Assay. ACS Chem Biol. 13(6): 1480–1486.
Article Information
Publication history
Received: Jan 12, 2024
Accepted: Jun 28, 2024
Available online: Jul 19, 2024
Published: Aug 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Molecular Biology > Protein > Protein-protein interaction
Cancer Biology > Cancer biochemistry > Protein
Drug Discovery > Drug Design
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5,048 | https://bio-protocol.org/en/bpdetail?id=5048&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Tetrazine Amino Acid Encoding for Rapid and Complete Protein Bioconjugation
AE Alex J. Eddins *
AP Abigail H. Pung *
RC Richard B. Cooley
RM Ryan A. Mehl
(*contributed equally to this work)
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5048 Views: 727
Reviewed by: Marion HoggEdwin AntonyPallavi Prasad
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Original Research Article:
The authors used this protocol in Bioconjugate Chemistry Dec 2023
Abstract
Generating protein conjugates using the bioorthogonal ligation between tetrazines and trans-cyclooctene groups avoids the need to manipulate cysteine amino acids; this ligation is rapid, site-specific, and stoichiometric and allows for labeling of proteins in complex biological environments. Here, we provide a protocol for the expression of conjugation-ready proteins at high yields in Escherichia coli with greater than 95% encoding and labeling fidelity. This protocol focuses on installing the Tet2 tetrazine amino acid using an optimized genetic code expansion (GCE) machinery system, Tet2 pAJE-E7, to direct Tet2 encoding at TAG stop codons in BL21 E. coli strains, enabling reproducible expression of Tet2-proteins that quantitatively react with trans-cyclooctene (TCO) groups within 5 min at room temperature and physiological pH. The use of the BL21 derivative B95(DE3) minimizes premature truncation byproducts caused by incomplete suppression of TAG stop codons, which makes it possible to use more diverse protein construct designs. Here, using a superfolder green fluorescent protein construct as an example protein, we describe in detail a four-day process for encoding Tet2 with yields of ~200 mg per liter of culture. Additionally, a simple and fast diagnostic gel electrophoretic mobility shift assay is described to confirm Tet2-Et encoding and reactivity. Finally, strategies are discussed to adapt the protocol to alternative proteins of interest and optimize expression yields and reactivity for that protein.
Key features
• Protocol describes site-specific encoding of the tetrazine amino acid Tet2-Et into proteins for bioorthogonal, quantitative, and rapid attachment of trans-cyclooctene-containing labels.
• Protocol uses auto-induction methods for the production Tet2-Et protein in E. coli.
• This protocol focuses on Tet-protein expressions in BL21(DE3) and B95(DE3) strains, which take approximately 4 days to complete.
• SDS-PAGE mobility shift assay using a strained TCO-PEG5000 (sTCO-PEG5000) reagent provides a simple, generalizable method for testing Tet-protein reactivity.
Keywords: 1,2,4,5-tetrazine (Tet) Noncanonical amino acid (ncAA) Genetic code expansion (GCE) Bioorthogonal Near-cognate suppression Strained trans-cyclooctene (sTCO) Aminoacyl tRNA synthetase/tRNA pair (aaRS/tRNA)
Graphical overview
Background
The facile ability to quantitatively and rapidly attach molecules onto proteins without needing to remove or modify native cysteine residues would greatly advance the study of proteins and the development of homogeneous protein reagents, therapeutics, and diagnostics. Previously, we have demonstrated that site-specific, translational encoding of tetrazine noncanonical amino acids (Tet-ncAAs) into proteins via genetic code expansion (GCE) provides the required qualities of ideal bioorthogonal reactions to advance protein labeling [1,2]. Three of these essential qualities are (1) an exceptionally fast bioorthogonal reaction that occurs under biological conditions enabling complete labeling in short reaction times, (2) a high-fidelity GCE system that ensures that all encoding sites contain the ncAA, and (3) the ncAA is stable during encoding in biological environments so that all sites are reactive. While a variety of encodable labeling strategies are available [3,4], the encoding of Tet-ncAAs into proteins and subsequent attachment of labels containing a trans-cyclooctene (TCO) functional group stands out as the only strategy that has these three qualities for routine, quantitative protein labeling. These advantages have been leveraged to produce systems for homogeneous protein conjugation to surfaces [5,6], highly effective anti-viral nanobody conjugates [2], and the attachment of spectroscopic probes both in vitro [7] and in vivo [8].
Here, we describe the GCE encoding of an ethyl-substituted 1,2,4,5-tetrazine ncAA (Tet2-Et) and its in vitro reaction with sTCO labels having a second-order rate constant of ~104 M-1·s-1 at room temperature and physiological pH (Figure 1). We use an engineered Tet2-Et RS/tRNA pair with optimized efficiency and fidelity and plasmid construct that provides high-fidelity encoding (i.e., > 95% encoding accuracy) and quantitative labeling [1]. This protocol outlines the expression of a control protein, superfolder green fluorescent protein (sfGFP) with a TAG codon at the N150 site (sfGFP150 [9]) that yields Tet-containing sfGFP (sfGFPTet2-Et) with yields of ~200 mg per liter of culture (Figure 1, Table 1). This Tet2-Et expression system requires two plasmids: (1) the pAJE-E7 GCE machinery plasmid expressing the Tet2-Et RS/tRNA pair, and (2) a plasmid that expresses a gene of interest containing the TAG stop codon at the intended site of Tet encoding. Compatible expression hosts include the standard IPTG/lactose-inducible BL21(DE3) cell line or B95(DE3) ΔAΔfabR, which minimizes premature protein truncation at TAG-encoding sites [10]. Both strains are compatible with target protein expression from classical pET vectors (e.g., pET28). Expressions are performed in auto-induction media (AIM) for reproducible, high-yielding Tet2-protein production. Finally, we describe an electrophoresis mobility shift assay using TCO-functionalized PEG5000 for quick confirmation of accurate Tet encoding into proteins, the stability of TCO reagents, and the efficiency of protein conjugation. Completion of this protocol, including the confirmation of Tet2-Et encoding into sfGFP, takes approximately four days. Discussions on adapting this protocol for encoding Tet2-Et into biologically relevant proteins are provided.
Figure 1. Genetic code expansion (GCE) encoding of tetrazine noncanonical amino acids (Tet-ncAA) and quantitative reaction with trans-cyclooctene (TCO) labels. (A) Structure of the Tet2-Et ncAA in its oxidized, reactive state. (B) During protein expression, sfGFP150 with encoded Tet2-Et exists in an equilibrium between an unreactive reduced state (green, top left) and the reactive oxidized state (orange, top right). The orange color of reactive sfGFPTet2-Et results from Tet2 quenching of sfGFP fluorescence when Tet2-Et is encoded at site N150. This fluorescence quenching does not occur when Tet2-Et is in the reduced form. Upon purification, buffer exchange, and exposure to ambient oxygen, any reduced (green) Tet2-sfGFP protein quickly oxidizes and can be quantitatively reacted with the desired sTCO labeling reagent. After reaction with sTCO, Tet2-Et no longer quenches sfGFP fluorescence. See General notes 1 and 2 for more information on Tet2 reactions and redox properties, respectively.
Materials and reagents
Biological materials
Strains
BL21(DE3) (Thermo Fisher, catalog number: EC0114). This strain of E. coli is optimized for over-expression of target proteins under the T7 transcriptional promoter that is commonly found in standard pET vectors. This strain contains a genomic copy of the T7 RNA polymerase controlled by the lacUV5 promoter so that upon introduction with IPTG, the T7 polymerase is expressed and transcribes the target gene to produce high quantities of recombinant target protein. This strain contains Release Factor 1 (RF1), the protein responsible for terminating translation at TAG amber stop codons so that when encoding Tet2-Et at TAG codons of target proteins, truncated protein will be produced along with full-length Tet2-protein. To avoid co-purification of truncated protein with full-length protein, C-terminal purification tags are recommended. For proteins that self-assemble into homo-multimers (dimers, trimers, etc.), purification can be challenging due to the possible co-purification of truncated forms that are incorporated as subunits in the assembly. If using N-terminal purification/solubilization tags, additional purification steps may be needed to remove truncated protein species. See Troubleshooting #1 for more discussion.
B95(DE3) ΔAΔfabR (Addgene, catalog number: 197655). This strain is a robustly growing BL21(DE3) derivative that lacks release factor-1 (RF1), the protein responsible for translation termination at TAG codons, as well as a spontaneously mutated fabR gene [10]. Endogenous TAG stop codons in 95 genes were mutated to TAA or TGA to maintain cellular health and minimize unwanted readthrough due to RF1 knockout. This strain is preferred over BL21(DE3) for protein expression because by lacking RF1, TAG codon suppression by GCE machinery is more efficient, and the production of truncated protein (caused by early TAG site termination) is minimized. B95(DE3) ΔAΔfabR cells require lower concentrations of antibiotics to maintain normal growth rates.
DH10B (Thermo Fisher, catalog number: EC0113). This strain can be used for faithful propagation of plasmids and for cloning needs when users clone their genes of interest into their preferred plasmid backbone. Do not use this strain for protein expression. Though we have not explicitly tested all of them, other classical cloning strains of E. coli can be used in place of DH10b, including NEB 10-beta (New England BioLabs, catalog number: C3019H), DH5α (e.g., Thermo Fisher, catalog number: 18258012), NEB 5-alpha (New England BioLabs, catalog number: C2987H), or TOP10 (Thermo Fisher, catalog number: C404010).
Plasmids
pAJE3-E7 (Addgene, catalog number: 214359). Machinery plasmid for Tet2-Et incorporation that expresses a copy of the Methanocaldococcus jannaschii (Mj)-TyrRS-(E7)-Tet2-Et RS for faithful Tet2-Et incorporation as well as a copy of its cognate amber codon suppressing tRNA under constitutive lpp promoters. This plasmid confers spectinomycin resistance and harbors a recently developed high-copy synthetic origin of replication [1,11]. This synthetic origin of replication in pAJE plasmids can be stably propagated in cells that contain other plasmids having any of the standard or typical origins of replications, such as ColE1/pBR322/pMB1, p15A, and CDF origins, adding to the utility and versatility of this machinery plasmid.
pET28-sfGFPWT (Addgene, catalog number: 85492). Expresses wild-type sfGFP control protein with C-terminal His6 tag, under a T7 transcriptional promoter, kanamycin resistance, and pBR322 origin of replication. sfGFP is expressed by the addition of IPTG or lactose.
pET28-sfGFP150 (Addgene, catalog number: 85493). Same as above except the sfGFP gene contains a TAG amber stop codon at site N150. The TAG codon is used to direct the translational encoding of Tet2-Et.
pET28-[GOI]WT (you must create). Expresses your wild-type protein of interest (POI). You can clone your POI into pET28 by removing the sfGFP gene via restriction digest with NcoI and XhoI enzymes and replacing it with your gene of interest (GOI) using standard cloning techniques (e.g., ligation, Gibson Assembly, or SLiCE [12]). For the reasons mentioned above, a C-terminal purification tag is preferred if using the BL21(DE3) strain for expression. For helpful tips on construct design, see Troubleshooting tip 1.
pET28-[GOI]TAG (you must create). Expresses your POI with Tet2-Et encoded at a TAG codon. You can clone your POI into pET28 backbone by removing the sfGFP gene by restriction digest with NcoI and XhoI and replacing it with your gene of interest (GOI) using standard cloning techniques (e.g., ligation, Gibson Assembly, or SLiCE). Using site-directed mutagenesis, change the codon to TAG (the amber stop codon) where you intend to encode Tet2-Et into your POI. See section B for recommendations on TAG site selection.
Reagents
Essential reagents
Tryptone (e.g., VWR, catalog number: 97063-386)
Yeast extract (e.g., VWR, catalog number: 97064-368)
NaCl (e.g., VWR, catalog number: 97061-274)
α-D-glucose (e.g., VWR, catalog number: 97061-168)
α-lactose (e.g., VWR, catalog number: AAA11074-0B)
Glycerol (e.g., VWR, catalog number: BDH24388.320)
Na2HPO4 (sodium phosphate dibasic) (e.g., VWR, catalog number: 97061-586)
KH2PO4 (potassium monobasic) (e.g., VWR, catalog number: BDH9268)
K2HPO4 (potassium dibasic) (e.g., VWR, catalog number: 97062-234)
NH4Cl (e.g., VWR, catalog number: 12125-02-9)
(NH4)2SO4 (e.g., VWR, catalog number: 7783-20-2)
Na2SO4 (e.g., VWR, catalog number: 7757-82-6)
MgSO4 (e.g., VWR, catalog number: 7487-88-9)
CaCl2ᐧ2H2O (e.g., VWR, catalog number: 10035-04-8)
MnCl2ᐧ4H2O (e.g., VWR, catalog number: 13446-34-9)
ZnSO4ᐧ7H2O (e.g., VWR, catalog number: 7446-20-0)
CoCl2ᐧ6H2O (e.g., VWR, catalog number: 7791-13-1)
CuCl2 (e.g., VWR, catalog number: 10125-13-0)
NiCl2 (e.g., VWR, catalog number: 7791-20-0)
Na2MoO4ᐧ2H2O (e.g., VWR, catalog number: 10102-40-6)
Na2SeO3 (e.g., VWR, catalog number: 10102-18-8)
H3BO3 (e.g., VWR, catalog number: 10043-35-3)
FeCl3 (e.g., VWR, catalog number: 7705-08-0)
L-arabinose (e.g., VWR, catalog number: 5328-37-0)
Kanamycin (e.g., VWR, catalog number: 75856-684)
Spectinomycin (e.g., VWR, catalog number: 89156-368)
Isopropyl β-D-1-thiogalactopyranoside (IPTG) (e.g., Anatrace, catalog number: I1003)
Agar (e.g., VWR, catalog number: 97064-336)
Tet2-Et amino acid (see General note 4 for reagent availability)
N,N-Dimethylformamide (DMF) (VWR, catalog number: BDH1117-4LG)
sTCO-PEG5000 (see General note 4 for reagent availability)
Sodium dodecyl sulfate (SDS) (VWR, catalog number: JT4095-4)
Silicone emulsion, Antifoam B® (VRW, catalog number: JTB531-5)
Reagents necessary only for defined auto-induction media
Glutamic acid, Na salt (e.g., VWR, catalog number: 56-86-0)
Aspartic acid (e.g., VWR, catalog number: 56-84-8)
Lysine-HCl (e.g., VWR, catalog number: 657-27-2)
Arginine-HCl (e.g., VWR, catalog number: 1119-34-2)
Histidine-HCl-H2O (e.g., VWR, catalog number: 5934-29-2)
Alanine (e.g., VWR, catalog number: 56-41-7)
Proline (e.g., VWR, catalog number: 147-85-3)
Glycine (e.g., VWR, catalog number: 56-40-6)
Threonine (e.g., VWR, catalog number: 72-19-5)
Serine (e.g., VWR, catalog number: 56-45-1)
Glutamine (e.g., VWR, catalog number: 56-85-9)
Asparagine-H2O (e.g., VWR, catalog number: 5794-13-8)
Valine (e.g., VWR, catalog number: 72-18-4)
Leucine (e.g., VWR, catalog number: 61-90-5)
Isoleucine (e.g., VWR, catalog number: 73-32-5)
Phenylalanine (e.g., VWR, catalog number: 63-91-2)
Tryptophan (e.g., VWR, catalog number: 73-22-3)
Methionine (e.g., VWR, catalog number: 63-68-3)
Solutions
Essential solutions
LB/agar media (see Recipes)
2× YT media (see Recipes)
SOC media (see Recipes)
Kanamycin stock (see Recipes)
Spectinomycin stock (see Recipes)
1 M IPTG (see Recipes)
25× M salts (see Recipes)
Trace metal stock solution (5,000×) (see Recipes)
50× 5052 solution (see Recipes)
Tet2-Et solution (see Recipes)
ZY media (see Recipes)
ZY non-induction (ZY-NIM) and auto-induction media (ZY-AIM) (see Recipes)
Solutions necessary only for defined auto-induction media
Aspartate [5% (w/v), pH 7.5] (see Recipes)
18 amino acid mix (25×) (see Recipes)
Defined-NIM and AIM media (see Recipes)
Recipes
LB/agar media (0.5 L)
Reagent Final concentration Amount
Tryptone 1 % (w/v) 5 g
Yeast extract 0.5 % (w/v) 2.5 g
NaCl 1.0 % (w/v) 5 g
Agar 1.5 % (w/v) 7.5 g
H2O n/a To 500 mL
Total n/a 500 mL
After mixing reagents thoroughly, autoclave on standard liquid setting to sterilize. Note the agar will not go into solution until autoclaved.
After autoclaving, gently swirl the bottle to ensure molten agar is evenly mixed.
Notes:
i. Store LB/agar bottle in a 55 °C oven and pour plates on an as-needed basis. LB/agar can be stored in molten form for ~2 weeks if sterility is maintained.
ii. If an oven is not available, plates can be poured with antibiotics once LB/agar is sufficiently cooled to touch. Plates can be stored at 4 °C for up to a week.
2× YT media (1 L)
Reagent Final concentration Amount
Tryptone 1.6 % (w/v) 16 g
Yeast extract 1.0 % (w/v) 10 g
NaCl 0.5 % (w/v) 5 g
H2O n/a To 1000 mL
Total n/a 1 L
After mixing reagents thoroughly, autoclave on the standard liquid setting to sterilize.
After autoclaving, allow it to cool to room temperature before use.
SOC media (50 mL)
Reagent Final concentration Amount
2× YT media n/a 49 mL
1 M MgSO4 10 mM 0.5 mL
40% (w/v) α-D-glucose 0.4% (w/v) or ~20 mM 0.5 mL
Total n/a 50 mL
1 M MgSO4 can be made by mixing 12.3 g of MgSO4·7H2O in water up to 50 mL total volume. Adjust mass of MgSO4 accordingly if using a salt with a different hydration status.
40% (w/v) α-D-glucose can be made by mixing 20 g of α-D-glucose with water up to 50 mL total volume. Mix thoroughly until glucose is dissolved. Gentle heating in a microwave may facilitate the dissolution of glucose.
Sterilize MgSO4, glucose, and 2× YT solutions individually by autoclaving. Allow each component to cool to room temperature and mix as indicated above. Maintain sterility while adding components together.
It is easy to contaminate SOC. We suggest breaking this into 5 × 10 mL aliquots before use or making smaller batches. If sterility is maintained, SOC can be stored at room temperature indefinitely. It can also be stored at -20 °C but avoid repeated freeze/thaws.
Kanamycin stock (10 mL)
Reagent Final concentration Amount
Kanamycin 50 mg/mL 0.5 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Spectinomycin stock (10 mL)
Reagent Final concentration Amount
Spectinomycin 100 mg/mL 1 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
Note: Do not confuse spectinomycin with streptomycin. These antibiotics are not interchangeable.
1 M IPTG (10 mL)
Reagent Final concentration Amount
IPTG 1 M 2.3 g
H2O n/a To 10 mL
Total n/a 10 mL
Sterilize by filtering with a 0.2 μm syringe-end filter.
Store in 1 mL aliquots at -20 °C.
25× M salts
Reagent 25× concentration Amount for 25×
Na2HPO4 0.625 M 88.7 g
KH2PO4 0.625 M 85.1 g
NH4Cl 1.25 M 66.9 g
Na2SO4 0.125 M 17.8 g
H2O n/a to 1 L
Total 1 L
Add the above components to a 2 L beaker containing a magnetic stir bar. Add water up to 900 mL and mix until all components have dissolved. Add the remaining volume of water to reach 1 L.
Weights indicated are based on anhydrous salts. If using hydrated phosphate salts, adjust the weights accordingly to maintain indicated molarities.
Trace metal stock solution (5,000×)
Reagent Concentration Amount for individual 30 mL stocks
5,000× 1×
CaCl2·2H2O 20 mM 4 µM 8.82 g
MnCl2·4H2O 10 mM 2 µM 5.93 g
ZnSO4·7H2O 2 M 2 µM 8.62 g
CoCl2·6H2O 2 mM 0.4 µM 1.32 g
CuCl2 2 mM 0.4 µM 807 mg
NiCl2 2 mM 0.4 µM 777 mg
Na2SeO3 2 mM 0.4 µM 1.03 g
Na2MoO4·2H2O 2 mM 0.4 µM 1.45 g
H3BO3 2 mM 0.4 µM 371 mg
FeCl3 50 mM 10 µM 486 mg
H2O n/a To 30 mL
For each of the metals above (except FeCl3), make individual stock solutions using the indicated masses and dissolve in Milli-Q water up to 30 mL of total volume. Autoclave each metal solution separately to sterilize. The FeCl3 must be dissolved in 0.1 M HCl up to 30 mL of total volume and then filtered (through a 0.2 µm filter) to remove insoluble material and sterilize (do not autoclave).
Once all individual stock solutions are prepared, add 500 µL of each stock solution (except FeCl3) to 20.5 mL of sterile Milli-Q water. Then, add 25 mL of the FeCl3 solution. The total volume should be exactly 50 mL.
This stock solution might show minor precipitation over time but is stable at 15–25 °C for years.
50× 5052 solution (500 mL)
Reagent Final concentration Amount
α-D-glucose 25 mg/mL 12.5 g
α-lactose 100 mg/mL 50 g
Glycerol 25% (v/v) 125 mL
H2O n/a to 500 mL
Total n/a 500 mL
Add the glucose, lactose, and glycerol components to roughly 300 mL of warm water in a 0.5 L beaker containing a magnetic stir bar. Mix until all solutions have dissolved. Additional heating may be required via microwave to encourage lactose dissolution (CAUTION: Remove magnetic stir bar before microwaving). Once fully dissolved, add the remaining volume of water to reach 500 mL.
Autoclave on liquid cycle to sterilize.
Tet2-Et solution
Reagent Final concentration Amount
Tet2-Et 100 mM 8.5 mg
DMF n/a to 275 µL
Total n/a 275 µL
Vortex solution after combining to ensure all Tet2-Et has dissolved.
Solution can be stored at -20 °C for months but may come out of solution and require additional vortexing upon freeze/thaw cycles. For optimal expressions, prepare the solution directly before use to avoid freeze/thaw cycles.
ZY media
Reagent Final concentration Amount
Tryptone 1% (w/v) 10 g
Yeast extract 0.5% (w/v) 5 g
H2O n/a to 1 L
Total n/a 1 L
Add the above components to a 1 L beaker containing a magnetic stir bar. Add water up to 900 mL and mix until all solutions have dissolved. Add the remaining volume of water to reach 1 L.
Autoclave for sterilization.
ZY non-induction (ZY-NIM) and auto-induction media (ZY-AIM)
Reagent ZY-NIM ZY-AIM
Amount Amount
ZY media 47 mL 47 mL
MgSO4 0.1 mL 0.1 mL
25× M salts 2 mL 2 mL
50× 5052 - 1 mL
40% (w/v) α-D-glucose 0.625 mL -
Trace metal (5,000×) 10 µL 10 µL
Total 50 mL 50 mL
When preparing media, dilute the concentrated components into ZY media. Do not mix concentrated stocks and then dilute with ZY media.
For BL21(DE3), the final concentration for spectinomycin and kanamycin should be 100 µg/mL and 50 µg/mL, respectively. For B95(DE3) expressions, the final concentration for spectinomycin and kanamycin should be 50 µg/mL and 25 µg/mL, respectively.
Prepare immediately before use with a sterile technique.
Aspartate [5% (w/v), pH 7.5]
Reagent Final concentration Amount
Aspartate 5% (w/v) 50 g
H2O n/a To 1 L
Total n/a 1 L
Mix by placing a suitable magnetic stir bar in a 2 L beaker and add 900 mL of water to the graduated cylinder. While stirring, add the appropriate amount of L-aspartic acid and adjust pH to 7.5 with 8 M NaOH. Add the remaining volume of H2O to bring the solution to the final volume of 1 L.
Sterilize by autoclaving on liquid setting.
18 amino acid mix (25×) (1 L)
Reagent Concentration Amount for 25×
25× 1×
Glutamic acid, Na salt 200 µg/mL 8 µg/mL 5 g
Aspartic acid 200 µg/mL 8 µg/mL 5 g
Lysine-HCl 200 µg/mL 8 µg/mL 5 g
Arginine-HCl 200 µg/mL 8 µg/mL 5 g
Histidine-HCl-H2O 200 µg/mL 8 µg/mL 5 g
Alanine 200 µg/mL 8 µg/mL 5 g
Proline 200 µg/mL 8 µg/mL 5 g
Glycine 200 µg/mL 8 µg/mL 5 g
Threonine 200 µg/mL 8 µg/mL 5 g
Serine 200 µg/mL 8 µg/mL 5 g
Glutamine 200 µg/mL 8 µg/mL 5 g
Asparagine-H2O 200 µg/mL 8 µg/mL 5 g
Valine 200 µg/mL 8 µg/mL 5 g
Leucine 200 µg/mL 8 µg/mL 5 g
Isoleucine 200 µg/mL 8 µg/mL 5 g
Phenylalanine 200 µg/mL 8 µg/mL 5 g
Tryptophan 200 µg/mL 8 µg/mL 5 g
Methionine 200 µg/mL 8 µg/mL 5 g
H2O n/a n/a To 1 L
Total n/a n/a 1 L
Add 800 mL of water to a 1 L beaker, then add 5 g of each amino acid while stirring with a magnetic stir bar. Since some amino acids have trouble dissolving in solution, warming the water prior to adding the amino acids can aid in the dissolution process. It may take several hours for each component to fully dissolve. Finally, bring the volume to 1 L with water.
Sterilize by filtration.
Aliquot 45 mL of 25× 18-amino acid mix into sterile 50 mL conicals.
Store aliquots at -20 °C. Thaw working aliquot as needed, which can be stored stably at 4 °C for several months provided sterility is maintained.
Defined-NIM and AIM media
Reagent NIM AIM
Amount Amount
Aspartate [5% (w/v) pH 7.5] 2.5 mL 2.5 mL
50× 5052 - 1 mL
18 amino acid mix 2.0 mL 2.0 mL
25× M salts 2.0 mL 2.0 mL
MgSO4 (1 M) 100 µL 100 µL
Glucose [40% (w/v)] 6.25 mL -
Trace metal solution (5,000×) 10 µL 10 µL
Sterile H2O 36.64 mL 41.89 mL
Total 50 mL 50 mL
When preparing media, add the concentrated components to sterile H2O, do not mix concentrated stocks, and then dilute with sterile H2O.
For BL21(DE3), the final concentration for spectinomycin and kanamycin should be 100 µg/mL and 50 µg/mL, respectively. For B95, the final concentration for spectinomycin and kanamycin should be 50 µg/mL and 25 µg/mL, respectively.
Prepare immediately before use with a sterile technique.
Laboratory supplies
1.7 mL Eppendorf tubes (e.g., VWR, catalog number: 87003-294)
100 mm plates (e.g., VWR, catalog number: 470210-568)
500 mL graduated cylinder (e.g., VWR, catalog number: 470344-338)
15 mL conical tubes (e.g., VWR, catalog number: 89126-798)
50 mL conical tubes (e.g., VWR, catalog number: 89039-656)
14 mL sterile culture tubes (e.g., VWR, catalog number: 60818-689)
250 mL baffled flasks (e.g., VWR, catalog number: 89095-266)
Micro pipette tips 10 µL (e.g., VWR, catalog number: 76323-394)
Micro pipette tips 200 µL (e.g., VWR, catalog number: 76323-390)
Micro pipette tips 1,000 µL (e.g., VWR, catalog number: 76323-454)
12% Mini-PROTEAN® TGXTM precast protein gels, 15 well, 15 µL (Bio-Rad, catalog number: 4561046)
Mini-PROTEAN® Tetra companion running module (Bio-Rad, catalog number: 1658038)
Mini-PROTEAN® Tetra vertical electrophoresis cell for mini precast gels, 4-gel (Bio-Rad, catalog number: 1658004)
Disposable PD-10 desalting column, with Sephadex G-25 resin, 1.0–2.5 mL samples (Cytiva, catalog number: 17085101)
TALON® SuperflowTM (VWR, catalog number: CA71006-006)
Nalgene® bottle-top sterile filter (Millipore Sigma, catalog number: Z358223-12EA)
Equipment
Autoclave capable of sterilizing liquid media and culturing materials at 121 °C and of a saturated steam pressure of 15 PSI
Expression equipment:
Static incubator for growing LB/agar plates (set to 37 °C) (e.g., VWR, catalog number: 97025-630)
Shaker incubator for growing liquid cultures (e.g., New Brunswick I26R, Eppendorf, catalog number: M1324-0004)
i. Shaker should be able to rotate at 200–250 rpm.
ii. Refrigeration is necessary for expressions below room temperature (< 25 °C).
iii. Shaker deck should have clamps to hold 250 mL and 2.8 L Fernbach flasks.
Optical density 600 nm spectrophotometer (e.g., Ultrospec 10, Biochrome, catalog number: 80-2116-30)
Fluorometer capable of reading sfGFP fluorescence (excitation 488 nm/emission 512 nm). Handheld fluorometers work well for routine fluorescence reads (e.g., PicoFluor from Turner Biosystems)
Freezer (-20 °C) for storing plasmids and antibiotics (e.g., Fisher Scientific, catalog number: 10-549-264)
Ice machine (e.g., Fisher Scientific, catalog number: 09-540-003)
Software and datasets
ImageJ
Procedure
Overview
In Part A, we discuss practical considerations for which strain to use for the expression of Tet2-protein and best strategies for preparing competent cells. In Part B, general guidelines for selecting TAG sites for your unique gene of interest are discussed. In Part C, the day-by-day steps for Tet-protein expressions at a 50 mL scale in BL21(DE3) and B95 cell lines are described in detail. Variations on autoinduction media (defined-AIM vs. ZY-AIM) and considerations when scaling up expressions are discussed where relevant. Part C also describes the evaluation of Tet2-protein reactivity after purification using a gel mobility shift assay.
Expression host and competent cell preparation considerations
Choice of expression strain. As discussed briefly above, we recommend (and this protocol is written for) using either the BL21(DE3) or B95(DE3) ΔAΔfabR expression strains. Of these two, the latter RF1-deficient strain may be preferred as it limits prematurely truncated protein at TAG codons where Tet2-Et is intended to be encoded, thus increasing overall Tet2-protein yields while also allowing purification of target proteins with N-terminal purification/solubility tags without co-purification of truncated protein. Although we have not tested all other options, we expect that alternative T7-based expression strains of E. coli such as Rosetta(DE3), pLysS(DE3), and C41/43(DE3) are compatible with this particular Tet2-Et encoding strategy. T7Express strains from New England Biolabs (NEB) are not compatible with AIM and, therefore, methods described here will not work for these strains. We have not yet been successful at adopting Tet2-Et encoding in Origami(DE3) or Shuffle T7 strains (unpublished data).
Tips for generating competent cells. Expression cultures tend to be the most reproducible when multiple colonies are used to inoculate starter cultures. Thus, BL21(DE3) or B95(DE3) ΔAΔfabR cells need to be made sufficiently competent to transform two plasmids at once and obtain at least several dozen transformants (or colony-forming units). Note that fresh double plasmid transformations must be performed for each expression; BL21(DE3) cells (and their derivatives) should never be frozen as glycerol stocks with plasmids in them for later expressions. If cells are frozen for storage with plasmids in them, the cells will grow with the necessary antibiotics giving the false impression that they are suitable for expressions, but they will not reliably produce target protein.
Two types of competent cells can be made: chemically competent and electrocompetent. Chemically competent cells are the less efficient option of the two, but when made properly, they are sufficiently competent to generate hundreds of colonies from a double plasmid transformation using the pAJE3-E7 and pET28-[GOI] plasmids. The advantages of chemically competent cells are that they do not require special electroporation equipment and can be prepared less frequently if users are expecting to conduct many expressions. We recommend users follow the so-called “Inoue” method when generating chemically competent cells [13]. Chemically competent cells are not recommended if triple plasmid transformations are required, as seen in Eddins et al. [1] when additional accessory plasmids were used (e.g., for expressing protein folding chaperones). In these cases, electrocompetent cells can be prepared and used with an electroporator to greatly improve the efficiency of transformation and the number of colony-forming units [14]. This protocol is written for the preferred chemical transformations. For a detailed electrocompetent cell preparation protocol, see Zhu et al. [14]. Aliquots of competent cells are stable for at least two years at -80 °C without loss of competency, provided they do not experience notable temperature fluctuations or thaw.
Selecting TAG sites to screen for encoding
When optimizing this protocol for your GOI, it is important to screen multiple TAG sites, as some protein locations are more amenable to alteration than others and some TAG sites suppress more efficiently, both of which can affect protein expression and stability. When possible, structural information of the target protein should be used to guide the placement of TAG codons so that Tet2 incorporation does not perturb protein structure or function. We find it is typically best to install Tet2 at solvent-exposed sites, within flexible loops, or residues that do not make interactions important for protein stability. Yet even with such a priori information, the ideal placement of Tet2 installation is not easily predictable, and screening multiple sites in parallel (~3–6) is often the best practice to determine the sites for efficient encoding and downstream applications without affecting protein function. For our control protein, sfGFP, we have screened for efficient encoding sites like the N150 site used in this protocol. See Figure 2 for examples of two sites that allow for high expression of sfGFPTet2-Et that follow the described guidelines.
Figure 2. Superfolder GFP (PDB ID: 2B3P) as a model for TAG site screening. Examples of successfully encoding TAG sites N150 and D134 are highlighted. These sites adhere to TAG site placement guidelines: both sites are solvent-exposed and do not engage in structurally critical interactions, while site D134 is located within a flexible loop region.
Expression of Tet2-protein in BL21(DE3) and B95(DE3) cell lines
Note 1: Volumes of media can be changed depending on the scale of expression desired. Described below are volumes for a 50 mL sfGFP test expression.
Note 2: Reproducible expressions via auto-induction methods benefit from overnight non-inducing starter cultures that reach the stationary phase (total growth time ~12–18 h). AIM cultures inoculated with non-inducing cultures that did not reach the stationary phase may not always express the target protein appropriately in AIM.
Day 1: Transformations
Prepare two LB/agar plates, one for sfGFPWT and one for sfGFP150 expressions, with antibiotics, as follows:
i. Sterilize LB/agar as described above. After autoclaving, mix the contents of the bottle and allow the bottle to cool sufficiently to touch while the agar still remains liquid.
ii. Pour 50 mL of LB/agar into a sterile 50 mL conical tube. Add 50 μL each of spectinomycin and kanamycin stock solutions for BL21(DE3) cells. For B95 cells, add 25 μL of each. Mix thoroughly and pour ~15–20 mL into each 100 mm plate. The final working concentrations of antibiotics for BL21(DE3) expressions should be 100 μg/mL spectinomycin (for the pAJE plasmid) and 50 µg/mL kanamycin (for the pET28 plasmid). The final working concentrations of antibiotics for BL21(DE3) expressions should be 50 µg/mL spectinomycin and 25 µg/mL kanamycin for BL21(DE3) or B95 cells, respectively.
iii. Allow LB/agar to cool and solidify beside a flame with the plate lid slightly ajar for ~30 min.
Label the plates accordingly, e.g., “pAJE3 + pET28-sfGFPWT” and “pAJE3 + pET28-sfGFP150”.
For each expression, label two 1.7 mL Eppendorf tubes (e.g., “pAJE3-E7 + sfGFPWT” and “pAJE3-E7 + sfGFP150”).
Add 1 µL of pAJE plasmid (~200–400 ng) and 2 µL of pET28-sfGFPWT (~100 ng) to one tube. To the other tube, add 1 µL of pAJE plasmid and 2 µL of pET28-sfGFP150.
Place both tubes containing plasmids on ice for 5 min to pre-chill.
Thaw aliquots of chemically competent cells [BL21(DE3) and/or B95(DE3)] and place on ice once thawed. Cells can be thawed rapidly with the warmth of your fingers, but immediately place the tube on ice once thawed.
Add 50 µL of competent cells to each tube with plasmids, gently mix cells with plasmids by briefly pipetting up and down or flicking gently, and place back on ice for 30 min. Do not vortex cells.
Heat shock cells by submerging the end of the Eppendorf tube in a 42 °C water bath for exactly 45 s.
Immediately place the tubes back on ice and incubate for 2 min.
Add 1 mL of SOC media.
Note: Make sure SOC is not contaminated from prior use.
Allow cells to recover at 37 °C with shaking at >200 rpm. It is convenient to simply tape Eppendorf tubes horizontally to the shaker deck.
i. For BL21(DE3) cells, recover for 90 min.
ii. For B95(DE3) cells, recover for 120 min.
Plate recovered culture onto LB/agar plates with appropriate antibiotics for the strain used.
i. To ensure a sufficient number of colonies, plate all cells. To do this, centrifuge Eppendorf tubes at 3,000× g for 3 min, remove 900 µL of the supernatant, resuspend the cell pellet by gentle pipetting in the remaining 100 µL, then plate and spread the fully resuspended 100 µL of cells.
ii. Let plate(s) dry with lid partially open for ~20 min near a flame (maintaining sterility) and then incubate the plate upside down overnight at 37 °C.
Day 2: Non-inducing starter cultures
Remove the LB/agar plates from the 37 °C incubator and place at room temperature or 4 °C for the day.
Note: Several hundred colonies should be obtained for BL21(DE3) co-transformations while B95(DE3) transformation should have several dozen; see Figure 3.
Figure 3. Example LB/agar transformation plates. Transformations of B95(DE3) cells (left) and BL21(DE3) cells (right) commonly produce ~100 colonies and ~1,000 colonies, respectively, after 20 h of growth at 37 °C.
At the end of the day (~3–5 pm), prepare starter cultures.
Note: Defined-NIM and ZY-NIM have separate recipes as listed above. Here, researchers can choose between using defined-NIM and ZY-NIM, depending on their required expression conditions. These tetrazine GCE systems were developed and optimized using defined media because this media offers high reproducibility in expression yields and tet-encoding fidelity from batch to batch. However, the number of reagents required and the time to assemble them into defined media can be cumbersome. ZY-based media is a great alternative to defined media because it is easier to make and requires fewer reagents to assemble; however, depending on the source of tryptone and yeast extract, expression yields may vary slightly between batches. As shown in Figures 4 and 5 and Table 1, we generally see comparable Tet-protein expression yields and fidelity using ZY-based vs. defined media.
i. Prepare 50 mL of NIM with the appropriate antibiotic concentrations. See Recipes for details on making defined-NIM and ZY-NIM.
ii. Prepare starter cultures for sfGFPWT and sfGFPTet2-Et expressions:
1) Label two 15 mL sterile culture tubes with the plasmid combination (i.e., pAJE3-E7 + pET28-sfGFPWT and pAJE3-E7 + pET28-sfGFP150) and add NIM (5 mL) to each tube.
2) To inoculate these 5 mL cultures, scrape a glob of cells constituting several dozen colonies from overnight LB/agar plate with a sterile pipette tip, shake the glob off into the culture media, and break apart by gentle pipetting. Enough cells should be transferred to the 5 mL starter culture such that it is slightly turbid upon inoculation.
Note: Since expression levels can vary across different BL21(DE3) clones, we recommend inoculating starter cultures with several dozen colonies to obtain an averaged population for highly reproducible results from one expression to another.
3) Grow starter cultures at 37 °C with shaking at 250 rpm overnight.
Figure 4. Example BL21(DE3) defined-AIM expressions displaying the characteristic green color of sfGFPWT (left) and orange color of sfGFPTet2-Et (right). Cultures expressing pET28-sfGFP150 and lacking Tet2-Et amino acid should be colorless since no full-length sfGFP is expressed (middle).
Figure 5. Expected sfGFP fluorescence values of 50 mL expression cultures for (A) sfGFPWT and (B) sfGFPTet2-Et. By plotting the normalized fluorescence of each expression (i.e., the raw fluorescence values divided by the OD600), relative yields per cell can be estimated for each expression. It is important to remember that oxidized encoded Tet2-Et quenches sfGFP’s fluorescence and so these values do not necessarily reflect the actual yield of sfGFPTet2-Et protein. sfGFPTet2-Et normalized fluorescence values shown in this representative expression should give approximate fluorescence values to expect for control sfGFP expressions. B95(DE3) expressions lacking Tet2-Et ncAA display expected high normalized fluorescence values due to near-cognate suppression [1]. These expressions were performed in duplicate and allowed to express for 20 h.
Table 1. Characteristic yields (milligrams per liter of culture) for sfGFPWT and sfGFPTet2-Et from purified 50 mL expressions using 500 mL of TALON affinity resin calculated using absorbance values at A280
Media condition Cell line sfGFP variant Yield (mg/L per 500 μL TALON resin)
Defined-AIM BL21(DE3) WT 250
sfGFPTet2-Et 230
B95(DE3) ΔAΔfabR sfGFPWT 210
sfGFPTet2-Et 160
ZY-AIM BL21(DE3) sfGFPWT 240
sfGFPTet2-Et 230
B95(DE3) ΔAΔfabR sfGFPWT 170
sfGFPTet2-Et 160
Day 3: Expressions
Note 1: Expression cultures should have a starting OD of ~0.05 after inoculation.
Note 2: A 50 mL expression is best grown in a 250 mL baffled flask. It is important to use baffled flasks as they increase the aeration of expressions, promoting proper growth of E. coli and higher protein expression levels. Increased aeration, in turn, increases the formation of foam, making it necessary to use antifoam in expression cultures.
Note 3: When scaling for larger expressions, consider the size of the baffled flask in reference to the volume of media used. We recommend a total flask volume that is ~3–5 times the volume of the media to ensure adequate aeration, e.g., a 2 L flask works well with 500 mL of media.
Note 4: We recommend mixing all media stock components together in a single batch and then dividing working media into appropriate 250 mL baffled culture flasks to ensure all expressions contain the same media. Below, we perform 3 × 50 mL expressions: one for sfGFPWT, one for sfGFP150 with Tet2-Et in the media, and one for sfGFP150 lacking Tet2-Et in the media. In this case, prepare 150 mL of AIM and split into three flasks.
Preparing and inoculating auto-induction expressions
i. Measure the OD600 of starting cultures after overnight growth. OD600 readings are a measurement of culture density through light scattering and transmittance of a given culture—optical density at a 600 nm wavelength. This measurement provides an easy assessment of the E. coli culture’s growth phase, allows inoculation of expression cultures at equal densities, and provides normalization of protein production by cell density. OD600 for NIM is generally low and is expected to be between 1.5 and 4, depending on the culture vessel, media type used, and metabolic burden of constitutively expressed protein components.
ii. Dissolve Tet2-Et ncAA in DMF as described in Recipes. We recommend making the stock with ~10% more volume than is needed for expressions. For example, for a 50 mL expression, you need 250 µL of 100 mM Tet2-Et solution to reach 0.5 mM final concentration, and so we recommend making ~275 µL of stock for this expression.
iii. Prepare defined-AIM or ZY-AIM as described in Recipes.
After adding all reagents and antibiotics, and before adding Tet2-Et, split the media into Tet2-Et-containing and Tet2-Et-lacking expression batches.
1) For Tet2-Et-containing expressions, add 250 μL of 100 mM Tet2-Et stock to each 50 mL expression (0.5 mM Tet2-Et final).
2) For expressions lacking Tet2-Et (wild-type expressions and minus ncAA culture expressions), add 250 μL of DMF (100%) for each 50 mL expression.
iv. Inoculate expression cultures with non-inducing overnight cultures so that the starting OD600 upon incubation is 0.05 (e.g., if overnight starter cultures have an OD600 of 5, add 0.5 mL to a 50 mL culture)
v. Add antifoam to each culture to enable proper aeration. Two drops or ~50 µL should be sufficient to eliminate foam in 50 mL cultures.
Note: For 1 L cultures, add 6 drops or ~150 μL.
vi. Grow at 37 °C at 250 rpm for 20–24 h.
Note: If expressing a protein that requires lower expression temperatures, monitor OD600 until it reaches ~1.5, then lower to the desired temperature, and continue culturing for another 16–24 h.
Day 4: Evaluating expressions and harvesting cells
Note 1: After expression, it is convenient to estimate the amount of sfGFP produced by measuring culture fluorescence, since only full-length sfGFP (and not protein that was prematurely truncated at the TAG codon) will fluoresce. Yields determined by fluorescence are not directly comparable between sfGFPWT and sfGFPTet2-Et as the Tet moiety affects fluorescence properties (see Figure 1, and General note 2 for information on tetrazine redox and quenching).
After 20–24 h of expression, measure OD600 and fluorescence of sfGFPWT and both sfGFP150 expressions (Tet2-Et-containing and Tet2-Et-lacking cultures; ex/em: 485/510 nm). The sfGFPWT culture should be visibly green, while the sfGFP150 expression culture should be orange (quenched sfGFP, Figure 4). The OD600 values will vary depending on the target protein. Normal values will range from ~2.5 to 15. Final OD600 values below 2 are indicative of poor cell growth and/or protein expression. Characteristic fluorescence values for all discussed expression conditions are summarized in Figure 5.
Harvest cells by centrifugation.
i. Centrifuge cells at 5000–10,000× g for 10–20 min at 4 °C and then decant or aspirate the media.
ii. Resuspend the cell pellet in the appropriate buffer for the downstream application, and either store at -80 °C (flash-freezing cells in liquid nitrogen may help maintain the integrity of unstable proteins) or proceed with purification.
1) Buffer choice is often contingent on protein purification strategy.
2) Here, the buffer can be supplemented with a cryoprotectant [e.g., 10% (v/v) glycerol] to minimize adverse effects associated with freezing sensitive or unstable proteins.
3) For His6-tagged proteins to be purified via TALON resin, a recommended resuspension/lysis buffer would be 50 mM Tris pH 7.5, 500 mM NaCl, 10% (v/v) glycerol, and 5 mM imidazole. Avoid the use of reducing agents such as DTT or b-mercaptoethanol as they can react with (reduce) and temporarily inactivate the Tet2 amino acid [1].
Evaluation of Tet2-protein reactivity with sTCO-PEG5000 and other sTCO-probes
Purifying Tet2 proteins: important considerations
The sfGFPWT and sfGFPTet2 proteins expressed above contain C-terminal His6 tags and can be purified using TALON of Ni-NTA metal affinity resins according to manufacturers’ recommendations.
Avoid the use of reducing agents during the purification of Tet2 proteins as they can reduce the Tet2 amino acid, rendering it temporarily unreactive.
Quantifying purified Tet2-containing proteins: important considerations
Quantifying Tet2-protein concentration can be performed with normal standard methods including Bradford, BCA, Lowry assay, or measuring protein absorbance at 280 nm (A280) using UV-VIS as described below. See General note 2 for more information on accurate Tet2-protein concentration determination using A280 measurements.
For sfGFPWT and sfGFPTet2-Et proteins, the molar extinction coefficients at 280 nm (ϵ280) of 24,080 and 37,640 M-1·cm-1 can be used, respectively [1].
The concentration can be determined using Beer’s Law: A280 = ϵ280 × l × c where l is the pathlength (usually 1 cm) and c is the concentration of protein.
Example: An A280 of 1.8 with a 1 cm path length corresponds to 76 μM sfGFPWT or 48 μM sfGFPTet2-Et.
Evaluating Tet2-protein reactivity with sTCO-PEG5000
Accurate Tet2 encoding and its reactivity on a protein of interest is most easily assessed by measuring electrophoretic mobility shifts upon conjugation with sTCO-PEG5000. After successful conjugation, the attached PEG polymer will slow the migration of reactive target protein while unreactive protein will migrate identical to that of wild-type (unmodified) protein. This assay requires only a few minutes for the reaction to occur and about 1 h to run a standard SDS-PAGE gel, and only small quantities (<10 μg) of protein are needed. While treatment of Tet2-protein with other sTCO reagents can be used to ensure the Tet2-protein is reactive (e.g., by visualizing in-gel fluorescence after reacting with an sTCO-fluorophore), it is not trivial to evaluate the extent or the stoichiometry of conjugation with these methods. See General note 1 for preparation and handling of sTCO-PEG5000 and General note 2 regarding tetrazine reactivity. As a quick and clear diagnostic test, in the following assay, a 10-fold molar excess of sTCO-PEG5000 is reacted with purified Tet2-protein for ~5 min, and then excess sTCO-PEG5000 is quenched with free Tet2 amino acid to eliminate potential nonspecific reaction with the protein prior to boiling for SDS-PAGE analysis.
Prepare a 1 mM stock of sTCO-PEG5000 stock in water and a 20 mM Tet2-ethyl stock in DMF. Very little Tet2-Et is needed for each quenching reaction; we recommend making 100 µL of Tet2-Et at 100 mM in DMF as described in Recipes and then diluting this to 20 mM. The stock can be frozen at -20 °C and repeatedly thawed for quenching steps.
For each 30 μL reaction, 10 μM sfGFPWT or sfGFPTet2-Et protein is incubated with or without 100 μM sTCO-PEG5000 and then quenched with ~1 mM Tet2-Et amino acid. Table 2 shows an example reaction scheme in which the sfGFPWT and sfGFPTet2-Et protein concentrations were determined to be 50 and 20 μM by A280 measurement.
Table 2. sTCO-PEG5000 reactions
sfGFPWT sfGFPWT + sTCO-PEG5000 sfGFPTet2-Et sfGFPTet2-Et + sTCO-PEG5000
Buffer (50 mM Tris pH 7.5, 100 mM NaCl) 24 μL 21 μL 15 μL 12 μL
Protein
(10 μM final)
6.0 μL 6.0 μL 15 μL 15 μL
1 mM sTCO-PEG5000
(100 μM final)
0 μL 3 μL 0 μL 3 μL
Final volume 30 μL 30 μL 30 μL 30 μL
Incubate 5 min at room temperature
20 mM Tet2 amino acid
(~1 mM final)
2 μL 2 μL 2 μL 2 μL
Incubate 5 min at room temperature
4× Laemmli loading buffer [containing + 10% (v/v) β-mercaptoethanol] 10 μL 10 μL 10 μL 10 μL
Denature by incubation at 95 °C for 5 min
Centrifuge samples for 2 min at >10,000× g
Run 10–15 μL of each sample on a 12% SDS-PAGE gel. This equates to running ~2 μg of each sample.
Stain the SDS-PAGE gel using common Coomassie staining methods and evaluate the reactivity of Tet2-protein with sTCO-PEG5000 (Figure 6). This can be done qualitatively, by estimating the amount of protein that has shifted, or quantitatively, by taking a high-resolution scan of the gel and using densitometry software (e.g., ImageJ) to evaluate the exact percentage of reacted sTCO-PEG.
Figure 6. sTCO-PEG5000 gel mobility shift assay to evaluate the reactivity of purified Tet2-protein expressions.
Proteins expressed in defined-AIM (panel A) and ZY-AIM (panel B) media were evaluated for their reactivity by conjugating with a 10-fold molar excess of sTCO-PEG5000. A clear upward electrophoretic shift due to the added molecular weight of PEG5000 is observed for sfGFPTet2-Et proteins but not sfGFPWT, confirming the specificity of Tet2-Et labeling. Complete (> 95%) reactivity is observed for proteins expressed in either BL21(DE3) or B95(DE3) ΔAΔfabR cell lines. See General notes 1–3 for considerations when Tet/TCO reactivity is not complete.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Eddins et al. [1]. Truncation-Free Genetic Code Expansion with Tetrazine Amino Acids for Quantitative Protein Ligations. Journal name (Figure 4, panel D) (Figure S9).
Two biological replicates were used to produce the dataset in the protocol and the dataset referenced.
General notes and troubleshooting
General notes
Considerations for Tet-TCO labeling reactions:
TCO structure and reactivity: TCO molecules can come in a variety of forms, having different stabilities and rates of reactivity with tetrazines. In this protocol, we describe the use of strainedtrans-cyclooctene (sTCO) functionalized reagents (containing the fused cyclo-propyl ring, Figure 1) for conjugation to encoded tetrazines because of their ultra-fast reactivity with tetrazines. sTCO displays second-order kinetics approximately two orders of magnitude higher than standard TCO [15], which greatly reduces conjugation time and the requirement for excess TCO reagent to ensure complete reactivity. Currently, sTCO reagents are not commercially available; however, methods to synthesize them have been well-established [5,15]. To accelerate access to sTCO reagents, we offer a select assortment of sTCO reagents (as well as tetrazine reagents, see General note 4) at the GCE4All Center (https://gce4all.oregonstate.edu/, [email protected]). TCO reagents that lack the strain-promoting cyclopropyl group of sTCO are commercially available and are effective for conjugating tetrazine-containing proteins. If using TCO-functionalized reagents, incubation times will need to be increased ~10-fold to compensate for slower reaction times; however, additional optimizations will be necessary depending on TCO used [15].
TCO stability: sTCO and TCO may undergo spontaneous trans-to-cis isomerization over time. Users should be mindful of how they store these reagents to maximize stability. We have found that large TCO-PEG polymers are relatively stable at -20 for months and appear to be stable dissolved in water at high concentrations. We recommend minimizing the number of freeze/thaw cycles of TCO stocks by aliquoting samples when they are received. TCO-PEG polymers can lose reactivity over time so when attempting to use stoichiometric amounts of label, it may be worth determining the percent reactivity of a TCO-label stock by titrating the sample with freshly prepared Tet-protein.
Causes of non-reactive Tet2-Et protein. Testing reactivity with the sTCO-PEG mobility shift assay validates successful tetrazine encoding when reactions are successful; however, if a reaction does not occur, tetrazine may still be encoded but is in a reduced (unreactive) state (see General note 2 below), the encoding site is buried and inaccessible to labeling reagents, the TCO reagents have degraded, or natural amino acids were encoded at the TAG/UAG codon instead of Tet2-Et. Natural amino acid encoding at amber codons is typically caused by a deficiency in the GCE machinery such that insufficient amounts of amber-suppressing tRNA amino-acylated with Tet2 are generated, allowing endogenous tRNAs to wobble-base pair their anticodons with the UAG codon so that natural amino acids are encoded instead. The most common natural amino acid encoded at UAG codons via near-cognate suppression is glutamine, and such events are more common in RF-1 deficient expression strains [e.g., B95(DE3)]. However, as shown and discussed in Eddins et al. [1], the described Tet2-Et GCE system utilized here was optimized specifically for its ability to routinely and effectively out-compete any near-cognate suppression events in B95(DE3) or BL21(DE3) cells. Mass spectrometry (see General note 3 below) can help delineate whether any non-reactive protein was caused by near-cognate suppression, e.g., if the MS spectra of sfGFPTet2-Et protein contain a peak with a mass consistent with glutamine at site N150 instead of Tet2-Et; then, the GCE machinery system was not functioning adequately and improvements in expression parameters must be considered or more carefully followed as described.
Tetrazine reduction—considerations for reactivity and protein quantification:
The tetrazine amino acid can undergo reversible reduction when exposed to reducing environments. Reduction of free Tet2-Et amino acid by the cells in the culturing media may be observed during protein expression, as indicated by a media color change from pink to clear. Reduction of Tet2-Et amino acid during protein expression can be minimized using baffled flasks that provide high rates of aeration, high shaking rates (220–250 rpm), and antifoam to maximize air exchange rates. Once encoded into a protein, the reducing environment of the E. coli cytoplasm may cause the Tet2 residue to be reduced, but it will oxidize and be fully reactive to sTCO once purified and buffer exchanged (Figure 1). Some considerations on tetrazine redox properties are provided below.
Promoting the oxidized state of encoded Tet2 for complete bioconjugation: After cell lysis, exposure to ambient oxygen during purification and desalting/buffer exchange will cause the encoded, reduced Tet2-Et to spontaneously oxidize to the reactive form (see Eddins et al. [1]). This oxidation event occurs rapidly after purification and desalting, only requiring a few minutes (up to 1 h at most) for nearly all encoded Tet residues on sfGFPTet2-Et to oxidize. The kinetics of oxidation may change depending on the target protein and site of encoding. If proteins are stable overnight at 4 °C, allowing them to oxidize in the fridge after purification and after buffer exchanging into a buffer of choice lacking imidazole (extended exposure to imidazole at high concentrations can inhibit tetrazine reactivity), with a closed cap, is typically sufficient to achieve complete oxidation and maximal reactivity. If proteins are unstable, perform a reactivity-over-time assessment to determine when a given Tet2-protein will be fully oxidized. If overnight incubation is not sufficient for complete oxidation, an additional desalting/buffer exchange step may help.
Effect of Tet2-Et redox state on sfGFP fluorescence. The N150 site in sfGFP resides in close physical proximity to its chromophore, and so tetrazine encoded at this site will quench sfGFP fluorescence when the tetrazine is in its oxidized state, but not when in its reduced form (see Figure 1 [16]). Consequently, when the encoded Tet2-Et is oxidized, sfGFPTet2-Et is orange in color and is reactive to sTCO, but when it is reduced, the sfGFPTet2-Et will be fluorescent green and will not be reactive to sTCO. After a successful reaction with sTCO, the oxidized sfGFPTet2-Et will change from orange color to fluorescent green (see Video S1). This restoration of sfGFPTet2-Et fluorescence upon exposure to sTCO is a convenient strategy to evaluate successful reaction and labeling.
In-cell fluorescence of sfGFPTet2-Et: Because the redox state of tetrazine influences its ability to quench sfGFP fluorescence, and because it is difficult to quantify the ratio of oxidized to reduced sfGFPTet2-Et when inside the cell, it can be difficult to measure how much sfGFPTet2-Et is produced after expressions using in-cell fluorescence. Still, the fluorescence values plotted in Figure 5 are representative sfGFPTet2-Et values for a 20-h expression at this scale. As an approximate rule of thumb, when tetrazine is fully oxidized, sfGFPTet2-Et fluoresces at approximately one-sixth the amount of sfGFPWT.
Estimating Tet2 protein concentration using A280: Tet2-protein concentration can be determined using standard methods including UV light absorption, Bradford, BCA, or Lowry assays. It is convenient to estimate Tet-protein concentrations using UV-VIS and Beer’s Law; however, using an accurate molar extinction coefficient is necessary. Encoded Tet2-Et affects the extinction coefficient significantly when the residues tetrazine group is in its oxidized form, but to a much lesser extent when in reduced form. For consistency of protein quantification, we emphasize the importance of providing sufficient incubation time for all Tet2-protein to oxidize. To calculate the approximate extinction coefficient (ϵ280) of a Tet2-containing protein, use the following formula:
ϵ280 = (# of Tet2 residues) × (13,560 M-1·cm-1) + (# of Trp residues) × (5,500 M-1·cm-1) + (# of Tyr residues) × (1,490 M-1·cm-1)
Mass spectrometry to evaluate encoding fidelity:
Mass spectrometry enables direct evaluation of tetrazine ncAA encoding fidelity and can help delineate sources of non-reactive protein. We recommend using whole-protein mass spectrometry to evaluate tetrazine encoding fidelity in new proteins and expression constructs. We also recommend comparing tetrazine-containing protein to the wild-type variant so that the expected differences in mass can be confirmed (see Figure 7) Note that it is possible that no off-target MS peaks are detected even though notable quantities of non-reactive protein are observed in the sTCO-PEG5000 gel mobility assays. In these cases, the lack of reactivity is likely caused by the presence of multiple independent non-reactive protein species that individually are in too little abundance to be detected by MS but collectively are observed as a single aggregate band in the electrophoresis assays. Still, this appears to be a negligible amount of unreactive protein.
Figure 7. ESI-mass spectrometry comparison of sfGFPTet2-Et (green) and sfGFPWT (black). sfGFPWT: expected mass 27,827 Da, observed 27,826.5 Da. sfGFPTet2-Et: expected mass 27,968 Da, observed 27,967.8 Da. Observed mass difference is 141.3 Da, matching the expected 141 Da difference. Commonly observed methionine loss and sodium adduct peaks are highlighted. This data was adapted from Eddins et al. [1].
Site-specificity of Tet2-Et encoding can be confirmed by tryptic digestion of target proteins followed by MS/MS sequencing methods. Since different peptide fragments ionize with different efficiencies, these fragmentation methods should only be used to confirm the site of encoding and should not be used to assess the fidelity of encoding.
Availability of tetrazine and TCO reagents:
Tet2-Et
i. Tet2-Et is available for purchase from the GCE4All Center product #1001 (see https://gce4all.oregonstate.edu/tetrazine-amino-acids).
ii. See Blizzard et al. [17] for synthesis protocol.
TCO reagents
i. sTCO-PEG5000 can be requested via email at [email protected].
ii. See Bednar et al. [5] or Fang et al. [15] for synthesis protocol.
iii. TCO-coupled reagents are available from various commercial vendors.
Troubleshooting
Improving protein expression efficiency
While the sfGFPTet2 protein should express efficiently if following the above protocol, optimization may be necessary to achieve efficient expression of biologically relevant proteins containing the Tet2 amino acid.
Construct design:
Placement of affinity purification tag, N-terminus vs. C-terminus: C-terminal affinity purification tags are preferred when expressing Tet2 proteins in BL21(DE3) cells to avoid the co-purification of truncation products. If N-terminal tags are preferred, or if the target protein is not stable or compatible with non-native C-terminal affinity tags, ion-exchange and size-exclusion chromatography can be used to separate some full-length proteins from their truncation products. If TAG codons are placed in close proximity to the N-terminus, then truncation products may be sufficiently small that they get degraded or are insoluble, and therefore they will not be co-purified with N-terminal affinity tags. Alternatively, proteins can be expressed in B95(DE3) ΔAΔfabR cells and no truncation product will be produced.
Solubilization fusion proteins: fusing target proteins with N-terminal solubilizing fusion proteins such as maltose binding protein (MBP), glutathione-S-transferase (GST), and small ubiquitin modifying protein (SUMO) can often increase expression yields and solubility of challenging target proteins. Adding a proteolytic cleavage sequence (e.g., TEV, 3C) between the fusion protein and the target protein allows users to remove the solubility fusion protein during purification processes, so they do not interfere with downstream assays. SUMO proteins can be cleaved with their cognate SUMO proteases. See Esposito and Chatterjee [18] for additional information on using fusion proteins as solubility tags.
Alternative expression strategies
Manual induction: We generally find auto-induction expression methods as described above the most efficient strategy for encoding Tet2 into proteins. However, manual induction methods in which an inductant (e.g., IPTG when using pET vectors) is added to the media to initiate protein expression are also compatible with the pAJE-E7 expression system [1]. For example, protein can be expressed by growing freshly transformed BL21(DE3) cells in a standard rich media (e.g., LB, 2× YT) at 37 °C until cultures reach mid-log phase (e.g., OD600 ~0.6–0.8) and adding IPTG to a final concentration of 0.5–1 mM and Tet2-Et to 0.5 mM. Expression temperature can be adjusted to be between 18 and 37 °C as needed, and protein can be expressed for 3–24 h. As with standard manual induction and expression of wild-type proteins, it is important to optimize conditions (e.g., inductant concentration, temperature, media). These factors should be optimized using wild-type target protein constructs and TAG-interrupted proteins in parallel, including sfGFP controls.
Best practices for using ImageJ/other densitometry software to evaluate reactivity
Quantification of protein reactivity can be useful for determining the success of tetrazine fidelity, redox, and the stability of TCO probes. It is important to use densitometry software properly, without biasing the extent of reactivity in either direction. Because PEG mobility shifts cause a spreading effect on SDS page gels (see diffuse of higher MW bands in Figure 6), the best practice is to measure the remaining area of the band of unreacted protein instead of the area of the shifted protein.
When possible, it is best to use area boxes of equal sizes for each band. SDS-PAGE gels can sometimes have variable sizes for lanes, which may require multiple sizes of area boxes to be used.
Reactivity quantification is less accurate for gels that have a high background signal or gels with overloaded protein lanes. Background subtraction tools can be used to account for this; however, destaining SDS gels properly will help eliminate these concerns.
Acknowledgments
The original research paper in which this protocol was developed and validated is Eddins et al. [1]. This work was supported by the GCE4All Biomedical Technology Development and Dissemination Center supported by National Institute of General Medical Science grant RM1-GM144227. We thank Yogesh M. Gangarde and Subhashis Jana for chemical synthesis and preparation. We also thank Phillip J. Zhu and P. Andrew Karplus at the GCE4All Center for providing insights and discussions during project development and manuscript preparation.
Competing interests
The authors declare no competing interests.
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Supplementary information
The following supporting information can be downloaded here:
Video S1. sfGFPTet2-Et Reaction with sTCO-PEG5000
Article Information
Publication history
Received: Apr 20, 2024
Accepted: Jul 7, 2024
Available online: Jul 25, 2024
Published: Aug 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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5,049 | https://bio-protocol.org/en/bpdetail?id=5049&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Using Localization Microscopy to Quantify Calcium Channels at Presynaptic Boutons
BM Brian D. Mueller §
SM Sean A. Merrill
LD Lexy Von Diezmann
EJ Erik M. Jorgensen
(§ Technical contact)
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5049 Views: 770
Reviewed by: Oneil Girish Bhalala Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Calcium channels at synaptic boutons are critical for synaptic function, but their number and distribution are poorly understood. This gap in knowledge is primarily due to the resolution limits of fluorescence microscopy. In the last decade, the diffraction limit of light was surpassed, and fluorescent molecules can now be localized with nanometer precision. Concurrently, new gene editing strategies allowed direct tagging of the endogenous calcium channel genes—expressed in the correct cells and at physiological levels. Further, the repurposing of self-labeling enzymes to attach fluorescent dyes to proteins improved photon yields enabling efficient localization of single molecules. Here, we describe tagging strategies, localization microscopy, and data analysis for calcium channel localization. In this case, we are imaging calcium channels fused with SNAP or HALO tags in live anesthetized C. elegans nematodes, but the analysis is relevant for any super-resolution preparations. We describe how to process images into localizations and protein clusters into confined nanodomains. Finally, we discuss strategies for estimating the number of calcium channels present at synaptic boutons.
Key features
• Super-resolution imaging of live anesthetized C. elegans.
• Three-color super-resolution reconstruction of synapses.
• Nanodomains and the distribution of proteins.
• Quantification of the number of proteins at synapses from single-molecule localization data.
Keywords: C. elegans Super-resolution microscopy Quantitative localization microscopy Calcium channels Pre-synapse Neuroscience
Graphical overview
Emissions from blinking fluorophores are collected, and 3D point spread functions are fit to calculate emitter positions. Emitters are clustered by position and resolution. These clusters represent individual proteins or clumps of proteins. Based on single protein statistics, the number of proteins at any position is approximated by the number of blinks at a site or the total photon flux per site.
Background
Proteins’ scale is in the order of nanometers; for example, GFP is approximately 4 nm across. However, due to the diffraction limit of light, collecting and focusing the light from GFP onto a camera sensor will, at best, produce an Airy disk with a diameter of 440 nm. To understand the distribution of proteins in cells, microscopy methods that can bypass the diffraction limit of light are required. Resolution is the spatial limit at which the certainty of resolving two nearby proteins fails; thus, high spatial resolution becomes particularly important for small subcellular structures like synapses [1]. Synapses contain highly specialized domains, such as the active zone, but are just a few hundred nanometers in diameter. These highly organized domains allow rapid and reliable neurotransmission [2–4]. To understand how these domains work, the underlying proteins and their nanoscale organization relative to each other must be determined.
Super-resolution methods that achieve resolutions below 100 nm can be divided into stimulated emission microscopy (for example, STED) or localization microscopy (for example, PALM and STORM). Stimulated emission is a scanning method that, in essence, reduces the diameter of the laser beam. Localization microscopy illuminates the whole sample and collects a 2D image of fluorescence on a camera face but limits the number of molecules that contribute to any single frame of the movie so that they appear as single blinks of light. Next, a modeled 3D-point spread function is fit to the blinks of light to reconstruct an image in x, y, and z [5–8].
The coordinate data produced from fitting a 3D-point spread function to the signal from each emitter has a precision determined by the Cramér-Rao lower bound (reviewed in von Diezmann et al. [9]). In contrast, the entire set of coordinate data achieves a resolution that can be determined by several methods. Fourier ring analysis is one common method to determine the resolution that reflects both localization precision and how well a set of localizations samples the underlying structure [10]. By this metric, we typically obtain datasets with 40 nm resolution from live worms. Given that ion channel complexes are ~10 nm in diameter, these data fall short of resolving single channels.
Beyond single proteins, we aim to determine the organization of two different proteins relative to one another. The spatial distributions of distinct proteins can be related in four ways: the proteins can be organized into a single coincident domain, overlapping domains, or adjacent domains, or be completely disjointed. Critically, the borders of protein domains, like the borders of a country, describe the essential information. While more difficult to analyze than an image, coordinate data can more completely describe these spatial relationships.
Here, we demonstrate multicolor super-resolution imaging in live anesthetized C. elegans of directly tagged and labeled endogenous proteins. Further, we discuss quantifying proteins and analyzing images from 3D single-molecule localization microscopy data. This protocol describes how to collect and analyze data from individual synapses, to count the number of ion channels, and determine the spatial relationships between nanodomains of different proteins.
For counting single proteins, the use of endogenous labels vs. antibodies has two advantages. The first is saturation. The TMR dye saturates endogenous proteins tagged with HALOtag in one round of staining (Figure 1). Therefore, the number of proteins can be quantified without caveats from epitope availability or binding affinities of antibodies [11]. The second is precision. Antibodies introduce an additional ~20 nm of uncertainty into localization due to their size. By contrast, the size of self-labeling enzyme tags is approximately 4 nm. One drawback of live cell imaging vs. antibodies on fixed samples is that oxygen-scavenging buffers cannot be used, leading to increased photobleaching (and thus lower photon counts or fewer localizations). Given these considerations, our protocol produces a reconstruction with sufficient localizations and resolution considering the typical size and separation of neuronal protein microdomains [12–14].
Figure 1. HALO-TMRstar saturates UNC-10::HALO after 60 min. L4 animals were incubated with 5 μM HALO-TMRstar (Green) for 60 min. After initial staining, animals were allowed to recover on a nematode growth media (NGM) plate with OP50 bacteria for 1 h. The animals were stained again in 5 μM HALO-SIR (Red) for 60 min and then allowed to recover for 4 h. Synaptic regions near the pharynx were imaged by localization microscopy. Scale bar = 1 μm.
To determine if two protein microdomains are associated, Pearson’s or Mandel’s correlation coefficients are commonly used to determine colocalization in fluorescence microscopy [15]. These tests rely on diffraction-limited signals to create overlapping signals. However, for super-resolution methods, as spatial resolution approaches the size of a protein, the correlation coefficients in these tests will approach zero. For these reasons, it is preferable to analyze the data using nearest neighbor values. Importantly, nearest neighbor analysis distinguishes domains that are coincident, domains that do not overlap but are adjacent, domains at fixed distances, and domains with no relationship. In contrast to nearest neighbor measurements, the spatial relationship of localizations to a center axis or biological landmark has been successful at describing the underlying biology [1,16]. In the following protocol, we describe how to count the number of proteins in a nanodomain, how to measure the size of the domain, and how to determine relationships between protein domains based on localization coordinates.
For this study, we used a Bruker Vutara SRX352. In brief, this is a super-resolution microscope that uses wide-field illumination of the sample. Wide-field illumination activates fluorophores and collects light from an entire vertical column; thus, thin samples or samples positioned with regions of interest near the objective are crucial for reducing background emissions. The microscope uses a proprietary optical biplane that splits the focal plane by ~700 nm in Z-space onto the camera face. Biplane imaging permits improved axial resolution during reconstructive microscopy and provides 1.2 μm of optical thickness at a single z-position. Further, this microscope is capable of 3-color super-resolution imaging with 488, 561, and 641 nm excitation lines. These laser lines are paired with emissions filters for green (BP497-538), orange (BP570-629), and red (BP647-739) fluorophore imaging. Although we describe the specific routine for a Vutara SRX352, in principle the steps listed here are applicable to any super-resolution microscope.
Here, Sections 1–4 deal with the details for mounting and imaging C. elegans nematodes, and Sections 4–8 concern imaging and analysis of localizations.
Materials and reagents
Biological materials
C. elegans strain EG9617 (Jorgensen Lab, elks-1::Skylan-S, egl-19::SNAP IV; unc-2::HALO X)
C. elegans strain EG9667 (Jorgensen Lab, elks-1::Skylan-S, egl-19::SNAP IV; unc-68::HALO V)
Nematode growth media (NGM) plates seeded with OP50 (see Wormbook.org)
Reagents
DMSO (Millipore Sigma, catalog number: D2650-5X5ML)
JF549cp-SNAPtag Ligand (Janelia Labs), store freeze-dried at -80 °C. Available by request from: https://janeliamaterials.azurewebsites.net/
JF646-HALOtag Ligand (Janelia Labs), store freeze-dried at -80 °C. Available by request from: https://janeliamaterials.azurewebsites.net/
Sodium azide (NaN3) (Millipore Sigma, catalog number: S2002)
Agarose (Gold Biotechnology, catalog number: A-201-1000)
NaCl (Millipore Sigma, catalog number: S5886-5KG)
K2HPO4 (Millipore Sigma, catalog number: P8281-500G)
Na2HPO4 (Fisher Scientific, catalog number: S373-3)
MgSO4 (Millipore Sigma, catalog number: M2643-500G)
Solutions
M9 buffer (see Recipes)
25 mM sodium azide in M9 (see Recipes)
4% agarose in M9 (see Recipes)
Recipes
M9 buffer
Reagent Final concentration Amount
K2HPO4 22 mM 3 g
Na2HPO4 34 mM 6 g
NaCl 86 mM 5 g
MgSO4 (1 M) 1 mM 1 mL
H2O n/a 1,000 mL
Total n/a 1,000 mL
M9 has a shelf-life of one year. Dispose if fungal or bacterial contamination occurs.
25 mM sodium azide in M9
Reagent Final concentration Amount
Sodium azide (powder) 25 mM 16.26 mg
M9 buffer n/a 10 mL
Total n/a 10 mL
Store at -20 °C in single-use aliquots to avoid repeated freeze-thaw cycles. The paralytic properties of sodium azide solution begin to degrade after one year in storage.
4% agarose in M9
Reagent Final concentration Amount
Agarose 4% 2 g
M9 buffer n/a 50 mL
Total n/a 50 mL
Heat in the microwave until dissolved. Store at 4 °C in aliquots to avoid evaporation from heating cycles. Agarose aliquots have a shelf-life of one year. Dispose if evaporation or fungal or bacterial contamination occurs.
Laboratory supplies
1.5 mL microcentrifuge tubes (Life Science Products, model: Omniseal, catalog number: M-1700C-1M)
Pasteur pipettes (Fisher Scientific, catalog number: 13-678-20B)
Microscope slides (Fisher Scientific, model: Gold Seal, catalog number: 3048)
Zeiss 1.5 high-performance coverslips (Zeiss, catalog number: 474030-9000-000)
Worm pick (see Wormbook.org)
Equipment
Vutara SRX 352 (Bruker)
Benchtop centrifuge (Thermo Fisher, model: mySPIN6, catalog number: 75004061)
Heat block set to 95 °C (e.g., Eppendorf, model: ThermoMixer F1.5, catalog number: 5384000020)
Software and datasets
Vutara SRX (7.00.00rc39, 2020)
MATLAB (R2022a)
Microsoft Excel
Procedure
Synchronize worms by staging embryos
Day 1
At 9:00 am, use a worm pick to move 20 young adult animals onto a new NGM plate seeded with OP50; do this three times to create three plates total. This procedure takes approximately 10 min.
At 5:00 pm, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three additional plates.
Days 2–3
At 9:00 am, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three additional plates.
At 5:00 pm, move the 20 adult animals from each plate onto new NGM plates seeded with OP50 to create three new plates.
Move the original adult animals to fresh plates in the morning and evening for one more day.
Label SNAP and HALO
Day 4, evening
Inspect plates to select for broods with an abundance of L4 hermaphrodite animals. This is a synchronized plate.
Wash animals off the three synchronized plates using ~1 mL of M9 buffer and a glass Pasteur pipette. Place animals into a microcentrifuge tube. Worms will stick to the plastic of a micropipette, so use glass to move animals.
On a benchtop centrifuge, gently spin (10 s each) to remove bacteria, removing the supernatant and washing with 1 mL of M9 buffer.
After three washes, remove the supernatant and resuspend the animals in approximately 195 μL of M9 buffer.
Prepare JF dyes by resuspending at 1 mM in DMSO (e.g., 5 nmol of dye in 5 μL of DMSO). Move dye + DMSO suspension into the next dried dye aliquot to create a combination of dyes in suspension. This will create 5 μL of 1 mM JF646-HALOtag Ligand + JF549cp-SNAPtag Ligand dye.
Add dye + DMSO solution to worms in M9 buffer to create a 25 μM final concentration of dye. Mix gently by vortexing.
Note: DMSO above 5% is toxic to worms.
Put the tube with the lid in a freezer box on an orbital shaker and stain for 120 min at RT. Gently flick and spin down worms every 15 min to prevent worms from forming clumps on the bottom of the tube or becoming stuck on the sides of the tube. It is essential that the dye and worms incubate in a dark place like a lidded freezer box.
After 2 h, spin down the worm solution, aspirate off the supernatant, and wash four times with 1 mL of M9 buffer.
With a Pasteur pipette, transfer worms to at least three OP50-seeded NGM plates. Let the M9 droplet dry completely before inverting the plate. This should be performed in the dark (we use a drawer). After the droplet has dried, cap the plates and invert. Let the worms recover in a lidded freezer box for 10–12 h. It is important to not expose the stained worms to light. Ideally, the L4 to adult molt will occur during recovery to minimize background staining of the cuticle and gut.
Note: L4 hermaphrodite animals can be identified by the absence of a vulva, which appears as a white crescent at the center of the ventral side of the animal under a typical dissection microscope. Starting with synchronized plates is perhaps the most crucial step of the protocol because the dye sticks to the cuticle of the animal. If adults are exposed to dye, they will have non-specifically labeled cuticles forever, which interferes with imaging of the nervous system. L4s are the final larval stage of the animal and must molt their cuticle before adulthood. Thus, if L4s are stained and then imaged as young adults, a molt must have occurred during recovery.
Imaging preparation
Day 5, morning
Prepare a 4% agarose pad by heating an agarose aliquot in a 95 °C heat block and placing a small drop on a microscope slide. Then, flatten the pad with another microscope slide (Figure 2A).
Trim the agarose pad to about 10 mm × 5 mm using a microscope slide. The agarose pad should fit entirely under the coverslip (Figure 2B).
Add 2–4 μL of 25 mM NaN3 to the pad.
Use the worm pick to distribute the NaN3 drop evenly across the agarose pad. If NaN3 flows over the edges of the agarose pad, reduce the amount used.
Transfer 20 young adult worms to the agarose pad. Older animals may have been young adults during staining and will be highly fluorescent because they have not shed their cuticle.
Wait for animals to paralyze and straighten. This should take a few minutes. If the droplet of NaN3 is drying out, place the slide into a humidity chamber. Seal the pad with a coverslip. Then, seal the coverslip with nail polish and image (Figure 2B).
Notes:
i. Do not image animals that have been in NaN3for more than 60 min. The paralytic sodium azide causes neuron blebbing and disruption of cell morphology after approximately 1 h.
ii. Avoid air bubbles in the agarose pad or trim them.
Figure 2. Creating a worm imaging pad and slide. (A) The agarose pad acts as a support for the worm sample so it is not crushed by the microscope slide, also immobilizing the animal. (B) Once sealed with nail polish, the worms should last for hours. However, the sodium azide paralytic causes cell blebbing and disruption of cell morphology after about an hour. (C) Left: Under the brightfield illumination of the Vutara SRX 352, the worms will appear as pictured. Right: The roll can be in part determined by the position of the ventral nerve cord exiting the neuropil if there is a fluorescent marker in the nervous system or position of the vulva. Scale bar = 5 μm.
Superresolution imaging of C. elegans
Image worms by 3-color imaging
Organize laser lines in descending order of wavelength, e.g., 1) 646 nm, 2) 549 nm, 3) 488 nm. This reduces the photobleaching of the sample.
In widefield mode: Locate worms on the pad and coverslip.
In super-resolution mode:
i. Use only extremely low laser powers (< 1%).
ii. Set the exposure time to 300 ms.
iii. Focus both imaging planes as much as possible on the dorsal nerve cord (Figure 2A).
iv. Turn lasers off as soon as possible. We try to have the laser on for under 10 s while focusing on the synapses we plan to image.
After the nerve cord is in focus, reduce the exposure time to 30 ms and proceed to record. Tune laser powers for optimal blinking vs. the lifespan of the fluorophores. An example of a properly tuned laser power is shown in Figure 3B.
Notes:
i. The best samples are worms that are oriented with their dorsal nerve cord tilted toward the objective.
ii. Be very cautious with the laser power while finding your sample. The worm is not in oxygen scavenging buffer; thus, fluorophores will bleach. To focus on the cord, use very low laser power and a 300 ms exposure time.
iii. Optimizing laser power while imaging: A laser power that is too low will not induce blinking of the fluorophores. A laser power that is too high will bleach the sample in a few frames. The user should find these limits and note them for their microscope. The localization counts over time displayed in Figure 3B is an example of an ideally tuned laser power.
Image worms—counting channels
Organize laser lines by starting with the protein you wish to quantify, followed by fiducial markers. Fiducial markers label a known structure, like the synaptic dense projection, and are used to spatially orient the experimental channel.
In widefield mode: Locate worms on the pad and coverslip.
In super-resolution mode:
i. Use only extremely low laser powers (< 1%).
ii. Set the exposure time to 300 ms.
iii. Focus both imaging planes as much as possible on the dorsal nerve cord.
iv. Turn lasers off as soon as possible.
After the cord is in focus, reduce the exposure time to 30 ms and proceed to record. Increase the laser powers for optimal blinking vs. lifespan of the fluorophores. This is typically between 4% and 12% with 200 mW lasers and the 40 μm biplane module. For counting emitters, the dyes should be completely bleached after imaging; this typically occurs within 4,000 frames.
Notes:
i. The best samples are worms that are oriented with their dorsal nerve cord tilted toward the objective.
ii. Optimizing laser power while imaging: A power that is too low will not induce blinking of the fluorophores. A laser power that is too high will bleach the sample in a few frames. The user should find these limits and note them for their microscope. The localization counts over time displayed in Figure 3B is an example of an ideally tuned laser power.
Localizing
Proceed to the Localization tab.
Set the background threshold. This should be optimized to minimize background localizations (false positive) but not exclude real emitters (false negative). Adjust the background threshold to produce cutouts (colored boxes) around real signals but minimal cutouts in areas with no signal. An example of the effect of background thresholds is displayed in Figure 3C. For C. elegans, we find the ideal threshold to be between 3 and 5 and rely on denoising (detailed below) to remove false positives.
Select images that demonstrate blinking by adjusting the region of interest frame (Probe x) slider. This is primarily to exclude frames where fluorophores are not blinking. Blinking can be determined by moving 10 frames forward and 10 backward (600 ms range). If emitters are turning on and off between those frames, then they are suitable to analyze.
Notes:
At the beginning of imaging, all fluorophores will be active, and separate light flashes will not be visible. Blinking may not occur for a few hundred frames into the acquisition. Non-blinking frames should not be analyzed.
Background threshold should be optimized across the acquired frames.
Localization cutouts can be turned on in Expert Options under the Localizations tab.
Figure 3. Biplane, blinking, and background thresholding. (A) The Vutara SRX simultaneously images in two focal planes (“biplane”), which splits the light path into two focal planes separated by about 700 nm. To focus on a structure like the dorsal nerve cord, it should be equally in focus on both planes—which ensures that the emitters are in between the two focal planes. Scale bar = 5 μm. (B) Graph of localization cutouts over time. At the start of imaging, there will be many fluorophores in an excitable state. However, as imaging progresses, fluorophores will begin to enter the dark state. This change can be confirmed by the drop in localization cutouts over time. Stochastic blinking is required for super-resolution and can be confirmed by the spikes and troughs in the number of cutouts over time. (C) Examples of background thresholding on image reconstruction of CaV2 in the dorsal nerve cord (background threshold = 2, 10, or 30). Upper: Magenta boxes represent successful PSF fitting. Red arrow shows an example of a false positive. Black arrow shows an example of a successful localization. White arrow shows an example of a false negative. Lower: Magenta points represent protein localizations. Scale bar = 1 μm.
Visualizing
Proceed to the Visualization tab.
Discard localizations that are not accurate in the Advanced Particle Filters dialogue. For live worms, we set an arbitrary threshold of 50 nm radial precision and display localizations on the visualization plot as a 50 nm particle size (Figure 4A).
Denoise the image using Method: Mean Distance and denoise by Percent. Lower the slider until most of the signal outside the cord is removed but the signal inside the cord appears unperturbed (Figure 4B).
Save this image to the View Manager.
For analysis of proteins at individual synaptic boutons, select a region of interest 700 nm radius from the center of an in-focus dense projection. Save this view in View Manager. Export the data and save it as a particle .csv file. Out-of-focus synapses and synapses with chromatic aberrations in the XY plane are discarded. Some chromatic aberrations will be present in the Z plane. Aim to analyze five synapses per animal from five different animals, with a total of 25 synapses analyzed. We recommend saving particle files ordered by “animal#_synapse#”, e.g., “3_1.csv.”
Figure 4. Data filtering. (A) First, reconstructions are filtered to exclude imprecise localizations. (B) Next, localizations are trimmed based on nearest neighbors using the denoising tool. Finally, a region of interest is drawn around a synapse to be analyzed to exclude localizations that cannot be within the synapse based on the size of the bouton. Numbering and green arrows show the click order for the user. Magenta = CaV2, Yellow = CaV1.
Cluster analysis
Click on Advanced Statistics Dialogue and go to the Cluster Analysis tab.
Set the parameters for calling clusters based on the size of the protein and resolution of the image. See note.
Click Compute.
Export the resulting output as a .csv file.
Note: We assume that multiple localizations within a close distance are the same channel blinking multiple times. That distance is defined by the size of the calcium channel and the resolution. Voltage-gated calcium channels are approximately 10 nm in size, and localizations worse than 50 nm in precision were discarded. Thus, any localizations within 60 nm could be from the same channel. Therefore, the Maximum particle distance is set to 60 nm. The Minimum particle count to form cluster is set to 2. An example of clustering is available in Figure 5A.
Data analysis
Mapping distributions of proteins from particle files (produced in section F):
Open MATLAB and load the following script into a command window.
https://github.com/bdmscience/probeDistances/blob/main/probeDist.m
Change the address field to the folder where the particle files are saved.
Modify the underlined section of the first line of code to load the particle file you wish to analyze:
particles = readtable('____.csv')
Change “mkdir” to the worm and synapse you are analyzing, e.g., mkdir 3_1. Then, cd to that new directory.
Lines 4–9 set the x and y coordinates of each imaging channel. “particles.vis_probe == 0” is channel 1 in SRX, “==1” is channel 2, and “== 2” is channel 3. Keep this in mind when considering which channels to analyze. The script default is to compare channel 3 to channel 1.
Run each section of the code to measure the distances between the localizations of the first channel to the center of the second channel, and the localizations of the second channel to the center of the second channel.
Each section will output two variables called “dist”; these are your measurements.
Note: After each section of code, save the “dist” distance outputs before analyzing a new set of channels. We recommend you plot the data in MATLAB. Other graphing software has a maximum number of points that can be displayed on a plot.
Counting calcium channels
Foreword: Presented here are two methods to quantify the number of calcium channels at synaptic boutons. Both methods rely on clustering localizations into possible single protein domains. A caveat for both of these methodologies is that tight clusters of proteins can skew the estimate for blinks or for photon yield for a single protein. These methods of counting are more accurate for dispersed proteins than densely localized proteins but likely produce an undercount of the total number of proteins.
The methods differ in the quantification of signal within those protein clusters. The first calculates the number of localizations per protein. The second calculates the number of photons per protein. The advantages of one over the other are unclear; we recommend users attempt both methods and compare results. For a full discussion of counting methods refer to the methods section titled “Counting calcium channels” in Mueller et al. [14].
Counting proteins by position and particle count (Figure 5B and 5C):
Open the cluster file in Excel from section G in Microsoft Excel.
The ID column refers to cluster ID. Each row is a different cluster of localizations. Cluster 0 are single blinks. “particleCount” contains the number of localizations from channel 1 in each cluster. Using this information, we can calculate how many blinks each protein emitted.
Create a new Excel file called spreadsheet2. Name columns as Number of blinks, Count, Particles in Clusters, Total particles, and Single particles.
Create rows numbered 1 through the max number of “particleCount” in a cluster ID.
Manually count how many clusters contain each number of particles and input the total count in the “count” column. Do this for each synapse and replicate.
Combine these data across synapses or replicates and fit a Poisson distribution to the cumulative data. Lambda equals the mean number of blinks emitted from a protein in the dataset. Record this value on spreadsheet2.
Take the sum of the total number of localizations at each synapse and then divide the total number of blinks by the lambda value to yield how many proteins are present at each synapse. Plot the mean and SEM of these data. We recommend analyzing at least six synapses from three different animals.
Counting proteins by photon flux (Figure 5D and 5E):
Open the particle file from section G in Microsoft Excel.
Sort particles by clusterID. Note that clusterID = 0 are non-clustered localizations.
Sum the photons (“photon-count”) for each non-clustered localization and each clusterID.
Sum the total photons for all ClusterIDs.
Repeat 2a–2d for each synapse.
Take the average “photon-count” per cluster across the dataset.
Divide the “photon-count” from each synapse by the average “photon-count” per cluster (total photons divided by the average number of photons per channel). The result is the number of channels present at that synapse by photon flux. Plot the mean and SEM of these data. We recommend analyzing at least six synapses from three different animals.
Note: We found that counting by photon flux had higher variability than counting by particle count; however, there was agreement between the results. An example of blink number and photon yield for clusters is present in Figure 5F.
Figure 5. Counting calcium channels. (A) CaV2 (magenta) and CaV1 (yellow) localizations with cluster shells. (B–C) Quantification output from counting by particle count. Poisson curve fitting revealed that each protein blinked 2.7 times. (D–E) Quantification output from counting by photon flux. Each protein emitted on average 1996 photons. (F) Example statistics from voltage-gated calcium channel clusters at C. elegans synapses. Scale bar = 250 nm. Figure adapted from Mueller et al. [14].
Data availability and example data
All data mentioned in this study is available as source data files with the original publication. For ease of access, we have included particle files, cluster files, and channel count files as supplementary files with this protocol.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Mueller et al. [14]. CaV1 and CaV2 calcium channels mediate the release of distinct pools of synaptic vesicles. eLife (Figure 4, panels B, C; Figure 5, panels A–C; Figure 5 supplement 1, panels A–E; Figure 6, panels A–D; Figure 7, panels A–D; Figure 8, panels A–D; Figure 10, panels A–G).
Acknowledgments
We thank Luke Lavis and the Lavis lab for providing all JF dyes used in this study. We thank Brittany Graham for feedback on the manuscript, and Josh Kaplan who suggested we attempt to count calcium channels from these datasets. An application of this protocol was originally described in Mueller et al. [14]. Funding sources for this work were the NSF NeuroNex 2014862 to E.M.J., and NIH R01 NS034307 to E.M.J., F31 NS084826 to S.A.M. E.M.J is an investigator at Howard Hughes Medical Institute.
Competing interests
E.M.J holds a patent on optical biplanes, which are integral to the design of the microscope discussed in this study. However, E.M.J does not have any financial stake in Bruker.
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Supplementary information
The following supporting information can be downloaded here:
Cluster files
Particle files
Counting channels by photon flux.xlsx
Counting channels by position and particle count.xlsx
Article Information
Publication history
Received: May 16, 2024
Accepted: Jul 7, 2024
Available online: Jul 25, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Cellular mechanisms > Synaptic physiology
Cell Biology > Cell imaging > Super resolution imaging
Molecular Biology > Protein > Ion channel signaling
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5,050 | https://bio-protocol.org/en/bpdetail?id=5050&type=0 | # Bio-Protocol Content
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Extraction of Bacterial Membrane Vesicle and Phage Complex by Density Gradient Ultracentrifugation
SL Shangru Li *
AR Anmin Ren *
ML Menglu Li
GL Guobao Li
LY Liang Yang
TJ Tianyuan Jia
(*contributed equally to this work)
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5050 Views: 479
Reviewed by: Alba BlesaNuttavut KosemManousos E. Kambouris
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Abstract
The bacterial membrane vesicles (MVs) are non-replicative, nanoscale structures that carry specific cargos and play multiple roles in microbe–host interactions. An appropriate MV isolation method that mimics complex pathogen infections in vivo is needed. After bacterial MVs extraction, flagella or pili can be frequently observed along with MVs by transmission electron microscope (TEM). Recently, MVs from Pseudomonas aeruginosa were found to coexist with Pf4 phages, and this MV–phages complex exhibited a different impact on host cell innate immunity compared with MVs or phages solely. The presence of this MVs–phages complex simulates the real condition of complex pathogen infections within the host. This protocol outlines the extraction of the MVs and Pf4 phages complex of P. aeruginosa PAO1, including the respective isolation and qualification approaches. Our step-by-step bacterial MVs–phages complex extraction protocol provides valuable insights for further studying microbe–host cell interactions and the development of novel phage therapies.
Key features
• Detailed density gradient extraction procedures of MVs–phages complex
• TEM, plaque assay, and PCR to verify the coexistence of MVs and phages
• The obtained MVs–phages complex can be used for exploring phage–microbe–host cell interactions
Keywords: Pseudomonas aeruginosa Membrane vesicles Pf4 phages TEM Plaque assay
Graphical overview
Background
Pseudomonas aeruginosa is an opportunistic pathogen that often causes serious hospital-acquired infections and can lead to a variety of infectious diseases including pneumonia, wound infections, and septicemia [1]. Due to the high mortality rate of these infections, P. aeruginosa was recognized as one of the most life-threatening bacteria and listed as a priority pathogen for research and development of new antibiotics by the World Health Organization [2].
The Pf phage, a filamentous inovirus specifically associated with P. aeruginosa, is a type of temperate phage, which can be classified from Pf1 to Pf8. Approximately half of P. aeruginosa isolates harbor Pf phage operons within their genomes [3,4]. Pf phages are about 6–7 nm in diameter and vary in length from 0.8 to 2 µm, with a single-stranded DNA (ssDNA) as the genetic material packaged within a helical filamentous structure major coat protein CoaB [3]. Of these, Pf4 phage is an important member, and its genome is integrated into the P. aeruginosa PAO1 as the prophage. Thus, Pf4 phage can emerge inside the bacteria [5]. The core genes of the Pf4 phages mainly include xisF4, PA0720, PA0723, PA0724, and PA0726, which encode excisionase, single-stranded DNA binding protein, major coat protein CoaB, minor coat protein CoaA, and morphogenesis protein, respectively [3,5]. A recent study showed that Pf4 phages can be transmitted between bacteria and have various effects on P. aeruginosa, including reducing twitching motility, altering biofilm production, and affecting the release of virulence factors, which are important in the pathogenesis of P. aeruginosa [6–8]. However, how Pf4 is transmitted has remained elusive.
Bacterial membrane vesicles (MVs) are structures produced by bacteria with a diameter ranging from 40 to 400 nm that carry specific cargo and travel a long distance (centimeter scale) to play essential roles in cell–cell communications [9,10]. Gram-negative bacteria MVs could be divided into two types: B (blebbing)-type vesicles, which are formed via blebbing from the outer membrane, and E (explosive)-type vesicles, which are formed due to explosive cell lysis [11,12]. Studies have suggested that P. aeruginosa E-MVs can be formed under certain stresses and bacteriophage lysis conditions induced by Pf4 phages [3].
Many previous works observed that after the crude MV extraction step, flagella can also be observed by TEM [13]. A recent study investigated the combined effects of P. aeruginosa MVs with phages upon the recognition function of the host’s innate immune system. In a mouse model of acute lung infection, treatment with MVs-Pf4 phages complex resulted in reduced infiltration of neutrophils in the lung alveoli, thus suppressing the occurrence of inflammatory reactions [3].
In this protocol, we establish a bacterial MVs–phages co-extraction platform from P. aeruginosa PAO1 as a model organism. The MVs–phages extract effectively mimics the complex MVs–phage coexistence condition in vivo within the host during pathogen infections. Overall, this is an optimized method for extracting the MV–phage complex based on density gradient centrifugation (Figure 1), with feasibility verified through TEM, PCR, and plaque assays. It will provide a reliable protocol for studying bacteriophage–microbe–host cell interactions.
Figure 1. Flowchart of MV–phage complex extraction. The bacterial culture is centrifuged to retain the supernatant. After filtering and ultrafiltering, bacteria and small molecules can be removed. After density gradient ultracentrifuging, flagella and pili can be removed. Then, purified MVs–phage samples can be obtained.
Materials and reagents
The following materials and equipment are recommended for this protocol, but alternatives from other sources can also be used when proven to be equivalent.
Biological materials
P. aeruginosa PAO1 strain (strains are preserved at -80 °C in our laboratory)
P. aeruginosa PAO1 ΔPf4 strain (strains are preserved at -80 °C in our laboratory)
Reagents
Opti-Prep gradient medium (Sigma-Aldrich, catalog number: D1556)
Phosphate-buffered saline (PBS) (Servicebio, catalog number: G4202)
2% Phosphotungstic acid solution (Acmec, catalog number: AG1599)
LB liquid medium (Servicebio, catalog number: G3103)
Premix TaqTM Version 2.0 (Takara, catalog number: R004Q)
1% agarose gel (Baygene, catalog number: 192255)
DNA marker (Coolaber, catalog number: D2000)
Pancreatic digest of casein (peptone) (Beyotime, catalog number: ST800)
Soybean digest (Sigma-Aldrich, catalog number: C7210)
NaCl (Dogesce, catalog number: Sodium Chloride)
Agar (Dogesce, catalog number: Agar)
CaCl2 (Macklin, catalog number: C915443-500g)
MgCl2 (Macklin, catalog number: M813766-500g)
Mitomycin C (AbMole, catalog number: M5791)
GelRed 10,000× gel staining solution (I-PRESCI, catalog number: P202-01)
Solutions
TSB culture medium (see Recipes)
1.5% LB agar solid medium (see Recipes)
0.75% LB agar semi-solid medium (see Recipes)
Recipes
TSB culture medium
The composition of the medium includes distilled water, 1.5% pancreatic digest of casein (peptone), 0.5% soybean digest, and 0.5% NaCl.
1.5% LB agar solid medium
100 mL LB liquid medium with 1.5 g of agar.
0.75% LB agar semi-solid medium
100 mL LB liquid medium with 0.75 g agar, 1 mM CaCl2, and 1 mM MgCl2.
Laboratory supplies
LB agar plate (Servicebio, catalog number: G3104-0910)
50 mL cell culture tube (Bioland, catalog number: ATS05-12-50)
13.2 mL ultracentrifuge tube (Beckman, catalog number: 344059)
1.5 mL centrifuge tube (Eppendorf, catalog number: 0030108051)
PCR tube (Eppendorf, catalog number: 0030124359)
500 mL filter flask with 0.45 μm filter (Biosharp, catalog number: BS-500-XT)
Glow-discharged 200-mesh carbon grid (Beijing Zhongjingkeyi Technology, catalog number: BZ11022B)
250 mL centrifuge bottle (Beckman, catalog number: 356011)
Equipment
Intelligent high-efficiency centrifuge (Avanti, model: JXN-26)
Masterflex easy-load machine (Sartorius AG, model: VFA013)
100 KD Vivaflow membranes (Sartorius AG, model: VF20H4)
Ultracentrifuge machine (Beckman, model: Optima XE-100)
High-pressure sterilization pot (Yamato, model: SQ810C)
Constant temperature shaker (Shanghaiminquanyiqi, model: MQT-60R)
Tecan Spark 10 M plate reader (Tecan Group Ltd., model: Tecan Spark 10 M)
Transmission electron microscope (Hitachi, model: HT7700)
Nano Sight nanoparticle tracking analysis machine (Malvern Instruments Ltd, model: NS300)
Procedure
Preparation of the culture medium
Prepare 3 L of TSB culture medium (see Recipes).
Note: Adjust volume depending on bacteria. We use 3 L for one sample to obtain abundant MVs. The expected protein quantification concentration of the post-enriched MVs solution is 200 ng/μL.
Sterilize the medium in a high-pressure autoclave (121 °C, 21 min).
Bacterial culture
Select P. aeruginosa PAO1 from the -80 °C refrigerator, streak on a 1.5% solid LB plate, and culture upside down overnight at 37 °C.
Using a 50 mL cell culture tube, inoculate a single bacterial colony into the sterilized 20 mL of TSB culture medium and incubate it on a shaking bed at 37 °C until the OD600nm reaches 0.6.
Sub-culture bacteria into 3 L of TSB culture medium and culture on a shaking bed at 37 °C overnight until it reaches the exponential phase (OD600nm > 1.0).
Extraction of MV and filamentous structures complex
Transfer all the above-mentioned cultures to 250 mL centrifuge tubes and centrifuge at 10,000× g for 15 min at 4 °C.
Filter the supernatant through the 500 mL filter flask with the 0.45 μm filter to remove any remaining bacterial cells.
Transfer the above solution to a 3 L conical flask. Connect 100 KD Vivaflow membranes to the outlet end of the Masterflex Easy-Load system, start the system, and concentrate the solution to 72 mL.
Note: This is a rate-limiting step: 2 L of solution normally needs 1 h to be concentrated.
Transfer the concentrate into ultracentrifuge tubes with 12 mL in each tube. Ultracentrifuge concentrated supernatant at 50,000× g for 2 h at 4 °C to precipitate crude MVs. Remove the supernatant.
Add the Opti-Prep gradient medium into the ultracentrifuge tube and then add the crude extracted MV to the top of the solution. Perform density gradient centrifugation purification of the crude MVs using Opti-Prep gradient medium with a gradient range of 15%–60% at 4 °C and centrifuge at 100,000× g for over 16 h overnight. The purified MVs will be located in the 35%–45% density layer (Figure 2).
Figure 2. Stratification of the MVs-phages complex in the Opti-prep medium. After centrifugation at 4 °C for 16 h at 100,000× g, the MVs–phages complex is distributed in the 35%–45% layer of the Opti-prep gradient. Purified MVs can be taken using a 1 mL syringe.
Take the purified MVs using a 1 mL syringe into a 1.5 mL centrifuge tube and wash them continuously by ultracentrifugation with 4 °C 1× PBS at 50,000× g for 2 h at 4 °C.
Notes: The 1.5 mL centrifuge tubes must be soaked in 75% alcohol overnight.
Resuspend the purified MVs in 1 mL of PBS and store at 4 °C.
Validation of the MV–phage coexistence
Nanoparticle tracking analysis (NTA) to assess the quality of MVs
The quantification and size characterization of the MVs were analyzed using the Nano Sight NS300 for NTA.
Note: Nano Sight NS300 is an instrument used for size analysis and counting of nanoparticles and biological particles. It uses the laser to track the movement of particles in a fluid and analyzes these movement trajectories to calculate the size, concentration, and distribution of the particles.
MV samples were diluted (500×) in PBS to obtain a concentration within the recommended measurement range (1–10 × 108 particles/mL). The size and concentration measurements were conducted using the 448 nm laser in scatter mode.
Using a 1 mL syringe, the sample was injected into the instrument and videos were captured in triplicate for 30 s. The mean values for size and concentration were analyzed using NanoSight (NTA software, version 3.0).
Coexistence of MV and phage visualized by negative staining TEM
Incubate 5 μL of the purified MVs on a glow-discharged 200-mesh carbon grid for 1 min to allow binding.
Contrast the grids with a 2% phosphotungstic acid solution.
Fit the microscope with a high-sensitivity real-time charge-coupled device camera for image capture.
View the grids on an HT7700 transmission electron microscope operating at 100 kV.
Plaque assay to verify the coexistence of phages in bacterial MVs
Pour 25 mL of sterilized 1.5% LB medium with agar into a Petri dish and let it solidify as the underlayer. Take another sterilized 25 mL of LB semi-solid medium (containing 0.75% agar), add 600 μL of freshly cultured ΔPf4 bacteria (OD600nm = 0.6) into this medium, and mix thoroughly. Then pour onto the solidified underlayer as the upper layer.
Prepare the bacteriophage: add 0.5 μg/mL mitomycin C to the culture medium and culture the bacteria overnight. Take 1 mL of the culture fluid for centrifugation and filter the supernatant twice through a 0.22 μm filter.
The bacteriophages to be tested should be diluted in sterile PBS in a 10-fold serial dilution. After each dilution, take 5 μL of the bacteriophage solution, spot it onto the surface of a bacterial semi-solid agar plate, and allow it to dry.
Incubate the plate inverted at 37 °C overnight. The following day, count and analyze the plaques formed.
PCR validation of the coexistence of Pf4 phages in bacterial MVs
Design primers of Pf4 phages for the target genes PA0720, PA0724, and PA0726 based on the gene sequence of the Pf4 bacteriophage using SnapGene software (Table 1). Synthesize the primers and dissolve them.
Note: When validating the binding of other types of bacteriophages in experiments, it is necessary to design primers for specific genomic loci of the bacteriophage according to the actual situation.
Table 1. Primer sequences for PA0720, PA0724, and PA0726
PA0720 PA0724 PA0726
Forward primer ACGAACACCCGTGGTTGGCT AAGGCGTAACGGGTGCTCTG CTCGATCAGATCATCGCCTT
Reverse primer TAGGCCTGTTGCCATGCGAG GCGGCATACATGCTGCGGAT TGGTGTCGTCGAAGAACAAC
Using 1 μL of purified MVs as templates, perform PCR by adding primers. The PCR reaction system is as follows (Table 2 and Table 3).
Table 2. PCR reaction master mix
Reagent Amount
Premix TaqTM Mix 25 μL
Template 1 μL
Forward primer 2 μL
Reverse primer 2 μL
ddH2O 20 μL
Table 3. PCR cycling conditions
Steps Temperature Time Cycles
Initial denaturation 98 °C 10 min 1
Denaturation 98 °C 10 s
35 cycles
Annealing 55 °C 30 s
Extension 72 °C 1 Kb/min
Final extension 72 °C 5 min 1
Hold 4 °C Hold
After the reaction, prepare 50 mL of 1% agarose gel, heating it in a microwave until completely dissolved. Let it cool down to around 50 °C, then add 5 μL of GelRed 10,000× gel staining solution, mix well, and pour the mixture into a gel tray.
Load the samples and DNA marker onto the gel and perform electrophoresis at a constant voltage of 164 V for 20 min using an electrophoresis apparatus.
Visualize the gel using a gel imaging system for observation.
Data analysis
In this protocol, MVs require stringent validation and quality control measures. To this end, an initial step focuses on the measurement and analysis of the size of the extracted MVs, which is an essential aspect of assessing the quality of the MV extraction process. Techniques such as TEM, plaque assay, and PCR are performed to verify the coexistence of MVs and Pf4 phage. Herein, we analyze the data obtained to provide experimental references according to Liu et al. [14].
NTA test analysis of MVs size and concentration
Using the NTA method, the size and concentration of the extracted MVs can be measured to assess the quality of the extracted P. aeruginosa MVs. Results show that the (mean) size and concentration of MVs in PAO1 WT are 67.19 nm and 2.03 × 1010 particles/mL, respectively (Figure 3).
Figure 3. Bacterial membrane vesicles (MVs) concentration and size measured by the nanoparticle tracking analysis (NTA) method. Based on the NTA measurement and software analysis results, we can conclude that the concentration of the measured sample is 2.03 × 1010 particles/mL, with sizes distributed mainly between 30 and 80 nm and with a mean size of 67.19 nm.
TEM observation of the coexistence of MVs and filamentous structures
We used negative staining TEM to observe the structures of the extracted complex. MVs, flagella, and phages can be observed directly through this method (Figure 4). TEM results clearly show that MVs are nearly 100–200 nm in size and comprise spherical structures harboring membrane bilayers. However, it is hard to distinguish whether the filamentous structures are phage or flagella from the outside structure. Thus, we performed further verifications to confirm whether phages were included.
Figure 4. TEM observation to show the coexistence of membrane vesicles (MVs) and filamentous structures. A. In the image, white spherical structures with a size of approximately 100–200 nm can be observed, as indicated by the red arrow, which is the extracted MVs. B: Multiple filamentous structures can be observed in the image, as indicated by the red arrow, which is a mixture of flagella and bacteriophages.
Plaque assay to verify the coexistence of phages and MVs
We utilized the supernatant of the bacterial strain culture fluid as a positive control and the supernatant of the ΔPf4 strain culture fluid as a negative control, demonstrating the presence of bacteriophages in the extracted MVs. The concentration of bacteriophages in the original sample is expressed in plaque-forming units (PFU), and the formula is PFU = (The number of plaques * Dilution factor)/ the volume of the inoculum.
Quantification results show that the number of plaque-forming units (PFU) dropped from 5.2 × 104 in the P. aeruginosa phages to 600 in PAO1 MVs, thereby showing the coexistence of phages and MVs (Figure 5).
Figure 5. Plaque assay to verify the coexistence of phages in bacterial MVs. A. Plaque assay of PAO WT, ΔPf4 phages, and PAO1 MVs on the ΔPf4 bacteria lawn. B. Quantification of PFUs in PAO1 WT bacteria and PAO1 MVs. Data are presented as means ± SD (n =3, ***P ≤ 0.001, Mann-Whitney U test).
PCR verification of the coexistence of Pf4 phages and MVs
In this experiment, we selected three characteristic gene fragments, PA0720, PA0724, and PA0726, as important indicators for detecting the presence of the Pf4 bacteriophage. After conducting PCR on the purified P. aeruginosa MVs, we detected the characteristic gene fragments of the Pf4 phage, PA0720, PA0724, and PA0726, which further showed that P. aeruginosa MVs seemed to coexist with Pf4 phage (Figure 6).
Figure 6. PCR verification of coexistence of Pf4 phages in bacterial membrane vesicles (MVs). Lane 1: DNA marker. The size of PA0720 is 816 bp, and enrichment is visible in lanes 3 and 4; the size of PA0724 is 399 bp, and enrichment is visible in the 250–500 bp region of lanes 5–7; the size of PA0726 is 1937 bp, and significant enrichment is visible around 2,000 bp in lanes 8–10. This confirms the presence of the Pf4 phage in the extracts.
Validation of protocol
The whole procedure has been validated in Pennetzdorfer et al [3].
General notes and troubleshooting
In this experiment, we chose P. aeruginosa PAO1 as the experimental subject. When using the protocol to extract MV–phage complexes from other bacteria, it is important to select a more suitable culture medium.
In the bacterial culture step, it is necessary to cultivate the bacteria to the exponential growth phase and monitor that the OD600 value exceeds 1.0 to obtain a sufficient number of bacteria and MVs. If the OD600 value is too low, it may reduce the yield of MVs. If the MV extraction concentration is low, it may be helpful to attempt further bacterial expansion.
Density gradient centrifugation is the most critical step in this experiment, requiring careful attention to centrifugation speed, time, and temperature. Insufficient centrifugation time can lead to incomplete layer separation and result in mixed impurities in the extract. When using a syringe to extract MVs–phage complexes from the 35%–45% layer, caution is necessary to avoid contamination from other layers. If mixing between layers occurs during this step, it is necessary to repeat this procedure.
For experimental rigor, we employed NTA, TEM, plaque assay, and PCR to confirm the coexistence of extracted MVs with Pf4. During practical operations, depending on needs and lab equipment, you can select necessary inspection methods to quality-check the extracted complex.
After extraction, the protein concentration of the extracted MVs can be determined by the BCA method for preliminary quality inspection. Following the complete protocol procedure, the expected protein concentration is approximately 200 ng/μL. If the measured protein concentration is too low, it indicates that the quantity of extracted MVs is insufficient, and it may be necessary to consider repeating the bacterial culture steps to achieve a higher initial bacterial concentration.
Acknowledgments
This work was supported by the National Key Research and Development Program of China (2022YFC2304700); the National Natural Science Foundation of China (32300068, 32270196, 91951204, 32200155, 32200053, and 32300060); Guangdong Basic and Applied Basic Research Foundation (2019A1515110640 and 2020A1515010316); Shenzhen Science and Technology Program (KQTD20200909113758004, 2022303002); and Guangdong Pearl River Talent Plan.
Competing interests
The authors declare no competing interests.
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Received: May 9, 2024
Accepted: Jul 12, 2024
Available online: Aug 2, 2024
Published: Aug 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Calibrating Fluorescence Microscopy With 3D-Speckler (3D Fluorescence Speckle Analyzer)
CL Chieh-Chang Lin
AS Aussie Suzuki
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5051 Views: 405
Reviewed by: Keisuke TabataIvonne Sehring Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Jan 2023
Abstract
Fluorescence microscopy has been widely accessible and indispensable in cell biology research. This technique enables researchers to label targets, ranging from individual entities to multiple groups, with fluorescent markers. It offers precise determinations of localization, size, and shape, along with accurate quantifications of fluorescence signal intensities. Furthermore, an ideal fluorescence microscope can achieve approximately 250 nm in lateral and 600 nm in axial resolution. Despite its integral role in these measurements, the calibration of fluorescence microscopes is often overlooked. This protocol introduces the use of 3D-Speckler (3D fluorescence speckle analyzer), a semi-automated software tool we have recently developed, for calibrating fluorescence microscopy. Calibration of fluorescence microscopy includes determining resolution limits, validating accuracy in size measurements, evaluating illumination flatness, and determining chromatic aberrations. 3D-Speckler is user-friendly and enables precise quantification of fluorescence puncta, including nanoscale 2D/3D particle size, precise locations, and intensity information. By utilizing multispectral fluorescence beads of known sizes alongside 3D-Speckler, the software can effectively calibrate imaging systems. We emphasize the importance of routine calibration for imaging systems to maintain their integrity and reproducibility, ensuring accurate quantification. This protocol provides a detailed step-by-step guide on using 3D-Speckler to calibrate imaging systems.
Key features
• Semi-automated particle detection
• Accurate three-dimensional measurement of fluorescent particle sizes
• High-precision three-dimensional localization of fluorescent particles
• Precision analysis of point spread function and chromatic aberration in fluorescence microscopy
Keywords: Fluorescence microscopy PSF Resolution Calibration Confocal microscopy Super-resolution microscopy
Background
Fluorescence microscopy serves as an essential tool in a variety of research fields, offering the flexibility to analyze specimens ranging from in vitro to in vivo, with resolutions spanning from micrometers to nanometers [1,2]. An advantage of this technique is the ability to detect individual fluorescent targets and either simultaneously or sequentially visualize multiple fluorescent markers. Applications include the characterization of fluorescently labeled purified proteins, as well as the examination of proteins and genes within 2D/3D cell culture systems, tissues, and whole organisms. Fluorescence microscopy is utilized in various in vitro studies, such as investigating the liquid–liquid phase separation of proteins using fluorescently labeled purified proteins [3,4], examining spindle formation through droplet assays [5], and employing fluorescence resonance energy transfer (FRET) to analyze protein–protein interactions [6] and measure mechanical tension [7,8]. Its utility in developmental biology extends to research across a variety of models, including plants, Drosophila, Xenopus, C. elegans, zebrafish, and mice [9]. Moreover, fluorescence microscopy is a cornerstone in cell biology research, enabling the investigation of cellular processes in organisms from yeast to humans, across both 2D and 3D culture systems, as well as in living and fixed tissue samples [10,11]. This technique allows researchers to accurately determine spatial locations, relative localizations, dimensions, and shapes at the nanoscale. For instance, it has been demonstrated that standard confocal microscopy can achieve <5 nm precision in determining 2D/3D protein architecture within cells using a dual-color fluorescence approach [12,13]. Furthermore, fluorescence microscopy excels in quantifying protein copy numbers and assessing relative protein concentrations with high accuracy [14–16]. The resolution of fluorescence microscopy is constrained by the point spread function (PSF), which represents the 3D diffraction pattern of light emitted from an infinitely small point source [17]. An ideal fluorescence microscope, equipped with a high numerical aperture (NA) objective lens (NA > 1.4) and a high-resolution camera, can achieve resolutions of approximately 250 nm laterally and 600 nm axially [18,19]. However, the maximum resolution is also contingent on the signal-to-noise ratio (SNR) of the specimens. These resolution limits can be enhanced without altering the optical setup through expansion microscopy (ExM), which physically expands specimens without changing the optics setup, or through post-imaging processing techniques, such as deconvolution [18,20,21]. Additionally, ongoing developments in super-resolution microscopy significantly improve resolutions compared to conventional fluorescence microscopes, enabling the further detailed study of nano-scale cellular structures [1,22–24].
All fluorescence microscopes have the potential to achieve their theoretical resolution maximum and quantitative accuracy. However, this potential heavily relies on the components and the conditions under which microscopy is conducted. To understand the current conditions of the imaging system, it is critical to routinely perform calibrations. We have recently developed the 3D-Speckler software, which enables precise quantification of fluorescence puncta in both biological and non-biological samples [18]. Utilizing fluorescence beads of known sizes in conjunction with 3D-Speckler allows for the comprehensive evaluation of resolution limits, size quantification accuracy, chromatic aberration determination, and the flatness of illumination produced by imaging systems. The aim of this article is to present a comprehensive guide on utilizing the 3D-Speckler software for calibrating imaging systems. This guide is intended for a broad audience interested in fluorescence microscopy, rather than experts in optics and mathematics. Consequently, this protocol minimizes the explanation of the coding and mathematical principles behind 3D-Speckler. For detailed information, please refer to our original manuscript [18]. Briefly, 3D-Speckler identifies fluorescence particles primarily based on their relative signal intensities against the background and determines particle sizes through a full-width-at-half-maximum (FWHM) calculation based on a 2D or 3D Gaussian profile fitted to the intensity profile. The intensity measurements for fluorescence beads can be performed either without background correction or with a local background correction for each bead [14]. 3D-Speckler can determine the center of fluorophores with ~2 nm accuracy, which allows it to accurately identify chromatic aberrations, an unavoidable image distortion that occurs between different wavelengths, by measuring the distance between the centers of fluorophores with different wavelengths within a single bead. This process is designed to evaluate their current performance and ensure their integrity, accuracy, and reproducibility in quantification tasks.
Materials and reagents
TetraSpeck Fluorescent Microspheres Size kit (Thermo Fisher Scientific, catalog number: T14792) (see General notes 1–2)
Appropriate immersion medium [oil (e.g., Nikon, Type F), water (Mili-Q water), or silicone (e.g., Nikon, silicone immersion oil)] for the objectives you wish to calibrate
Equipment
Any type of fluorescence microscope (e.g., Nikon, model: Nikon-Ti2)
Example microscope setting: a Nikon Ti-2 inverted microscope equipped with a Yokogawa SoRa-W1 super-resolution spinning-disc confocal installed uniformizer, a high-resolution Hamamatsu Flash V3 CMOS camera, 4-line lasers (405, 488, 561, 640, 100 mW power), and a 60× or a 100× NA 1.4 oil objective
Image analysis PC [e.g., Dell, model: Precision 5820 Tower with Intel Core i7-9800X (3.8 GHz), Windows 10 Pro 64, 64 GB 2666 MHz DDR4, and Radeon Pro WX5100 8 GB]
Software and datasets
MATLAB (MathWorks, R2019-b and above, 10/1/2018, license required) (see General note 3)
3D-Speckler (https://github.com/suzukilabmcardle/3D-Speckler) (see General note 4) (07/20/2024, publicly available)
The source code, bfmatlab, user manual, and example test images are available at the following link: https://drive.google.com/drive/u/3/folders/1jKqiYFm31cJ0VVhGhLFhRuLI_ZiqmRjF (07/20/2024, publicly available)
Microscope control software (example: Nikon NIS Element, 04/05/2018, license required for NIS Element)
Procedure
This protocol mainly focuses on describing in detail the 3D-Speckler interface features for microscope calibration and thus offering a comprehensive, step-by-step workflow to operate it.
Obtaining images of fluorescent beads
For calibration with 3D-Speckler, we recommend utilizing fluorescent beads of at least two different sizes: one smaller than the point spread function (PSF) of your imaging system (less than 200 nm) and one larger (for example, 250 nm, 500 nm, or 1 μm). Smaller beads will be used to determine the lateral resolution limit of your system, which is equivalent to its PSF. Larger beads will be used to evaluate whether your system can accurately measure the size of beads larger than the PSF. In this protocol, we used TetraSpeck beads of 100 and 500 nm. 3D-Speckler can be used for both 2D and 3D images of fluorescence beads obtained from any fluorescence microscope. Additionally, 3D images can be utilized for 2D calibration (see details in our original manuscript [18]). Axial size measurement and 3D chromatic aberration correction require 3D stack images.
Power on the microscope system and select the objectives you plan to use for calibration.
Place a slide with TetraSpeck beads (100 nm) on the microscope stage.
Locate a field of view (FOV) that contains TetraSpeck beads. We recommend finding a FOV containing fluorescence beads that are evenly distributed and not aggregated or too crowded. This helps minimize the difficulty and errors in identifying individual beads and taking measurements. Then, finely adjust the focus, the camera's exposure time, and the light source's power for optimal imaging (see General note 5).
(Optional) Set the z-range to encompass the entire depth of the beads, ensuring that the interval between each step is less than 200 nm (see General note 6).
Acquire images at the wavelengths you wish to calibrate. It is recommended to capture images at different wavelengths at the same z-plane for chromatic aberration measurements.
Repeat steps A2–A5 at several different locations on the slide to ensure thorough calibration (see General note 7).
Using a slide with larger TetraSpeck beads (for example, 500 nm), repeat steps A2–A6 (Figure 1).
Figure 1. Example of a TetraSpeck bead image. The 500 nm TetraSpeck beads were imaged using a 60× objective lens. The full field of view (FOV) image is displayed on the left. On the right, the XY and XZ views of a single 500 nm bead from the left image are presented. Visualization of XY and XZ views using Imaris software (Andor).
Workflow of 3D-Speckler
3D-Speckler interface features
Figure 2 displays the primary user interface of 3D-Speckler. A detailed description of each module is listed below.
Figure 2. Main interface of 3D-Speckler. This figure displays the primary user interface of the 3D-Speckler software.
Open File: Enables the import of new image files (e.g., *.stk, *.nds, *.tif) for analysis (see General note 8).
Import Data: Users can import data previously analyzed for further visualization (*.xlsx, *.xls).
Import CA (chromatic aberration) Calibration: Facilitates the import of polynomial calibration surfaces or affine transformation calibrations to correct chromatic aberrations.
Analysis Options: Offers several options for local background correction, exclusion of overlapping particles, and by-size particle filtering.
Bounding Box Buffer: Users can adjust the bounding box padding for 2D/3D analysis, but the default value (0.3) should suffice for most fluorescence bead analyses.
Particle Size Filter: Allows users to filter particles for analysis by setting a range of lateral pixel sizes.
Commit Current ROI (Region of Interest): Enables the analysis of selected areas within a larger image, with the option to reset the ROI to the original image dimensions.
Channel Choice: Provides options for users to select channels to analyze or visualize.
Image Viewer: Visualize the current image or image stack under analysis.
Contrast Adjustment: A pop-up window can enable users to adjust contrast to enhance visualization.
Reset View: Resets the image zoom to the max outward level.
Package Option: Provides options for users to select an analysis pipeline, such as Particle Analysis or Chromatic Aberration Calibration (see General note 9).
Start Analysis: Opens and displays the bounding boxes for detected particles at the selected threshold, offering single or multi-thresholding.
Fit Gaussians: Performs the 2D/3D Gaussian fitting at the specified threshold and displays the fitted particles with localizations and bounding boxes.
Fitting Options: Offers a choice between 2D and 3D Gaussian fitting for users. The 2D option is compatible with both 2D and 3D images, providing measurements of detected objects at their optimal focus plane when applied to 3D images.
Match & Align Points: Performs matching of corresponding fluorescent particles across different channels after their analysis for further examination (see General note 10).
Distance Matrices: Generates distance matrices for every particle set within the same or other channels.
Aberrations: Facilitates the characterization and generation of polynomial calibration surfaces or affine transformations for calibrating between different channels.
Save Channel Results: Saves analysis results for the current channel, visualized when switching between channels.
Export Results: Exports all analysis parameters to an Excel sheet and a folder, including a reference image and aberration calibration.
Measurements of size below and above PSF, illumination flatness, and chromatic aberration
Set the MATLAB working directory to the location that includes the “bfmatlab” folder and the 3D-Speckler script.
Use the command “Open3DSpeckler” in the command window to run the 3D-Speckler or click Run (see General note 11).
Select a bead image file in 3D-Speckler. Users can choose image stacks that are either single-wavelength or multi-wavelength (see General note 12).
For multi-wavelength images, select the appropriate channel wavelengths and adjust the image scales. If users select an image that is single-wavelength, 3D-Speckler will prompt users to decide whether to add more wavelength images (Figure 3) (see General note 13).
Figure 3. Selecting image wavelengths. Upon selecting an image file, it will be displayed. You will then need to choose the wavelengths and review both lateral and axial scales.
The main user interface will launch upon clicking choose. After completing this step, the 3D-Speckler base interface will appear (Figure 4), allowing you to select either Particle Analysis or Local Chromatic Aberration Correction for downstream analysis.
Figure 4. 3D-Speckler main interface. After completing wavelength selection, this interface will be displayed. You will need to select either Particle Analysis or Local Chromatic Aberration Correction for downstream analysis.
Choose the Particle Analysis pipeline from the top right-hand corner (see General note 14).
Select your preferred analysis options from the panel on the middle left-hand side (refer to feature d). When you choose the Local Background Correction option in 3D-Speckler, the software will subtract local background signals from each detected object (see General note 15). 3D-Speckler performs local background correction using a user-defined percentage for a larger bounding box (default is 15% larger in all dimensions) with automated BG exception handling. The Exclude Overlapping Particles feature ensures that particles with overlapping bounding boxes are removed to avoid measurement inaccuracies. The Sort by X < Y < Z FWHM option organizes results in an Excel file based on the full width at half maximum (FWHM) values for the X, Y, and Z axes. Before starting the analysis, confirm the ROI by using the zoom button to select a specific area if you aim to analyze a particular region (refer to feature g).
To adjust the image contrast, utilize the Contrast feature (refer to feature j). For Z-position adjustments, slide the bar situated to the right of the image. To choose a specific wavelength, click on the Channel feature (refer to feature h).
Begin by clicking on Start Analysis to set the optimal detection threshold (Figure 5). This allows users to view bounding boxes, which can be adjusted according to the thresholding bar or via the bounding box buffer fields (refer to feature e). Once the ideal threshold is determined, confirm by clicking SET THRESHOLD (see General note 16).
Figure 5. Thresholding bar. It pops up when you click Start Analysis. Users can adjust the threshold by sliding the bar to the right or left.
Select the fitting options (either 2D Gauss Fit or 3D Gauss Fit; refer to feature o) and click on Fit Gaussians to fit particles.
(Optional) If you would like to perform local background correction, you need to select the parameters below (Figure 6) (see General note 15).
Figure 6. Background correction buffer. This option becomes available after you click on Fit Gaussians. If you wish to perform background correction, please select the size of the bounding box for background measurements. Choosing 1 will double the dimensions of the bounding boxes used for quantification (see General note 15).
Review and manage detected particles: To remove unwanted particles, first click the Select button found beneath the Fit Gaussians button. Then, within their bounding boxes, double-click on the particles you wish to eliminate (Figure 7). Once you have reviewed the detected objects, click on Save Channel Results to save the data for the current channel. Ensure you save your data before moving to the next channel; otherwise, unsaved measurement results will be lost.
Figure 7. Review fitting results. After fitting, particle locations will be shown in blue, not green. Click the select button located under the Fit Gaussians button, and then double-click on particles within their bounding box to remove them.
Repeat steps B2h–B2l to analyze each channel.
After completing the particle analysis for all channels, select Match & Align Pts (refer to feature p) to match corresponding particles and align all channels (Figure 8). This step is optional for users who only require measurements of size and intensities. The Match & Align Pts feature identifies corresponding beads across various wavelengths, enabling the comparison of measurement values from different wavelengths within the same TetraSpeck beads. However, this process will eliminate detected particles that do not match. Utilizing this option is essential for further analysis of chromatic aberrations.
Figure 8. Matching and aligning detected particles across all channels. The Match & Align Pts function enables the selection of corresponding particles across different channels, eliminating those that do not match. The white arrow indicates a particle unmatched in the Blue channel, resulting in its removal from quantification in all channels, as shown by the Red arrows.
To measure chromatic aberrations, clicking Aberrations (refer to feature r) will open the wavelength selection tab. Choose the wavelength at which you wish to determine aberrations, and then click Generate (see General note 17).
Select Export Results (refer to feature t). 3D-Speckler will prompt you to decide if you wish to include Field Dependence Plots in your export. If you are interested in assessing the uniformity of illumination, please choose Yes. Then, specify the desired location to save the files. Upon doing so, multiple plots will be displayed, followed by a confirmation message from 3D-Speckler indicating “Data Export Completed Successfully.”
Reviewing results
3D-Speckler will create a new folder that includes an Excel file, MATLAB images for each channel, and a subfolder containing field-dependent plots of FWHM and signal intensities. The MATLAB images of beads will feature assigned numbers for each detected bead, corresponding to the numbering in the Excel file.
To review and obtain the 2D and 3D size of TetraSpeck beads, open the Excel file (Figure 9). 3D-Speckler offers three distinct methods for measuring FWHM: true (tFWHM), interpolated (iFWHM), and Gaussian FWHM (gFWHM). The differences lie in the methods used to obtain the intensity profile of detected particles for measuring FWHM. Detailed descriptions of these measurements can be found in our original article [18]. Briefly, gFWHM and iFWHM use Gaussian and interpolation fitting methods, respectively, whereas tFWHM does not employ mathematical fitting methods. We recommend using tFWHM for lateral measurements and iFWHM for axial measurements [18]. The average lateral size (tFWHM) can be determined by averaging the values of the blue columns in the Excel file (Column Q and R in an Excel file in 3D measurements). Similarly, the axial size (iFWHM) can be calculated by averaging the values in the black column (Column V in an Excel file in 3D measurements) (see General note 18).
The lateral and axial sizes of beads smaller than the PSF represent the resolution limit of your system (measured PSF in your system). You can compare and evaluate these values against the theoretical resolution limit of your system. The measured size of beads larger than the PSF should be equal to or close to their actual size. If not, it may indicate that the bead size you used is still smaller than the PSF of your system or suggest potential defects in your system.
Figure 9. Quantification results. Example results of the green channel. The output is in Excel file format. Abbreviations’ definitions are listed in Table 1.
Table 1. Definition of the abbreviations in Figure 9. Explanation of the abbreviations in the output Excel file.
Abbreviation Explanation
Gauss_X, _Y, -Z x, y, and z coordinates of the center position of detected particles.
Theta_X, _Y, _Z x, y, and z angles based on the optimum center of detected particles, as described in the original 3D-Speckler manuscript.
STD_X, _Y, _Z Standard of deviation of Gaussian fitting.
GaussBase The base signal intensity of Gaussian fitting. If users select local background correction option, this value is used for local background correction.
GaussPeak The peak signal intensity of Gaussian fitting.
tFWHM_X, _Y, _Z tFWHM values of each detected particle in x, y, and z-axis.
iFWHM_X, _Y, _Z iFWHM values of each detected particle in x, y, and z-axis.
gFWHM_X, _Y, _Z gFWHM values of each detected particle in x, y, and z-axis.
ResNorm Normalized residual value: how well the objects fit the 2D/3D Gaussian model.
3D-Speckler measures integrated and maximum signal intensities for each bead [the red columns (O and P) in Figure 9]. Using this information, 3D-Speckler can generate field dependence plots that illustrate variations in the FWHM measurements and intensity across the FOV of an imaging setup. Users will have the option to export these plots. If chosen, 3D-Speckler will save the plots as .fig files in the Field Dependence Plots folder. These plots can then be analyzed to identify areas within the field of view that provide the most accurate measurements (Figure 10).
Figure 10. Visualization of field dependency plots. Surface plots of integrated intensity and maximum intensity variations illustrate changes across the imaging field of view (FOV). The light source in fluorescence microscopy is typically brightest at the center of illumination and gradually becomes dimmer toward the edges.
When the Aberrations case (step B2o) is executed, the Excel file contains chromatic aberration measurements across the selected wavelengths. These results are presented in the worksheet titled “aberration_ wavelength” in the Excel file (Figure 11 and General note 19). The file includes aberration data for each axis and calculates the 2D/3D distances for individual beads. To obtain average aberration values and distances, users can average these data points.
Figure 11. Aberration between different channels. The Excel file contains the worksheets named “aberration_wavelength”. These worksheets include aberration in x, y, and z-axis and 3D distance measurements (µm scale) (see General note 19-20).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article
Loi, J. et al. (2023). Semi-automated 3D fluorescence speckle analyzer (3D-Speckler) for microscope calibration and nanoscale measurement. J Cell Biol. (Figures 1–8).
General notes and troubleshooting
General notes
Any fluorescence bead slide can be utilized; however, we recommend using beads of at least two sizes: one smaller than the PSF (< 200 nm) and one larger than the PSF (e.g., 250 nm, 500 nm, and 1 µm). We observed that the mounted beads on slides tend to shrink over time. Therefore, we advise using fresh beads or those used shortly after purchase. Commercial or homemade bead slides should be stored at 4 °C and protected from light.
Fluorescent bead calibration slides can be made by yourself.
For users who do not have access to a MATLAB license, we provide a standalone, executable application made through MATLAB compiler. The standalone 3D-Speckler and user manual are available at the following website (Windows: https://drive.google.com/drive/u/3/folders/1iV5AbqgJhIkQV2lAoZyrNN0BRRq5_JrX and Mac: https://drive.google.com/drive/u/3/folders/1wrVNDN-6H0QAMgidj0Ql_XTvt44GGkFR.). Running standalone 3D-Speckler requires installation of the correct version of MATLAB Runtime, thus we provided an installer for the correct version in the same download folder.
3D-Speckler can be installed on both Windows and Mac OS with MATLAB (2019 and above).
We recommend obtaining images with high SNR, ideally > 20–50, to calibrate the optimal performance of your system. However, the specific SNR requirements may vary depending on your system. You can test the effects of SNR on your system's performance by preparing bead images with different SNRs, which can be achieved by adjusting the light source power and/or camera exposure time.
We recommend imaging with 100 or 200 nm steps.
Since calibrating with a single 2D/3D bead image carries risks, we recommend using multiple images taken from several different locations.
3D-Speckler uses the Bio-Formats (OME) software tool, which can open most image file formats [25].
Protein Quantification is under development and currently unavailable.
Unmatched points will be removed from further analysis.
Executing 3D-Speckler by clicking Run does not require changing the working directory to include the “bfmatlab” folder. However, to use the “Open3DSpeckler” command for operation, it is necessary to change the working directory.
3D-Speckler automatically reads and categorizes images into different channels. If only one single is detected, it allows users to import data for more channels.
3D-Speckler automatically detects the lateral and axial resolution by using the image metadata. If these resolutions are not automatically filled in or the numbers have errors, users must manually input or correct the scaling values.
The 3D-Speckler interface automatically updates based on the selected analysis options and workflow. Particle Analysis is the default analysis method for analyzing general fluorescent particles.
For advanced settings, 3D-Speckler provides a Local Background Correction option for signal intensity and Gaussian fitting measurements. Since background signals are not uniform across the FOV, local background correction emerges as one of the most accurate measurement methods [14,15,18]. To utilize this feature, users need to optimize parameters for further analysis. The default is set at 0.15 (which is 15% larger in all dimensions than the bounding box used for measurements in each detected particle to obtain local background signals). For most analyses involving TetraSpeck beads, this default setting proves effective.
For an additional optimization step, by clicking on multi-thresholding, users can select a threshold to commit an image. 3D-Speckler will then remove those objects, allowing users to set another threshold until all desired objects are accurately detected. Conclude by clicking on Finish Thresholding.
The Distance Metrics option offers 2D or 3D distance measurements between all detected beads, both within a single channel and across different channels [18]. This feature is not necessary for microscope calibration.
3D-Speckler offers three different measurement methods for FWHM. Please see the details in the original article [18].
This measurement boasts a precision of less than 5 nm [18].
3D-Speckler provides correction for chromatic aberration and outputs the corrected images, as detailed in the original article [18].
Acknowledgments
The 3D-Speckler was primarily developed by Jonathan Loi under the guidance of Aussie Suzuki (McArdle Laboratory for Cancer Research, Department of Oncology, University of Wisconsin-Madison). The original work was published in Loi et al., Journal of Cell Biology, 2023 [18]. Part of this work is supported by Wisconsin Partnership Program, the University of Wisconsin-Madison Office of the Vice Chancellor for Research with funding from the Wisconsin Alumni Research Foundation (Research Forward), start-up funding from University of Wisconsin-Madison SMPH, UW Carbone Cancer Center, and McArdle Laboratory for Cancer Research, and NIH grant R35GM147525 (to A.S.).
Competing interests
The authors declare no conflicts of interest within this work.
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Loi, J., Qu, X. and Suzuki, A. (2023). Semi-automated 3D fluorescence speckle analyzer (3D-Speckler) for microscope calibration and nanoscale measurement. J Cell Biol. 222(4). https://doi.org/10.1083/jcb.202202078.
Liu, W. J., Toussaint, K. C., Okoro, C., Zhu, D. Z., Chen, Y. H., Kuang, C. F. and Liu, X. (2018). Breaking the Axial Diffraction Limit: A Guide to Axial Super-Resolution Fluorescence Microscopy. Laser Photonics Rev. 12(8). https://doi.org/10.1002/lpor.201700333
Chen, F., Tillberg, P. W. and Boyden, E. S. (2015). Optical imaging. Expansion microscopy. Science. 347(6221): 543–548. https://doi.org/10.1126/science.1260088
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Article Information
Publication history
Received: Apr 9, 2024
Accepted: Jul 4, 2024
Available online: Jul 24, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Cell Biology > Cell imaging > Fluorescence
Biophysics > Microscopy
Computational Biology and Bioinformatics
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5,052 | https://bio-protocol.org/en/bpdetail?id=5052&type=0 | # Bio-Protocol Content
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Protocol for Imaging the Same Class IV Neurons at Different Stages of Development
SS Sonal Shree
JH Jonathon Howard
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5052 Views: 461
Reviewed by: Nafisa M. JadavjiEhsan Kheradpezhouh Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Jun 2022
Abstract
In this protocol, we focused on analyzing internal branches of Drosophila class IV neurons. These neurons are characterized by their highly branched axons and dendrites and intricately tile the larval body. As Drosophila larvae progress through developmental stages, the dendritic arbors of Class IV neurons undergo notable transformations. As Drosophila larvae develop, their Class IV dendritic arbors grow. In the initial 24 h after egg laying (AEL), the dendrites are smaller than segments. During the subsequent 24 h of the first instar larval stage, dendritic arbors outpace segment growth, achieving tiling. After 48 h, arbors and segments grow concurrently. Epidermal cells near Class IV dendrites expand in proportion to segment growth. This observation suggested that Class IV cells might grow via branch dilation—uniformly elongating branches, akin to Class I cells [1,2]. To understand whether the class IV complex arbor structure is formed by dilation or simply from growing tips, we developed this protocol to introduce a systematic approach for quantitatively assessing the growth dynamics of internal branches.
Key features
• This protocol employs imaging the same neuron over different development times
• Drosophila embryo and larvae genotype is ;;ppkCD4-tdGFP, which explicitly tags class IV neurons
• This protocol for the preparation of agar pads to mount and image Drosophila larvae is adapted from Monica Driscoll's method
• Neurons are imaged without the use of anesthetics and for a short duration of time
• This technique involves the use of a spinning disk confocal microscope
Keywords: Class IV neurons Branching morphogenesis Neuronal development Dendrite Drosophila larvae Tip growth Confocal microscopy
Graphical overview
Schematic overview of the process for mounting larvae on an agar pad for imaging using a confocal microscope
Background
Dendrites serve as extensions of neurons and play a pivotal role in receiving signals from other neurons or the surrounding environment. These extensions often have a branched structure, which enhances their capacity to gather information. An excellent model for exploring the development of dendrites is found in the extensively branched Drosophila Class IV neurons. These neurons exhibit a wide array of shapes in their dendritic structures, with each dendrite being unique. This suggests that the growth of dendrites is more likely a random process rather than a predetermined one.
Despite this diversity, the specific underlying mechanisms guiding the development of dendrites in these neurons remain largely unclear. The larval body is divided into segments, and each segment harbors a pair of class IV neurons. These neurons, in our hand, are first visible around 16 h after egg laying (AEL). At this stage, neurons are small as compared to their harboring segment, but they grow fast and catch up with growing segments. After that, neurons grow in proportion to the expansion of the segments [3]. This has led to the hypothesis that Class IV cells expand through branch dilation—that is, by uniform expansion of branches along their lengths [4]. Such dilation occurs in Class I cells [1]. To form the highly branched late-stage dendrite, dilation would need to be complemented by the formation of new branches on extant ones, to infill the structure.
Through our investigation, we have made a noteworthy observation: dendritic branches lengthen primarily through extension from their tips, rather than by expanding their non-terminal branches. We developed this protocol to image the same neurons over developmental time without the use of anesthetics. Our focus was to image neurons for short durations and take static images, placing larvae back in apple agar plates in the Darwin chamber at 25 °C so that neurons and larvae do not get stressed by continuous imaging and use of anesthetics. The LarvaSPA method [5] allows imaging neurons continuously for 10 h; however, our goal was to image the same neuron every 24 h at various developmental time points.
Materials and reagents
Biological material
Fly line ;;ppk-cd4-tdGFP (homozygous) was used to image class IV dendritic arborization neurons and was a gift from C. Han (Cornell University)
Reagents
Apple juice unsweetened frozen concentrate (4×) [Stop & Shop (grocery store), 12 oz can]
Dextrose (Sigma-Aldrich, catalog number: G5767)
Sucrose (Sigma-Aldrich, catalog number: 57-50-1)
Bacto agar (Becton Dickinson, catalog number: 214010)
NaOH pellet (Sigma-Aldrich, catalog number: 221465)
Propionic acid (Sigma-Aldrich, catalog number: 402907)
Phosphoric acid (Sigma-Aldrich, catalog number: 695017)
Halocarbon oil 700 (Sigma-Aldrich, catalog number: H8898)
Agarose (AmericanBio, catalog number: AB9012-36-6)
PBS 10× solution (AmericanBio, catalog number: AB11072-01000)
Laboratory supplies
High-precision microscope cover glass 22 × 22 mm (VWR International, catalog number: 16004-302)
35 mm dish, No. 1.5 coverslip 10 mm glass diameter uncoated (MatTex, catalog number: P35G-1.5-10-C)
Mini embryo collection cages (Flystuff.com) (Genesee Scientific, catalog number: 59-105)
Tissue culture dish, 35 × 10 mm style (Falcon, catalog number: 353001)
Tray of 100 NARROW 1-oz PS vials, each with 10 mL of glucose media, pre-loaded with cellulose-acetate plugs (Archon Scientific, catalog number: D20102)
Frosted micro slides (VWR, catalog number: 48312-004)
Equipment
For imaging, samples were mounted on the microscope stage, illuminated with Nikon lasers (488 nm at 18%–21% laser power), and imaged on a spinning disk microscope: Yokogawa CSU-W1 disk (pinhole size 50 μm) built on a fully automated Nikon TI inverted microscope with perfect focus system, an sCMOS camera (Zyla 4.2 plus sCMOS), and Nikon Elements software with either a 40× (1.25 NA) or 20× (0.5 NA) water immersion objective.
Software and datasets
Images were stitched and analyzed using Fiji (NIH, 2021). Detailed procedure of stitching is mentioned in the procedure section. https://imagej.net/software/fiji/
NIS-Elements Imaging Software (Nikon, 2022 This imaging software comes with the spinning disk microscope. It is an interface that enables the user to capture images using Nikon microscope and also helps to export the raw images in the TIFF format, which could later be processed by FIJI for stitching
Prism 9 (GraphPad Prism, 2023). https://www.graphpad.com/scientific-software/prism/
Procedure
This protocol for the preparation of agar pads to mount and image Drosophila larvae is adapted from Monica Driscoll's method (https://www.wormatlas.org/agarpad.htm) [7].
Preparing an agar pad for larvae imaging
Measure and dissolve agar in water to create a 5% agar solution. Utilize a heat block to maintain the solution's molten state at a steady 65 °C. This temperature ensures the agar remains in a liquid state for the subsequent steps.
Application of agar solution: Using scissors, cut the tip of a 1000 mL pipette to allow controlled dispensing. Pipette 100–200 mL of molten agar onto the center of a pristine glass slide, ensuring a smooth and even distribution.
Formation of the agar pad: Place another clean glass slide perpendicular to the one with the agar drop, creating a "sandwich" effect. Gently press the top slide to flatten the agar, forming a circular pad. Pay close attention to avoid trapping air bubbles within the agar during this process, as they can interfere with imaging.
Optimal technique: For a more precise agar pad suitable for imaging larvae, follow Monica Driscoll's method. This technique involves using two additional taped slides, ensuring a flat surface without any irregularities that might hinder the larvae's movement or positioning.
Solidification and separation: Allow the agar to solidify naturally. Carefully separate the slides, ensuring that the agar pad adheres to one of them without any damage or distortion.
Timely preparation: It is crucial to prepare the agar pad just before use to maintain its optimal consistency and prevent it from drying out, which could affect its efficacy as a base for imaging.
Halocarbon application: Before mounting the larvae for imaging, add a drop of Halocarbon 700 onto the agar pad. Spread the halocarbon evenly across the surface to ensure proper lubrication and facilitate smooth movement of the larvae during imaging.
Chilling for optimal performance: To further enhance the agar pad's performance, chill it at 4 °C for approximately 5 min before mounting the larvae. This step can assist in achieving better adherence and stability.
Mounting of live larvae
Preparation of larvae: Wash the larvae sequentially with 20% and then 5% sucrose solutions to ensure cleanliness and hydration.
Positioning on agar bed: Gently place the larvae, dorsal side up, onto a 5% agar bed secured on a glass slide. Create an imaging environment by adding a drop composed of 50% PBS and 50% Halocarbon oil 700.
Immobilization technique: Further immobilize the larvae by applying gentle pressure with a 22 × 22 mm coverslip lined with Vaseline or vacuum grease.
Temperature optimization: Allow the setup to rest at 4 °C for 5 min to enhance stability.
Optional temperature control for reduced movement: Optionally, minimize larval movement during imaging by maintaining a temperature of 4 °C for 2–5 min. Employ an OKO lab temperature control module connected to a spinning disk confocal microscope (Nikon) for this purpose.
Image acquisition: Samples were imaged using a sophisticated spinning disk microscope setup, incorporating a Yokogawa CSU-W1 disk with a 50 μm pinhole size. The agar pad setup was mounted on a fully automated Nikon TI inverted microscope, equipped with perfect focus capabilities. For illumination, a 488 nm laser was used at a power setting of 18%–21%. The imaging was performed using either a 40× water immersion objective or a 20× air objective. The detection system included an sCMOS camera (Zyla 4.2 plus) and was controlled using Nikon Elements software.
Prior to image acquisition, samples were manually focused to accurately identify the abdominal third and fourth segments (A3 or A4 neurons). Static images intended for morphometric studies were captured using the 40× water immersion objective for larvae that were 24 h AEL, while the 20× objective was utilized for imaging larvae at later stages (such as 48, 72, and 96 h AEL). To create comprehensive images, the individual images were stitched together using the stitching plugin available in ImageJ software [6].
Post-imaging procedure: Return the imaged larvae to an apple agar plate containing yeast paste within a Darwin chamber set at 25 °C. Conduct a subsequent imaging session after 24 h, allowing for the continuous observation of the same class IV neuron throughout larvae development without the use of an anesthetic.
Protocol for stitching microscope images using FIJI plugin
This protocol is used to stitch microscope images using the FIJI plugin's "Stitch Directory with Images (unknown configuration)" under the Deprecated section. This process is suitable for combining multiple images into a single cohesive image, especially when the configuration of the images is not known beforehand.
Requirements: FIJI (Fiji Is Just ImageJ) software installed and a directory containing microscope images to be stitched.
Steps:
Open FIJI: Launch the FIJI software on your computer.
Prepare your images: Ensure all the images you want to stitch are in a single directory.
Verify that the images have consistent naming conventions for easier processing.
Access the stitching plugin: Go to Plugins in the menu bar. Navigate to Stitching and then select Stitch Directory with Images (unknown configuration). Select the Image Directory:
A dialog box will appear prompting you to select the directory containing your images (Figure 1).
Figure 1. Screenshot of FIJI Software highlighting access to the stitching plugin
Click Browse or Choose and navigate to the folder where your images are stored.
Select the directory and click OK (Figure 2).
Figure 2. Snapshot of FIJI Stitching window demonstrating access to the image directory
Fusion Method: Choose the method for merging images; typically, "Linear Blending" or "Max Intensity" work well.
Start stitching: Once all parameters are set, click OK to begin the stitching process.
FIJI will process the images; this may take some time depending on the number and size of images.
Review the stitched image: After the process is complete, the stitched image will be displayed. Inspect the image for any stitching errors or misalignments. If adjustments are necessary, you can tweak the parameters and repeat the process.
Save the stitched image: Once satisfied with the stitched result, go to File > Save As.
Choose the desired file format and location to save your stitched image.
Post-processing (Optional): Perform any additional image processing if required, such as cropping, adjusting contrast, or applying filters using FIJI.
Tips: For best results, ensure your images have consistent exposure and focus.
If the images have significant overlap, reduce the overlap percentage to speed up processing.
Save intermediate results periodically to avoid data loss in case of software crashes.
By following these steps, you should be able to successfully stitch microscope images using the FIJI plugin, even when the configuration of the images is unknown (Figure 3).
Figure 3. Overview of stitching microscope images using the FIJI stitching plugin. In this example, four side images are taken using the soma of a neuron as the reference point. The images are then stitched together to generate a complete view of the neuron.
After stitching, segment/crop the neuron using FIJI’s Image and then Crop function.
Validation of protocol
This protocol was used to visualize the same class IV neuron (A3 and A4 segments) at discrete times over development, at 24 and 48 h and at 48 and 96 h, to test whether arbor expansion of neurons is by dilation of internal branches or tip growth and infilling. The larvae were mounted on an agar pad and imaged as described above, without using anesthetics. To minimize movement, imaging was performed at 4 °C for 2–5 min. After imaging, the larvae were returned to an apple agar plate in the Darwin chamber. The imaging was conducted using 20× and 40× objectives.
For image analysis, the same neurons imaged at 24 and 48 h (Figure 4) were segmented (FIJI/Image/Crop) and aligned (FIJI/Image/Rotate) using ImageJ software. This alignment allowed us to identify conserved non-terminal internal branches in the proximal region. This detailed approach enabled a precise study of the potential role of internal branch elongation in arbor growth. The fractional increases in branches and segment lengths were defined as
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Figure 4. The same neuron imaged at 24 h (A) and 48 h (B) after egg laying (AEL) was segmented and aligned using ImageJ. Conserved internal branches were identified by overlaying the aligned images over each other using FIJI image tab and overlay tool (C) and marked with the same color-coded dotted lines and corresponding numbers (D&E). The length was measured using FIJI’s segmented line tool. The table on the right displays the lengths of these conserved internal or non-terminal branches along with their fractional length changes.
The detail of data analysis of this method is also described in the Result section “Dendrite growth is not due to elongation of all branches in the arbor” and Figure 2 of Shree et al., 2022 [6]. Our measurements of internal branches of the same class IV neurons over developmental time using an agar pad method were also consistent with earlier assessments done by Baltruschat et al. [8].
General notes and troubleshooting
This protocol was designed specifically to visualize the consistent observation of class IV neurons located on the dorsal side of Drosophila larvae at discrete developmental stages (24, 48, and 96 h after egg laying). The focus was solely on class IV neurons within the A3 and A4 abdominal segments.
Image the larvae for not more than 2–5 min as long-term imaging could stress larvae, which might halt their growth.
Following imaging sessions, it is crucial to promptly return the larvae to their apple agar plates and transfer them back into Darwin incubators set at 25 °C. This swift return process aids in maintaining the natural developmental conditions for the larvae. Use a paintbrush to pick larvae instead of tweezers for gentle handling, as Class IV neurons are highly sensitive to mechanical pressure; excessive pressure can halt their growth and lead to neuron degeneration. During imaging, larvae were immobilized with minimal pressure under a coverslip and positioned on an agar pad.
Emphasize speed: To preserve the integrity of observations and ensure consistency in developmental stages, prioritize the quick and efficient return of larvae to the apple agar plates post-imaging.
Acknowledgments
This study was supported by NIH grants DP1 MH110065 and R01 NS118884 awarded to Jonathon Howard. The abbreviated version of this protocol is published in Science Advances, Jun 2022. DOI: 10.1126/sciadv.abn0080.
Competing interests
The authors declare that they have no competing interests.
References
Palavalli, A., Tizón-Escamilla, N., Rupprecht, J. F. and Lecuit, T. (2021). Deterministic and Stochastic Rules of Branching Govern Dendrite Morphogenesis of Sensory Neurons. Curr Biol. 31(3): 459–472.e4. https://doi.org/10.1016/j.cub.2020.10.054
Ferreira Castro, A., Baltruschat, L., Stürner, T., Bahrami, A., Jedlicka, P., Tavosanis, G. and Cuntz, H. (2020). Achieving functional neuronal dendrite structure through sequential stochastic growth and retraction. eLife. 9: e60920. https://doi.org/10.7554/elife.60920
Parrish, J. Z., Xu, P., Kim, C. C., Jan, L. Y. and Jan, Y. N. (2009). The microRNA bantam Functions in Epithelial Cells to Regulate Scaling Growth of Dendrite Arbors in Drosophila Sensory Neurons. Neuron. 63(6): 788–802. https://doi.org/10.1016/j.neuron.2009.08.006
Yang, W. K. and Chien, C. T. (2019). Beyond being innervated: the epidermis actively shapes sensory dendritic patterning. Open Biol. 9(3): e180257. https://doi.org/10.1098/rsob.180257
Ji, H. and Han, C. (2020). LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging. J Visualized Exp. 156: e60792. https://doi.org/10.3791/60792
Shree, S., Sutradhar, S., Trottier, O., Tu, Y., Liang, X. and Howard, J. (2022). Dynamic instability of dendrite tips generates the highly branched morphologies of sensory neurons. Sci Adv. 8(26): eabn0080. https://doi.org/10.1126/sciadv.abn0080
Driscoll, M. (2006). Mounting Animals for Observation with Nomarski DIC Optics. In Shaham, S. (Eds), In Methods in cell biology WormBook, 1551–8507. https://www.ncbi.nlm.nih.gov/books/NBK19784/
Baltruschat, L., Tavosanis, G. and Cuntz, H. (2022). A developmental stretch-and-fill process that optimises dendritic wiring. bioRxiv. 2020.07.07.191064. https://doi.org/10.1101/2020.07.07.191064.
Article Information
Publication history
Received: Jun 30, 2024
Accepted: Jul 12, 2024
Available online: Aug 2, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Developmental Biology > Morphogenesis > Cell structure
Neuroscience > Cellular mechanisms > Neuronal fate
Cell Biology > Cell imaging > Confocal microscopy
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5,053 | https://bio-protocol.org/en/bpdetail?id=5053&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Simple Analysis of Gel Images With IOCBIO Gel Software
LJ Lucia Jaska
RB Rikke Birkedal
ML Martin Laasmaa
MV Marko Vendelin
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5053 Views: 481
Reviewed by: Marcelo S. da SilvaThaise Lara Teixeira Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in BMC Biology Oct 2023
Abstract
Gel image analyses are often difficult to reproduce, as the most commonly used software, the ImageJ Gels plugin, does not automatically record any steps in the analysis process. This protocol provides detailed steps for image analysis using IOCBIO Gel software with western blot as an example; however, the protocol is applicable to all images obtained by electrophoresis, such as Southern blotting, northern blotting, and isoelectric focusing. IOCBIO Gel allows multiple sample analyses, linking the original image to all the operations performed on it, which can be stored in a central database or on a PC, ensuring ease of access and the possibility to perform corrections at each analysis stage. In addition, IOCBIO Gel is lightweight, with only minimal computer requirements.
Key features
• Free and open-source software for analyzing gel images.
• Reproducibility.
• Can be used with images obtained by electrophoresis, such as western blotting, Southern blotting, isoelectric focusing, and more.
Keywords: Data analysis Reproducibility FAIR Western blotting Southern blotting Isoelectric focusing Python Electrophoresis gel
Background
When analyzing signal intensity from images, for example from western blots, researchers mainly use either proprietary software or the ImageJ Gels plugin [1]. Unfortunately, using those programs for image analysis contributes to a reproducibility crisis [2]. Proprietary software is closed source, so users do not know exactly how the analysis is performed, and the results can only be reproduced using the same software. While the ImageJ Gels plugin is open source, it does not automatically document the steps in the analysis process, leading to decreased reproducibility. IOCBIO Gel [3] solves both of these problems—it is free and open source, and it also records the original image together with all analysis steps performed on it, such as cropping, background subtraction, and lane selection. This makes it easy to reproduce the results and make corrections at any analysis stage. In addition, the software can store the data either on a PC or in a central database, facilitating efficient collaboration between researchers. Taken together, IOCBIO Gel is an efficient tool for image analysis, contributing to the implementation of FAIR (findability, accessibility, interoperability, and reusability) principles [4]. Further information can be found in the original article describing the software and testing its performance against the ImageJ Gels plugin [3].
Equipment
Minimal requirements are mainly imposed by the corresponding operating system. The requirements of the application itself are minimal
Software and datasets
IOCBIO Gel
Homepage: https://iocbio.gitlab.io/gel/
Downloads: https://gitlab.com/iocbio/gel/-/releases (version 1.0.3, release date July 11, 2024)
Video tutorials on YouTube, links also available on homepage
Installation
First start and setup
Basic tutorial
Advanced tutorial
Procedure
Installation
Linux/Mac
To use the automatic installation script for Linux/Mac, first make sure that you have the latest pip installed by running:
python3 -m pip install --user --upgrade pip
Then, open a terminal and go to the folder where you want to install the program. Then, run the following command:
curl https://gitlab.com/iocbio/gel/-/raw/main/install.sh | bash
or
wget -qO - https://gitlab.com/iocbio/gel/-/raw/main/install.sh | bash
and run by:
iocbio-gel/bin/iocbio-gel
Windows
Select the release under Releases and download Windows-executable packaged as a ZIP. Unpack the ZIP into any location of your PC and start the application by running gel.bat in the extracted folder. A video tutorial is available on YouTube and a written tutorial is available on homepage.
Running IOCBIO Gel
Starting the application
Linux/Mac users run the command in the folder that was used to install the software: iocbio-gel/bin/iocbio-gel
Windows users go to the extracted folder and run: gel.bat
On the initial launch, you will need to choose the image source and the database, as described below under Settings.
Settings
When starting the application for the first time, you will be asked to choose a few settings. A video tutorial is available on YouTube. Settings are accessible in the main application window via a button on the left.
First, select Image source and Database connection. Your selection depends on whether you want to store your files and results locally, i.e., on your own computer, or whether your files and results are stored in a shared space.
Image source
Images can be stored and accessed either as files or through specialized image server interfaces.
If you want to access your images as files, simply select Local files under Image source selection. This includes files stored on the local hard drives or files on some network-mounted filesystems, such as through network shares. It also allows you to use network storage and share the images in workgroups. The main requirement is that the images are visible through a local PC file path and are accessed in the same manner as local files.
Your workplace may also have a specialized image database for central storage of microscopy and other images from all lab members. For example, in our laboratory, all images are stored in our OMERO central repository (https://www.openmicroscopy.org/omero/). In contrast to the local files, these central storage servers use specialized interfaces that can be used by the program to retrieve images.
If your images and results are stored centrally in OMERO, select Image source OMERO.
Insert:
• The host name/address: should be available from your local systems administrator.
• The port: you probably have to use the default port (4064).
• Your username.
• Your password.
Database connection
Under Database connection, select SQLite if you want to store analysis data locally.
Alternatively, if you have a central database, select PostgreSQL from the drop-down lists.
Insert:
• The hostname/address: should be available from your local systems administrator.
• The port: we suggest using the default port (5432).
• The SSL mode: we suggest using the default (prefer encryption).
• The name of your database: should be available from your local systems administrator.
• The schema: we suggest using the default (gel).
• Your username.
• Your password.
View and edit modes
The software has two modes to avoid accidental changes, viewing and editing (see the upper right corner, Figure 1). By default, the software opens in viewing mode. In order to make any changes, click on the toolbar button in the upper-right corner. It will change to editing mode, and you can add types of measurements, analyze gels, and perform other similar operations. Select the action you want to perform by pressing the corresponding button in the panel on the left, such as Gels, Projects, or Types.
Figure 1. Edit and view mode shown while defining measurement types
Types
Under Types, you define the types of measurements. First, click on Types in the panel on the left. To add types, highlight the Name field and start typing. In the example shown above (Figure 1), we did a western blot to assess the expression of AMPK. Before antibody incubation, the membrane was stained with Ponceau to visualize and assess the overall protein content in each lane. Therefore, we put the types Ponceau and AMPK. With the types of measurements defined, continue with projects.
Projects
When doing multiple analyses, you might want to split your data according to which project they belong to. First, click on Projects in the panel on the left. Click Add a new project. By double-clicking on the Name and Comment fields, you can name the project and add a comment. Projects can be organized into a tree. For that, drag and drop the project under another one to form a branch. Notice that the subprojects are shown with the full path (the last column) consisting of the parent and the child names (Figure 2).
Figure 2. Projects can be defined as a hierarchy
Gels
First, click on Gels in the panel on the left. Define a new gel by clicking on + New in the table. That is possible only when the application is in the editing mode.
Name the gel: Highlight the Name field and start typing.
Set the reference time. In our case, we used the date of transfer as the reference time, but this is open to interpretation, and each user can decide which reference time to use. Highlight the Date and time field and double-click to see the drop-down calendar, where you can select the correct date. You can also specify the time of the day. If this field is left blank, then the software will automatically put the current time.
Insert a comment, if needed, by highlighting the Comment field and typing.
Here, you can also select which project this gel belongs to. Double-click on the Projects field and select the project.
The number of lanes can be set later and is shown based on the current records available for this gel.
You are now ready to start analyzing your images.
Image analysis
There is a basic and advanced usage video tutorial available on YouTube.
Clicking on Gels shows you a list of all the gels you have. To start the image analysis of a particular gel, press its ID in the list of gels or the name of the gel in the left column.
Define the lanes of the gel. Press + New until you have the number of lanes that you want to analyze.
In our lab, we often have two ladders—one in the left-most and one in the right-most lane. As we do not include the ladder in the image analysis, we only analyze, for example, 13 out of 15 lanes (Figure 3). It is up to you how to define lane indexing, whether to take into account ladders or not.
There are multiple columns:
• ID: the ID of the lane
• Lane: the number of the lane
• Sample ID
• Protein: the amount of sample protein that was loaded into each well on the gel
• Reference: tick if the samples are used as references for comparing between gels (could be one or multiple)
• Comments
Notice that each lane has two IDs. Lane ID, marked as ID in the software, is a unique ID for each lane among all gels in the local database. This is generated by the software and does not have to be changed by the user. Sample ID allows you to link the sample in this lane to your other records. It can be either alphanumeric or numeric, and advanced users can impose more restrictions on it by changing the database rules directly using database software. Through the sample ID, you can also link your gel measurements to the data from other experiments in the database if you have other software using the same database. In our case, we keep animal lineage, sample descriptions, and number of experimental records all stored in the same database and analyzed or entered through an open-source web interface or specialized software for analysis of time series or calcium sparks [5,6].
If you want to shift the number of lanes, it is easier to do this already after adding the first lane. The lane number will increase with each new addition.
Note that you can sort the lanes in ascending or descending order. By sorting in descending order, you can keep adding new lanes on the top of the table.
When the details of the different lanes are added, you can add the images from this gel. Click on Add new image. If you did multiple images of the same membrane, for example, Ponceau and antibody staining, both images are added here. You are now ready to analyze the signal intensity of the lanes in the pictures.
Figure 3. Overview of a single gel with its lanes and connected images
Image selection
With the gel of interest selected, double-click on the image you want to analyze under Images. This will open the picture in the Adjust tab. Before that, there is the Raw tab, where you see the raw image as well as the image without the applied colormap.
Defining the region of interest (ROI)
In the Adjust tab, click on Add ROI to select the ROI for your analysis. The ROI will be marked with a red square. The ROI can be moved by clicking within the ROI and dragging it to where you want it to be.
The ROI has two handles. In the upper-left corner, there is a circular handle for changing the rotation of the ROI. In the lower-right corner, there is a diamond-shaped handle for changing the size of the ROI. The rotation can also be set on the slider in the upper right corner. When you have defined your ROI, click Apply.
Background subtraction
After defining the ROI, you will automatically be taken to the Background tab, where you can subtract the background. First, you must define whether the background color is dark or light. Then, you select the kind of background subtraction that suits your image. Three kinds of background subtractions are available: Flat, Ball, and Ellipsoid.
Select the kind of background subtraction most suitable for your image. If the background has little variation, select the flat background. If the background is lighter in one part of the image than another, select Ball or Ellipsoid. For the latter two options, you need to define the radius of the ball or the radii of the ellipse. Select a relatively large radius (for example, 500 or more). If the radius is too small, part of the signal will also be subtracted. Click Apply to see the background that was subtracted as well as the final result. If you are unhappy with the result, try out another kind of background subtraction or change the diameter used for the background subtraction.
Marking and positioning the lanes for analysis
With the background subtracted, go to Lanes and click Add new lane. Position the lane on your image. Click add new lane and position the second lane as well. After that, every new lane will be positioned automatically based on the location of the previous lanes. Add new lanes until you have marked all the lanes that you want to analyze with an analysis lane. In each analysis lane, the center is marked with a green dotted line, and the sides are marked with green dashed lines. The intensity profile of each analysis lane will show up in the right-side panel (Figure 4).
You can zoom in and out by clicking on the gel picture and using the scrolling wheel of your mouse.
You will need to adjust the position and shape of the analysis lanes.
To move an analysis lane, position the mouse over the central dotted line—it will turn to a solid, red line. Click to drag and drop.
To change the width of the analysis lane, position the mouse over the little red square on the right side of the lane; this little square will turn yellow. Click and drag to adjust the width of the lane.
You can choose to adjust the width of just one analysis lane at a time or to keep the same width of all analysis lanes. For this, there is a toggle button in the upper-right corner, where you can choose Lane widths: Individual to change the width of one analysis lane, or Lane widths: Synced to keep the same width of all analysis lanes.
If the lane on your image is curved, you can incline or curve the analysis lane to follow the lane on the image.
To incline the analysis lane, the central line has a top and a bottom handle. They are marked as little red squares, and they can be moved left or right to incline the analysis lane according to the lane in your picture.
To curve the analysis lane, double-click on the central line to add extra handles. They will appear as little red squares along the central line, where you double-click. To grab a handle, position the mouse over it, and it will turn yellow. Click to grab and drag the handles on the analysis lane to follow the shape of the lane on the image. You can add multiple handles, but you must move the handle you made before you can add an additional handle (Figure 4).
All actions can be reversed by pressing Undo in the toolbar (shortcut Ctrl + z).
To remove a handle, place the mouse over the handle and check that the little red square changes color to yellow. Then, right-click on the handle and choose Remove handle from the drop-down menu. The analysis lane will straighten between the remaining handles.
When all the analysis lanes are adjusted according to the picture, you can examine the intensity profile for each lane.
Figure 4. Gel lanes on gel image
Setting the baseline of the intensity profile
The intensity profile of each lane is shown in the right-side panel (Figure 4). Here, it is possible to adjust the position of the baseline. We usually adjust the baseline to follow the dips of the intensity profile.
To interact with a plot, you have to activate it. To do that, click on it. The plot will be deactivated automatically as soon as the mouse pointer moves out of the plot area.
You can zoom in and out by clicking on the intensity plot and using the scrolling wheel of your mouse. To return to the original view, you can press the A button in the lower-left corner of the plot.
To add a handle, double-click on the baseline, and the handle will appear as a little black square.
To grab a handle, click on the window to ensure it is active. Place the mouse over the handle and check that the little black square becomes bold. Then, you can click and drag to position it.
To remove a handle, place the mouse over the handle and check that the little black square changes to bold. Then you can right-click on the handle and choose Remove handle from the drop-down menu.
When you are happy with the position of the baseline in all the intensity profiles, you can move on to the Measurements tab.
Obtaining signal intensity measurements
On the Measurements tab, you first add a new measurement. The measurement corresponds to the integrated signal intensity over the region of interest. Within the family of methods analyzed by IOCBIO Gel, it is assumed that the measurement of interest m is given by the formula:
where x is a position along the intensity profile of a lane of interest, I(x) corresponds to intensity at position x, and Ibg(x) is the baseline intensity. Note that since the pixel intensities are unitless and determined by specific camera hardware and other conditions, the measurement result m is also unitless. Therefore, m should be used in subsequent analysis only for relative comparisons with other measurement results, typically by dividing the compared measurements by a reference value.
To add a measurement, you can either click Add new measurement in the upper-right corner or click + New in the lower-right corner (Figure 5). Next, select the type of measurement that you are doing by double-clicking on the Type field and selecting it from the drop-down list. In the example shown below, we selected Ponceau.
Now, you simply click with the mouse on all the lanes that you want to use for your analysis. The dashed green lines marking the edges of each lane will be highlighted by a solid orange line below, and the intensity value will be shown in the Measurement Lanes field below the image. In this field, each measurement is shown with:
• Its own ID.
• The lane number.
• The value of the intensity.
• Success—tick if the measurement is successful.
• Comments—if you have any.
It is quite common that you do not want to measure the intensity of all the bands in each lane, but rather one or—in the case of protein staining—some of the bands, i.e., you want to specify the height of your analysis lanes. To select the band(s) whose intensity you want to measure, go to the intensity profile for each lane. There are sliders at each end of the profile, which are displayed as yellow dashed lines. Click on the intensity profile to activate the window and position the mouse over the slider; it will turn into a solid, yellow line. Click with the left mouse button and hold it down while adjusting the slider’s position. Notice that the value in Measurement Lanes changes as you adjust the sliders.
You can choose to adjust the height of just one analysis lane at a time or to keep the same height of all analysis lanes. For this, there is a toggle button in the upper right corner, where you can choose Measurement regions: Individual to change the height of one analysis lane, or Measurement regions: Synced to keep the same height of all analysis lanes.
In the example shown, we uploaded two images from the same membrane: In one image, the overall protein was stained using Ponceau, and in the other image, the protein of interest was labeled with antibodies.
After analyzing the overall protein stain intensity in each lane, go back to Gels (in the upper-left corner) and click on the ID of the gel you are analyzing.
Double-click on the next image that you want to analyze. In this example, it is the image with the protein of interest, t-AMPK.
Adjust the ROI of the image (see step B7b) and analyze this image as before, subtracting the background, marking the lanes for analysis, setting the baseline of the intensity profile, and obtaining signal intensity measurements.
Figure 5. Selecting measurement type for an image (bottom right) and adjusting the region where the integration is done (top-right graph)
Data analysis
Statistical analysis should be performed in another application. You can choose to directly fetch the data from the database or export it into a spreadsheet first.
Accessing the data from the database directly
Many of the statistical analysis programs or environments support connection to SQL databases. Those include R and Python. Depending on the environment, you have to define the connection to the database and fetch the data using an SQL script. Two example SQL scripts are available at main/sql. Example code for data fetching in R is available at iocbioR.
Exporting the results
As an alternative to fetching the data from the database directly, you can export it to a spreadsheet. To do so, click on Export in the lower-left corner. You will be prompted to name the file and select a location for your exported data. The Excel file has several sheets with the information from the gel analysis software:
• Gels
• Measurement types
• Gel lanes
• Images
• Measurement (raw)
• Reference measurement
• Measurements (normalized)
For most users, the last sheet with the normalized data will be the most interesting. It shows the measurement intensity values normalized to the reference sample that was defined when specifying the number of lanes to analyze (see Image analysis). If you have more than one reference lane, the values from each lane will be normalized to an average of the values from the reference lanes. These normalized values take into account the protein content that was specified for each of the lanes.
The normalization is only within one type and within one gel. Thus, the overall protein stain intensity (in this example by Ponceau, P) is normalized to the protein stain intensity of the reference sample (Pref), i.e., P/Pref. The signal intensity of the protein of interest (S) is normalized to the signal intensity of the reference sample (Sref), i.e., S/Sref.
To analyze the protein of interest, it is common practice to relate its intensity to the overall protein stain intensity of each lane. Thus, you compare S/P between lanes within one image.
However, it is common to have so many samples that you want to compare between images as well. To compare between images, you must calculate S/Sref/P/Pref.
If you have only a few images to compare, this can easily be done in Excel. However, for larger datasets, we recommend using a database so these ratios can be calculated in the SQL command to fetch the data.
Validation of protocol
This protocol or parts of it have been used and validated in the following research article:
Kütt et al. [3]. Simple analysis of gel images with IOCBIO Gel. BMC Biology (Figure 3, panels B–C).
General notes and troubleshooting
General notes
Limitations of the software.
Currently, setting the baseline of the intensity profile has to be done individually for each lane. We plan to add a syncing feature, as in Lane width or Measurement region selection. We are also working on improving the export feature to include the export of the database snapshot to share with other research teams.
Bugs and feature requests.
IOCBIO Gel is an open-source software and relies on feedback from users in its development. This includes new features and fixing bugs.
Bugs and new feature requests are considered as “Issues” and are reported on the project page. To see current issues and report new ones, follow the link Issues on the project page. Project is hosted at GitLab. If you want to open a new issue or comment on the reported one, you have to register as a user on that platform.
To open a new issue, press the button New issue on the GitLab issues page.
Next:
• Write the title of the issue.
• Keep Type as the default, “Issue”.
• Write a description of the issue.
• Assignee, milestone, and labels do not have to be selected.
• Press the blue button Create issue.
While filing the issue, feel free to paste screenshots that describe the problem. These include screenshots with the error messages and configuration settings. While care has been taken to avoid showing database login passwords in the configuration settings, please check all the screenshots for sensitive information before pasting them into GitLab.
Troubleshooting
Problem 1: Error while installing ZeroC ICE
Possible cause: OMERO Python library requires a specific version of ZeroC ICE. At the moment of writing, 3.6.5.
Solution: That version is not possible to install with Python 3.11 and newer due to a bug. Fixes have been available but for the newer ZeroC Ice version (https://github.com/zeroc-ice/ice/pull/1394). We have backported these bug fixes to 3.6.5 and made the patched version available in a separate repository. It can be installed using the following URL: https://gitlab.com/iocbio/libs/zeroc-ice-py/-/archive/v3.6.5/zeroc-ice-py-v3.6.5.tar.gz.
Problem 2: Background subtraction not working properly.
Possible cause: The original image-capturing software changed the background, for example from dark to light.
Solution: Try with a dark background if the subtraction does not work with a light background or the other way around.
Acknowledgments
This work was supported by the Estonian Research Council (PRG1127). Protocol was described and validated in Kütt et al. [3].
Competing interests
The authors declare that they have no competing interests.
References
Schneider, C. A., Rasband, W. S. and Eliceiri, K. W. (2012). NIH Image to ImageJ: 25 years of image analysis. Nat Methods. 9(7): 671–675.
Schooler, J. W. (2014). Metascience could rescue the ‘replication crisis’. Nature. 515(7525): 9.
Kütt, J., Margus, G., Kask, L., Rätsepso, T., Soodla, K., Bernasconi, R., Birkedal, R., Järv, P., Laasmaa, M., Vendelin, M., et al. (2023). Simple analysis of gel images with IOCBIO Gel. BMC Biol. 21(1): 225.
Wilkinson, M. D., Dumontier, M., Aalbersberg, I. J., Appleton, G., Axton, M., Baak, A., Blomberg, N., Boiten, J. W., da Silva Santos, L. B., Bourne, P. E., et al. (2016). The FAIR Guiding Principles for scientific data management and stewardship. Sci Data. 3(1): e18.
Laasmaa, M., Karro, N., Birkedal, R. and Vendelin, M. (2019). Iocbio Sparks Detection and Analysis Software. Biophys J. 116(3): 384a.
Vendelin, M., Laasmaa, M., Kalda, M., Branovets, J., Karro, N., Barsunova, K. and Birkedal, R. (2020). IOCBIO Kinetics: An open-source software solution for analysis of data traces. PLoS Comput Biol. 16(12): e1008475.
Article Information
Publication history
Received: May 10, 2024
Accepted: Jul 12, 2024
Available online: Aug 2, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Molecular Biology > DNA > Electrophoresis
Computational Biology and Bioinformatics
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In Vitro Hyphal Branching Assay Using Rhizophagus irregularis
TT Takaya Tominaga
HK Hironori Kaminaka
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5054 Views: 637
Reviewed by: Andrea GramaticaRaju Mondal Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant Physiology Dec 2023
Abstract
Most terrestrial plants are associated with symbiotic Glomeromycotina fungi, commonly known as arbuscular mycorrhizal (AM) fungi. AM fungi increase plant biomass in phosphate-depleted conditions by allocating mineral nutrients to the host; therefore, host roots actively exude various specialized metabolites and orchestrate symbiotic partners. The hyphal branching activity induced by strigolactones (SLs), a category of plant hormones, was previously discovered using an in vitro assay system. For this bioassay, AM fungi of the Gigaspora genus (Gigasporaeae) are commonly used due to their linear hyphal elongation and because the simple branching pattern is convenient for microscopic observation. However, many researchers have also used Glomeraceae fungi, such as Rhizophagus species, as the symbiotic partner of host plants, although they often exhibit a complex hyphal branching pattern. Here, we describe a method to produce and quantify the hyphal branches of the popular model AM fungus Rhizophagus irregularis. In this system, R. irregularis spores are sandwiched between gels, and chemicals of interest are diffused from the surface of the gel to the germinating spores. This method enables the positive effect of a synthetic SL on R. irregularis hyphal branching to be reproduced. This method could thus be useful to quantify the physiological effects of synthesized chemicals or plant-derived specialized metabolites on R. irregularis.
Key features
• Development of an in vitro hyphal branching assay using germinating spores of Rhizophagus irregularis.
• This in vitro assay system builds upon a method developed by Kameoka et al. [1] but modified to make it more applicable to hydrophilic compounds.
• Optimized for R. irregularis to count the hyphal branches.
• This bioassay requires at least 12 days to be done.
Keywords: AM symbiosis Hyphal branching assay Rhizophagus species branching factor Axenic culture
Graphical overview
Simplified overview of the hyphal branching assay using Rhizophagus irregularis spores
Background
In nature, plants are surrounded by diverse and numerous microbes in their leaves (phyllosphere) and roots (rhizosphere). Since these microbes beneficially or detrimentally modulate the host’s growth, host plants utilize their own specialized metabolites to manipulate the assembly of microbes. In particular, Glomeromycotina fungi, termed arbuscular mycorrhizal (AM) fungi, are representative beneficial microbes that have been found to be associated with more than 70% of terrestrial plant species [2]. AM fungi allocate inorganic phosphate from the soil to the host plant and, in return, the host plant shares its photosynthates with the symbiotic fungi. This mutual interaction begins with a category of root-secreted plant hormones, the strigolactones (SLs), which possess hyphal branching activity in AM fungi [3]. SLs were initially identified from cotton root exudates and described as germination stimulants of root parasitic plants [4]. Forty years later, their hyphal branching activity was reported using an AM fungus of the Gigaspora genus, Gigaspora margarita [3,5]. Prior to that discovery, Gigaspora fungi had been used in bioassays to identify signal compounds in host root exudates [6–9]. Thus, in vitro assay systems using AM fungi are essential to clarify the chemical communication between AM fungi and host plants.
G. margarita usually forms large spores (approximately 200 μm in diameter), as suggested by the genus name [10]. In addition to a uniquely large spore size, the fungus exhibits a simple hyphal branching pattern, forming thick and linear hyphae [3,5]. Therefore, G. margarita has often been used for in vitro bioassays. However, recent studies aimed at revealing the molecular mechanisms regulating AM symbiosis have mainly applied Rhizophagus irregularis because fungal genomic information, including of several intraspecies lines, is available [11–13]. Another reason is that Rhizophagus fungi have more beneficial traits (i.e., plant growth promotion) than Gigaspora fungi [14,15]. On the other hand, Rhizophagus fungi have small spores (approximately 50 μm in diameter), thin and winding hyphae, and a complex branching pattern [16–19]. Taken together, the hyphal branching assay using R. irregularis is technically difficult [20]. This is the reason why bioassay using R. irregularis is performed by measuring the total length of hyphae [20–22].
In hyphal branching assays using Gigaspora fungi, chemicals of interest have been loaded onto paper discs [3,5]. On the other hand, diffusion assays are limited to the local treatment of reagents and require specialized techniques. In our previous study, we modified an in vitro bioassay system that was originally applied for asymbiotic sporulation of R. irregularis [1]. Using our protocol, which is technically easy, chemicals can be evenly applied to all parts of R. irregularis. Compared to the protocols for Gigaspora fungi, this system is useful for assessing the hyphal branching activity induced by various chemicals in this model AM fungus.
Materials and reagents
Biological materials
Rhizophagus irregularis DAOM197198 (4,000 spores/mL) (PremierTech)
Reagents
Acetone (FUJIFILM WAKO Pure Chemical, CAS number: 67-64-1)
Ethanol (99.5%) (FUJIFILM WAKO Pure Chemical, CAS number: 64-17-5)
Gastrografin for oral/enema use (Bayer, 597.3 g/L amidotrizoic acid, CAS number: 117-96-4)
Magnesium sulfate heptahydrate (MgSO4·7H2O) (FUJIFILM WAKO Pure Chemical, CAS number: 10034-99-8)
Potassium nitrate (KNO3) (FUJIFILM WAKO Pure Chemical, CAS number: 7757-79-1)
Potassium chloride (KCl) (FUJIFILM WAKO Pure Chemical, CAS number: 7447-40-7)
Potassium dihydrogen phosphate (KH2PO4) (FUJIFILM WAKO Pure Chemical, CAS number: 7778-77-0)
Calcium nitrate tetrahydrate [Ca(NO3)2·4H2O] (FUJIFILM WAKO Pure Chemical, CAS number: 13477-34-4)
Sucrose (FUJIFILM, CAS number: 57-50-1)
Fe(III)-EDTA (NaFeEDTA·3H2O) (DOJINDO, CAS number: 15708-41-5)
Potassium iodide (KI) (FUJIFILM WAKO Pure Chemical, CAS number: 7681-11-0)
Manganese chloride tetrahydrate (MnCl2·4H2O) (FUJIFILM WAKO Pure Chemical, CAS number: 13446-34-9)
Zinc sulfate heptahydrate (ZnSO4·7H2O) (FUJIFILM WAKO Pure Chemical, CAS number: 7446-20-0)
Boric acid (H3BO3) (FUJIFILM WAKO Pure Chemical, CAS number: 10043-35-3)
Copper sulfate pentahydrate (CuSO4·5H2O) (FUJIFILM WAKO Pure Chemical, CAS number: 7758-98-7)
Disodium molybdate(VI) dihydrate (Na2MoO4·2H2O) (FUJIFILM WAKO Pure Chemical, CAS number: 10102-40-6)
Potassium hydroxide (KOH) (FUJIFILM WAKO Pure Chemical, CAS number: 1310-58-3)
Glycine (FUJIFILM WAKO Pure Chemical, CAS number: 56-40-6)
Thiamine hydrochloride (FUJIFILM WAKO Pure Chemical, CAS number: 67-03-8)
Pyridoxine hydrochloride (FUJIFILM WAKO Pure Chemical, CAS number: 58-56-0)
Nicotinic acid (FUJIFILM WAKO Pure Chemical, CAS number: 59-67-6)
Myo-inositol (FUJIFILM WAKO Pure Chemical, CAS number: 87-89-8)
Phytagel (Sigma-Aldrich, CAS number: 71010-52-1)
rac-GR24 (StrigoLab, CAS number: 76974-79-3)
Solutions
M medium (see Recipes) [16]
Calcium stock of M medium (see Recipes)
1,000× vitamin stock of M medium (see Recipes)
0.3% phytagel containing 3 mM MgSO4·7H2O (see Recipes)
Recipes
M medium
Reagent Final concentration Quantity or Volume
MgSO4·7H2O 2.97 mM 731 mg
KNO3 0.79 mM 80 mg
KCl 0.87 mM 65 mg
KH2PO4 0.035 mM 4.8 mg
Sucrose 10% (w/v) 10,000 mg
NaFeEDTA·3H2O 0.022 mM 9.18 mg
KI 0.0045 mM 0.75 mg
MnCl2·4H2O 0.030 mM 6 mg
ZnSO4·7H2O 0.0092 mM 2.65 mg
H3BO3 0.024 mM 1.5 mg
CuSO4·5H2O 0.00052 mM 0.13 mg
Na2MoO4·2H2O 9.92 nM 0.0024 mg
Phytagel 4% (w/v) 4,000 mg
H2O n/a 1,000 mL
Total (optional) n/a 1,000 mL
Note: Adjust pH at 5.5 using KOH solution and autoclave at 121 °C for 20 min. n/a, not applicable.
Calcium stock of M medium
Reagent Final concentration Quantity or Volume
Ca(NO3)2·4H2O 1.22 M 2,880 mg
H2O n/a 10 mL
Total (optional) n/a 10 mL
Note: Sterilize the stock using a 0.45 μm filter and store at room temperature (23–25 °C) until use. n/a, not applicable.
1,000× vitamin stock of M medium
Reagent Quantity or Volume
Glycine 30 mg
Thiamine hydrochloride 1 mg
Pyridoxine hydrochloride 1 mg
Nicotinic acid 5 mg
Myo-inositol 500 mg
H2O 10 mL
Total (optional) 10 mL
Note: Sterilize the mixture using a 0.45 μm filter and store at -30 °C until use.
0.3% phytagel containing 3 mM MgSO4·7H2O
Reagent Final concentration Quantity or Volume
MgSO4·7H2O 3 mM 739.4 mg
Phytagel 0.3% (w/v) 3,000 mg
H2O n/a 1,000 mL
Total (optional) n/a 1,000 mL
Note: No need to adjust pH. Autoclave at 121 °C for 20 min. n/a, not applicable.
Laboratory supplies
24-well plates (TrueLine, catalog number: TR5002)
1.5 mL microcentrifuge tube (BIO-BIK, catalog number: CF-0150)
15 mL centrifuge tube (Labcon, catalog number: 3132-345)
50 mL centrifuge tube (Labcon, catalog number: 3181-345)
10 mL disposable serological pipette (ASONE, catalog number: 2-4131-14)
Cell strainer (40 μm mesh) (Falcon, catalog number: 352340)
1 mL needleless plastic syringe (TERUMO, catalog number: SS-01T)
0.45 μm PTFE filter (Shimadzu, catalog number: GLCTD-HPTFE1345)
Inspection film tape (aglis, catalog number: LP0002K)
Equipment
Electronic pipette (Labnet, model: FASTPETTE Pro)
Laminar flow hood (SANYO, model: MCV-B131S)
Centrifuge (KUBOTA, model: 3740)
Swing rotor for 50 mL centrifuge tubes (KUBOTA, model: SD-242)
Autoclave (TOMY SEIKO, model: LSX-300)
Vacuum concentrator (Thermo Fisher Scientific, model: Savant SpeedVac DNA130)
Plant growth chamber (NIPPON MEDICAL & CHEMICAL INSTRUMENTS, model: LH-411S)
Stereomicroscope (Olympus, model: SZX16) equipped with a digital camera (Olympus, model: DP-26)
Imaging software (Olympus, model: CellSens standard v1.18)
Software and datasets
Microsoft Office Excel 10
R v4.2.0 (https://www.r-project.org/)
Procedure
Spore separation from inoculum
To observe and count hyphal branches, fragments of fungal hyphae should be removed from the R. irregularis spore suspension in advance. This step builds upon the protocol developed previously [23]. Maintain the suspension and collected spores axenic by working on a laminar flow food.
Prepare 8%, 16%, 32%, and 50% (v/v) Gastrografin solutions in sterile 50 mL centrifuge tubes. Invert the tubes to mix the dense and thick reagents with sterile distilled water.
In a new 50 mL centrifuge tube, pour each Gastrografin solution very gently using an electronic pipette equipped with a 10 mL disposable pipette as follows:
First, pour 10 mL of 50% Gastrografin, followed by 5 mL of 32%, 16%, and 8% Gastrografin. Do not disturb the layers.
Second, load 5 mL of spore suspension onto the surface of 8% Gastrografin softly. The spore concentration of the final solution can be changed.
Centrifuge the tubes at 500× g for 10 min at room temperature in a swing rotor for 50 mL centrifuge tubes.
Note: Decelerate the rotor slowly if possible.
Collect the supernatant, including the upper three layers, using the electronic pipette (Figure 1).
Note: R. irregularis spores must be floating in the 16% Gastrografin layer, and hyphal fragments should be precipitated to the bottom of the tube. Spore density differs among AM fungal species. In the case of R. clarus, spores float in the 32% Gastrografin layer.
Figure 1. Isolation of Rhizophagus irregularis spores from the inoculum
Pour the supernatant onto a sterile cell strainer and rinse the collected spores with 10 mL of sterile distilled water three times.
Put the cell strainer on a Petri dish filled with sterile distilled water and move the spores to a new 15 mL centrifuge tube.
Note: Ideally, adjust spore concentration to approximately 2,000 spores/mL.
Keep the spore suspension at 4 °C in the dark.
Preparation of germinating R. irregularis spores and chemical treatment
Add 1 mL of sterile calcium stock of M medium and 1 mL of 1,000× vitamin stock of M medium (see Recipes) to 1 L of autoclaved M medium.
Pour 350 μL of M medium into each well of a sterile 24-well plate and keep the plate still until the gel solidifies.
Dilute the purified spore suspension to approximately 1,000 spores/mL with sterile distilled water.
Drop an 8 μL aliquot of the purified spore suspension onto the center of each M medium.
After placing spores on the media, check if approximately 8 spores are placed on the medium center using a stereomicroscope. Adjust the number under a laminar flow condition if one well contains more or fewer spores. In case of more spores, remove extra spores by sucking them with a 200 μL micropipette.
Open the 24-well plate in the laminar flow hood for approximately 5 min to evaporate excessive water (Figure 2).
Note: The spores move around on the medium surface if too much water remains.
Pour 150 μL of 0.3% phytagel containing 3 mM MgSO4·7H2O (see Recipes). The gel should be cooled sufficiently in advance to allow it to be touched (approximately 40 °C). The 0.3% phytagel should be poured gently into the perimeter of the M medium using a micropipette (see Figure 2).
Figure 2. In vitro culture of Rhizophagus irregularis spores and chemical treatment. A. R. irregularis spores (approximately 8 spores) are placed on the center of each well filled with M medium. After drying off excess water, cool 0.3% phytagel with 3 mM MgSO4·7H2O is poured gently into the perimeter of the M medium to cover the spores. B. To exchange the solvent of rac-GR24 (positive control) from acetone to distilled water, the acetone stock is centrifuged in vacuo. The residues are redissolved in distilled water and sterilized using the 0.45 μm PTFE filter. Images show germinating R. irregularis spores (arrowheads) treated with H2O (C) and 100 nM GR24 (D) for 5 days. E. The number of hyphal branches in R. irregularis treated with H2O and 100 nM GR24. Dots and error bars represent individual values and the standard deviation, respectively (n > 15). The E-value was lower than 0.05 in the Wilcoxon rank-sum test.
Close the 24-well plate lid and seal it with two layers of inspection film to prevent the dehydration of the gels.
Place the plate horizontally in a growth chamber at 25 °C and incubate it for 5 days in the dark.
Note: In our condition, R. irregularis spores usually germinate in this period.
After 5 days, treat 200 μL of chemicals diluted in sterile distilled water onto each gel containing germinating spores. Use the positive control rac-GR24 diluted in acetone as follows:
Add an 8 μL aliquot of 10 mM rac-GR24 in a 1.5 mL tube.
Centrifuge the tube in a vacuum concentrator at 35 °C for 5 min. If acetone is left in the tube, adjust the evaporation time to remove the solvent completely.
After evaporating the solvent, add 800 μL of distilled water to the tube.
In the laminar flow hood, sterilize the 100 nM rac-GR24 using a 1 mL needleless syringe with a 0.45 μm PTFE filter.
Pour 200 μL of the 100 nM rac-GR24 solution onto the gels.
Seal plates twice with inspection film tape and place them in the same growth chamber at 25 °C. Incubate plates for 7–10 days in the dark.
Quantification of R. irregularis hyphal branching
Under a stereomicroscope, count the number of hyphal branches except for that of the initially elongating hypha (Figure 3). Remove the 24-well plate lid if it becomes misty during observation.
Notes:
Count hyphal branches from the runner hyphae (see Figure 3A, B).
Ignore the short hyphae formed at the boundary of the subtending and runner hyphae. The hyphae often show vigorous branching without any chemical treatment (Figure 3C, D).
Basically, the hyphal branches of a single spore are considered to be one biological replicate. If multiple spores are tied with a subtending hypha, it is counted as one germinating spore (see Figure 3B).
Check the differences among treatments using a statistical calculation (see Data analysis).
Figure 3. Hyphal structures and growth patterns of Rhizophagus irregularis. A, B. Count the number of hyphal tips emerging from runner hyphae to quantify the hyphal branching. When you find multiple spores tied with a subtending hypha (B, right drawing), count them as one biological replicate. C, D. Germinating spores with highly branching hyphal clusters near the subtending hyphae. Exclude these spores from your quantification. Arrows indicate germinating R. irregularis spores drawn in (B) and (D).
Data analysis
Data can be analyzed using Microsoft Excel. We usually prepare three or four wells containing germinating spores for each treatment. For statistical analysis, R is used to perform Wilcoxon’s rank-sum test with a Bonferroni correction in the case of multiple comparisons. Approximately 20 spores per treatment at least should be assessed. The p-value cutoff is 0.05.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Tominaga et al. [24]. Monoterpene glucosides in Eustoma grandiflorum roots promote hyphal branching in arbuscular mycorrhizal fungi. Plant Physiology (Figure 3).
General notes and troubleshooting
General notes
Hyphal fragments should be removed as much as possible because some hyphal branches will emerge from them and this will mask the germinating spores.
Many spores, i.e., exceeding 8, and a longer (more than 10 days) incubation period can make hyphal branches indistinguishable. In addition, R. irregularis likely develops more hyphal branches when the spores are crowded in a medium, although the reasons remain unclear.
Exposing spores and hyphae to air influences R. irregularis growth and branching. The AM fungal spores exhibit straight hyphal elongation and moderate hyphal branching, although we are unsure why.
Troubleshooting
Problem 1: Spores do not germinate well.
Possible cause: The spore suspension includes many dead spores. Alternatively, phytagel is insufficiently cooled.
Solution: Increase the number of spores or prepare a new spore suspension. Make sure to pour sufficiently cooled phytagel onto the spores.
Problem 2: Cannot count the hyphal branches due to a complex branching pattern.
Possible cause: Each well may contain too many germinating spores, or the incubation period might be unnecessarily long.
Solution: Use fewer spores and shorten the period of chemical treatment. Instead, prepare more wells to assess sufficient spores (>10). Importantly, a germinating spore showing massive branches around subtending hypha without a long runner hypha can be ignored from the quantification. We observe germinating spores with traceable hyphal elongation and branching.
Acknowledgments
This protocol was developed by modifying a previous method [1]. The authors thank Dr. Hiromu Kameoka (China Academy of Science, China) for useful discussions on the in vitro culture of R. irregularis. This work was supported by the Japan Society for the Promotion of Science (JSPS) (KAKENHI grant numbers 20J21994 and 23KJ1578). The graphical abstract and figures were created using Biorender.com.
Competing interests
The authors declare that they have no competing financial or non-financial interests.
References
Kameoka, H., Tsutsui, I., Saito, K., Kikuchi, Y., Handa, Y., Ezawa, T., Hayashi, H., Kawaguchi, M. and Akiyama, K. (2019). Stimulation of asymbiotic sporulation in arbuscular mycorrhizal fungi by fatty acids. Nat Microbiol. 4(10): 1654–1660.
Brundrett, M. C. and Tedersoo, L. (2018). Evolutionary history of mycorrhizal symbioses and global host plant diversity. New Phytol. 220(4): 1108–1115.
Akiyama, K., Matsuzaki, K. and Hayashi, H. (2005). Plant sesquiterpenes induce hyphal branching in arbuscular mycorrhizal fungi. Nature. 435(7043): 824–827.
Cook, C. E., Whichard, L. P., Turner, B., Wall, M. E. and Egley, G. H. (1966). Germination of Witchweed (Striga lutea Lour.): Isolation and Properties of a Potent Stimulant. Science. 154(3753): 1189–1190.
Akiyama, K., Ogasawara, S., Ito, S. and Hayashi, H. (2010). Structural Requirements of Strigolactones for Hyphal Branching in AM Fungi. Plant Cell Physiol. 51(7): 1104–1117.
Besserer, A., Puech-Pagès, V., Kiefer, P., Gomez-Roldan, V., Jauneau, A., Roy, S., Portais, J. C., Roux, C., Bécard, G., Séjalon-Delmas, N., et al. (2006). Strigolactones Stimulate Arbuscular Mycorrhizal Fungi by Activating Mitochondria. PLoS Biol. 4(7): e226.
Nagahashi, G. and Douds, D. (2007). Separated components of root exudate and cytosol stimulate different morphologically identifiable types of branching responses by arbuscular mycorrhizal fungi. Mycol Res. 111(4): 487–492.
Besserer, A., Bécard, G., Jauneau, A., Roux, C. and Séjalon-Delmas, N. (2008). GR24, a Synthetic Analog of Strigolactones, Stimulates the Mitosis and Growth of the Arbuscular Mycorrhizal Fungus Gigaspora rosea by Boosting Its Energy Metabolism. Plant Physiol. 148(1): 402–413.
Ramos, A. C., Façanha, A. R. and Feijó, J. A. (2008). Proton (H+) flux signature for the presymbiotic development of the arbuscular mycorrhizal fungi. New Phytol. 178(1): 177–188.
Bécard, G. and Piché, Y. (1989). New aspects on the acquisition of biotrophic status by a vesicular—arbuscular mycorrhizal fungus, Gigaspora margarita. New Phytol. 112(1): 77–83.
Tisserant, E., Malbreil, M., Kuo, A., Kohler, A., Symeonidi, A., Balestrini, R., Charron, P., Duensing, N., Frei dit Frey, N., Gianinazzi-Pearson, V., et al. (2013). Genome of an arbuscular mycorrhizal fungus provides insight into the oldest plant symbiosis. Proc Natl Acad Sci USA. 110(50): 20117–20122.
Maeda, T., Kobayashi, Y., Kameoka, H., Okuma, N., Takeda, N., Yamaguchi, K., Bino, T., Shigenobu, S. and Kawaguchi, M. (2018). Evidence of non-tandemly repeated rDNAs and their intragenomic heterogeneity in Rhizophagus irregularis. Commun Biol. 1(1): 87.
Yildirir, G., Sperschneider, J., Malar C, M., Chen, E. C. H., Iwasaki, W., Cornell, C. and Corradi, N. (2021). Long reads and Hi‐C sequencing illuminate the two‐compartment genome of the model arbuscular mycorrhizal symbiont Rhizophagus irregularis. New Phytol. 233(3): 1097–1107.
Lendenmann, M., Thonar, C., Barnard, R. L., Salmon, Y., Werner, R. A., Frossard, E. and Jansa, J. (2011). Symbiont identity matters: carbon and phosphorus fluxes between Medicago truncatula and different arbuscular mycorrhizal fungi. Mycorrhiza. 21(8): 689–702.
Kaur, S., Campbell, B. J. and Suseela, V. (2022). Root metabolome of plant–arbuscular mycorrhizal symbiosis mirrors the mutualistic or parasitic mycorrhizal phenotype. New Phytol. 234(2): 672–687.
Hildebrandt, U., Janetta, K. and Bothe, H. (2002). Towards Growth of Arbuscular Mycorrhizal Fungi Independent of a Plant Host. Appl Environ Microbiol. 68(4): 1919–1924.
Hildebrandt, U., Ouziad, F., Marner, F. J. and Bothe, H. (2006). The bacterium Paenibacillus validus stimulates growth of the arbuscular mycorrhizal fungus Glomus intraradices up to the formation of fertile spores. FEMS Microbiol Lett. 254(2): 258–267.
Abdellatif, L., Lokuruge, P. and Hamel, C. (2019). Axenic growth of the arbuscular mycorrhizal fungus Rhizophagus irregularis and growth stimulation by coculture with plant growth-promoting rhizobacteria. Mycorrhiza. 29(6): 591–598.
Yamato, M., Yamada, H., Maeda, T., Yamamoto, K., Kusakabe, R. and Orihara, T. (2022). Clonal spore populations in sporocarps of arbuscular mycorrhizal fungi. Mycorrhiza. 32: 373–385.
Tsuzuki, S., Handa, Y., Takeda, N. and Kawaguchi, M. (2016). Strigolactone-Induced Putative Secreted Protein 1 Is Required for the Establishment of Symbiosis by the Arbuscular Mycorrhizal Fungus Rhizophagus irregularis. Mol Plant Microbe Interact. 29(4): 277–286.
Cosme, M., Fernández, I., Declerck, S., van der Heijden, M. G. A. and Pieterse, C. M. J. (2021). A coumarin exudation pathway mitigates arbuscular mycorrhizal incompatibility in Arabidopsis thaliana. Plant Mol Biol. 106: 319–334.
Serghi, E. U., Kokkoris, V., Cornell, C., Dettman, J., Stefani, F. and Corradi, N. (2021). Homo- and Dikaryons of the Arbuscular Mycorrhizal Fungus Rhizophagus irregularis Differ in Life History Strategy. Front Plant Sci. 12: e715377.
Furlan, V., Bartschi, H. and Fortin, J. A. (1980). Media for density gradient extraction of endomycorrhizal spores. Trans Br Mycol Soc. 75(2): 336–338.
Tominaga, T., Ueno, K., Saito, H., Egusa, M., Yamaguchi, K., Shigenobu, S. and Kaminaka, H. (2023). Monoterpene glucosides in Eustoma grandiflorum roots promote hyphal branching in arbuscular mycorrhizal fungi. Plant Physiol. 193(4): 2677–2690.
Article Information
Publication history
Received: May 20, 2024
Accepted: Jul 14, 2024
Available online: Jul 31, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Plant Science > Plant physiology > Endosymbiosis
Biological Sciences > Biological techniques > Microbiology techniques
Microbiology > Microbe-host interactions > Fungus
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Chloroform/Methanol Protein Extraction and In-solution Trypsin Digestion Protocol for Bottom-up Proteomics Analysis
TP Tess Puopolo
NS Navindra P. Seeram
CL Chang Liu
Published: Vol 14, Iss 16, Aug 20, 2024
DOI: 10.21769/BioProtoc.5055 Views: 718
Reviewed by: Dennis ProvinceNeha Nandwani Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Food & Function Jan 2024
Abstract
Bottom-up proteomics utilizes sample preparation techniques to enzymatically digest proteins, thereby generating identifiable and quantifiable peptides. Proteomics integrates with other omics methodologies, such as genomics and transcriptomics, to elucidate biomarkers associated with diseases and responses to drug or biologics treatment. The methodologies employed for preparing proteomic samples for mass spectrometry analysis exhibit variability across several factors, including the composition of lysis buffer detergents, homogenization techniques, protein extraction and precipitation methodologies, alkylation strategies, and the selection of digestion enzymes. The general workflow for bottom-up proteomics consists of sample preparation, mass spectrometric data acquisition (LC-MS/MS analysis), and subsequent downstream data analysis including protein quantification and differential expression analysis. Sample preparation poses a persistent challenge due to issues such as low reproducibility and inherent procedure complexities. Herein, we have developed a validated chloroform/methanol sample preparation protocol to obtain reproducible peptide mixtures from both rodent tissue and human cell line samples for bottom-up proteomics analysis. The protocol we established may facilitate the standardization of bottom-up proteomics workflows, thereby enhancing the acquisition of reliable biologically and/or clinically relevant proteomic data.
Key features
• Tissue/cell pellet sample preparation for bottom-up proteomics
• Chloroform/methanol protein extraction from murine tissue samples
• In-solution trypsin digestion proteomics workflow
Keywords: Proteomics Sample preparation In-solution digestion Protein extraction Chloroform/methanol
Graphical overview
Murine tissue sample chloroform/methanol protein extraction workflow for bottom-up proteomics analyses
Background
Sample preparation is a critical component of bottom-up proteomics analyses, yet it is often hindered by time constraints, performer errors, and intensive processes. Further, there remains a scientific need to develop reproducible, precise, and successful sample preparation protocols to advance proteomic discovery [1]. There are numerous sample preparation methods due to differences in sample type, protein heterogeneity, and physicochemical properties, which creates difficulty in standardizing workflows [2,3]. In-solution digestion (ISD) primarily employs varied buffers such as denaturants and detergents to denature protein samples followed by protein precipitation with organic solvents, such as chloroform/methanol, alcohol, or acetone [2]. A recent novel ISD protocol, sample preparation by easy extraction and digestion (SPEED), denatures proteins without the use of denaturants and detergents but rather through the use of trifluoracetic acid [2]. In-gel digestion (IGD), in contrast to ISD, involves the separation of proteins by size via gel electrophoresis, which has the capacity to reduce contaminants [4]. IGD is often coupled with SDS-PAGE, which utilizes the strong detergent SDS [5,6]. SDS may decrease the efficacy of proteases, such as trypsin, as well as reduce the signal/noise ratio, thereby limiting proteome coverage [5]. Therefore, it is advantageous to avoid the use of SDS in sample digestion steps or buffer solutions to prevent adverse effects and SDS removal steps. Similarly, another sample preparation method popularly employed includes device-based approaches such as filter-aided sample preparation (FASP) where SDS is also added to protein samples [7]. Samples are then filtered, and the protein is digested to elute peptides. However, FASP sample preparation protocols are time-consuming and have resulted in considerable variability, leading to the development of other methods [7]. Recently, the suspension trapping (S-Trap) method has garnered attention due to its protein extraction method via denatured size, which has improved time constraints and complexities in sample preparation [8]. However, the S-Trap method typically uses 5% SDS as a lysis buffer, and the inherent disadvantages of SDS still remain [7].
Therefore, ISD remains a promising sample preparation method compared to other approaches, yet standardization of methods for animal and cell culture samples is warranted. Herein, we developed an ISD protocol consisting of denaturation, alkylation, chloroform/methanol protein extraction, and digestion steps, as ISD methods are commonly employed in sample preparation of animal tissue samples [9]. Specifically, ISD complemented by a chloroform/methanol protein extraction method may be a reliable technique for animal and human tissue samples bottom-up proteomics as it is a fairly straightforward and fast method, which prevents protein degradation [10]. Chloroform/methanol protein extraction is a quantitative method first developed in 1984, utilizing a phase separation to precipitate proteins in solution; it has since been shown to be advantageous over acetone and ethanol for total peptides and peptides without missed cleavages [2,11]. Further, chloroform/methanol protein extraction produces a protein pellet that does not contain contaminants without reducing protein quantities [11]. In comparison to IGD, ISD is less error-prone and costly and has a higher likelihood of high peptide yield [2,4]. Further, studies have found that ISD is advantageous to identify peptides from the whole proteome, while other methods such as FASP may be better suited for specific protein recovery such as membrane proteins [12]. Our bottom-up proteomics tissue sample preparation technique using classic ISD and chloroform/methanol protein extraction methods provides a valid and reproducible workflow with protein yields suitable for in-depth, whole-proteome analyses.
Materials and reagents
Biological materials
Frozen mouse tissue sample
Note: This protocol has also been successfully conducted using cell lysis samples in the following manuscripts:
Li et al. (2024). Anti-Ferroptotic Effect of Cannabidiol in Human Skin Keratinocytes Characterized by Data-Independent Acquisition-Based Proteomics. J Nat Prod. DOI: 10.1021/acs.jnatprod.3c00759. (Figures 3 and 4) [13].
Li et al. (2023). Cannflavins A and B with Anti-Ferroptosis, Anti-Glycation, and Antioxidant Activities Protect Human Keratinocytes in a Cell Death Model with Erastin and Reactive Carbonyl Species. Nutrients. DOI: 10.3390/nu15214565. (Figure 2) [14].
Reagents
Dry ice (Airgas)
Urea (Sigma-Aldrich, catalog number: 51457-500 mL)
Triethylammonium bicarbonate buffer (TEAB) (Sigma-Aldrich, catalog number: T7408-100 mL)
PierceTM BCA Protein Assay kit with dilution-freeTM BSA protein standards (Thermo Fisher Scientific, catalog number: A55865)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A9418-5G)
DL-Dithioreitol (DTT) (Sigma-Aldrich, catalog number: D9779-1G)
Iodoacetamide (IAA) (Sigma-Aldrich, catalog number: I1149-5G)
Ammonium bicarbonate (Sigma-Aldrich, catalog number: A6141-25G)
Sodium deoxycholate (DOC) (Sigma-Aldrich, catalog number: 30970-25G)
Trypsin w/ CaCl2 (TPCK-treated), 10 pack (SCIEX, catalog number: 4445250)
Methanol CHROMASOLV LC-MS (Honeywell, catalog number: 34966-4L)
Chloroform-isoamyl alcohol mixture (Sigma-Aldrich, catalog number: 25666-100ML)
Milli-Q water
Formic acid (Sigma-Aldrich, catalog number: 695076-100ML)
Acetonitrile (ACN) CHROMASOLV LC-MS (Honeywell, catalog number: 34967-4X4L)
Solutions
TEAB (50 mM) and urea (8M) homogenization buffer (see Recipes)
50 mM ammonium bicarbonate (pH ~8) containing 3% w/v sodium deoxycholate (DOC) (see Recipes)
5% formic acid in acetonitrile and water (50:50 ratio) (see Recipes)
Recipes
TEAB (50 mM) and urea (8M) homogenization buffer
Reagent Final concentration
TEAB 50 mM
Urea 8 M
50 mM ammonium bicarbonate (pH ~8) containing 3% w/v sodium deoxycholate (DOC)
Reagent Final concentration
Ammonium bicarbonate 50 mM
DOC 3%
5% formic acid in acetonitrile and water (50:50 ratio)
Reagent Final concentration
Acetonitrile 47.5%
Water 47.5%
Formic acid 5%
Laboratory supplies
Pipette Tips (TipOne 10, 200, 1,000 μL) (USA Scientific, catalog numbers: 1111-3800, 1110-1800, 1111-2821)
Pipettes (ErgoOne Single-Channel Pipette 2.5, 10, 200, 1,000 μL) (USA Scientific, catalog numbers: 7100-0125, 7100-0510, 7100-2200, 7110-1000)
Soft tissue homogenizing mix 1.4 mm ceramic (2 mL tubes) nuclease free (Omni International, catalog number: 19-627)
1 mL Dounce tissue grinder (Avantor, catalog number: 357538)
Pour boat weigh dish 2-1/4"l × 1-3/4"w × 5/16"d, 20 mL cap (Wilkem Scientific, catalog number: 10177901)
96-well tissue culture plates (Cell Treat, catalog number: 229197)
96 150 μL PCT microtubes (Pressure Biosciences, Inc., catalog number: MTWS-MT-RK)
150 μL PCT microcaps (Pressure Biosciences, Inc., catalog number: MTWS-MC150-RK)
Pipet tip gel loading .57 mm O.D. 200 μL round non-sterile (Wilkem Scientific, catalog number: LABB13790)
Microcentrifuge tube 0.5 mL non-sterile (Cell Treat, Wilkem Scientific, catalog number: 72316004)
Microcentrifuge tube 1.5 mL non-sterile (Cell Treat, Wilkem Scientific, catalog number: 229441)
PK100 amber glass certified S/T vial (Sigma-Aldrich, catalog number: 29386-U)
Advantage 150 μL volume, 5 × 30 mm bottom spring glass inserts (Analytical Sales and Services Inc., catalog number: 20501)
Equipment
Bead Ruptor elite bead mill homogenizer (Omni International, model: 19-042E)
Microplate reader SpectraMax M2 (Molecular Devices, model: MDM2)
Pressure cycling technologies Barocycler (Pressure Biosciences Inc., model: NEP2320)
Circulating heater water bath (Thermo Fisher Scientific, model: CH 100)
Microcap tool Series 3 (Pressure Biosciences Inc., model: MTWS-CR-03)
Microtube adapter kit (Pressure Biosciences Inc., catalog number: MTTB-KEXT)
Centrifuge 5810 R (Eppendorf, model: 5811F)
Reciprocal shaking water bath (Precision Scientific, model: 66800)
Fisher vortex Genie 2TM (Fisher Scientific, catalog number: 12-812)
Software and datasets
Prism v10.0.3 (GraphPad, 07/05/2024)
Procedure
Homogenate preparation
Prepare homogenization buffer (see Recipes).
Add TEAB (50 mM) to urea (8 M).
Place the microcentrifuge tube with tissue sample on dry ice.
Place the Dounce tissue grinder glass homogenizer on dry ice.
Weigh 50 mg of mouse organ tissue with a weigh boat and place into the Dounce tissue grinder.
Add ~1 mL of homogenization buffer (50 mg/mL concentration) to the Dounce tissue grinder and grind the tissue sample until homogenized on dry ice.
Note: The protein concentration and total volume may be optimized to suit the user’s specific experimental needs. For example, if using nanoflow LC systems, a lower protein concentration may be suitable. Further, large proteomic studies where many samples need to be processed for proteomics analysis may utilize smaller lysate volumes to accelerate overall procedure time.
Transfer tissue lysis into an Omni bead homogenizer tube.
Homogenize tissue in an Omni bead homogenizer at 5 m/s for 30s.
Centrifuge samples at 14,000× g for 10 min at 4 °C.
Transfer the supernatant into new microcentrifuge tubes and place them on ice.
Protein quantification assay
Conduct BCA assay for protein quantification.
Use a standard curve (125–2,000 μg/mL) and multiple sample dilutions (e.g., 10-, 20-, 40-fold).
In a 96-well plate, add 25 μL of either standard or sample and then 200 μL of working reagent (prepared in a 50:1 ratio of reagent A to reagent B).
Cover the plate with aluminum foil and incubate at 37 °C for 30 min.
Read absorbance with the microplate reader at 562 nm.
In GraphPad Prism, interpolate sample protein concentrations based on a linear regression of the standard curve.
Prepare samples at 2,500 μg/mL (100 μL) and place them on ice for immediate use.
Note: Samples can be frozen at -20 °C for next-day use or at -80 °C long term and then thawed at 4 °C prior to use.
Adding BSA (optional) and DTT
Set the shaking water bath to 34 °C and centrifuge to 10 °C.
Prepare BSA at 0.2 mg/mL in milliQ water and spike samples with 10 μL.
Prepare DTT at 100 mM in milliQ water, add 25 µL to each tube, and vortex.
Caution: Wear protective equipment when handling DTT as it is harmful if swallowed, inhaled, or with skin contact.
Denature the proteins at 34 °C in the shaking water bath for 30 min (100 rpm).
Note: Ensure that the temperature does not exceed 35 °C. Once urea goes over this temperature, it will break down histidine bonds and form a gelatin-like substance.
Note: While a short denaturation time and the use of ammonium buffer in this protocol may decrease the chances of carbamylation developing during protein reduction with heat, users may optimize the reduction step by reducing for a longer time period at room temperature, if desired.
Chloroform/methanol precipitation
Place water, methanol, and chloroform on ice.
Prepare iodoacetamide (IAA) (200 mM) in milliQ water and add 25 μL to each sample.
Caution: Wear protective equipment and work under a vented fume hood when using IAA, as it is an acute toxin and harmful if swallowed, inhaled, or with skin contact.
Note: IAA is light-sensitive.
Place samples at room temperature (~20 °C) in the dark for 30 min for alkylation.
Concentrate and precipitate samples with the sequential addition (1:2:1 ratio) of ice-cold water (160 µL), methanol (320 μL), and chloroform (160 μL) and vortex after the addition of all solvents.
Caution: Wear protective equipment and work under a vented fume hood when using chloroform and/or methanol, as it is harmful if swallowed, inhaled, or with skin contact.
Centrifuge samples at 10,000× g for 5 min at 10 °C .
Remove the chloroform layer from samples (bottom layer) using a gel tip (Figure 1).
Note: Use precise care not to disrupt the pellet.
Critical: Removing the chloroform layer first will prevent the loss of protein sample as the protein pellet will transfer to the side of the tube.
Figure 1. Chloroform-methanol precipitation involves a phase separation identified by an immiscible methanol/water layer on top, a protein layer in the middle, and a chloroform layer on the bottom
Remove the methanol/water layer from samples (top layer) using a gel tip.
Note: Use precise care not to disrupt the pellet.
Add 200 μL of ice-cold methanol to the samples to wash them, and then remove it using a gel tip.
Note: Use precise care not to disrupt the pellet. The methanol wash will remove any additional organic solvents present.
Resuspend pellet in 130 μL of 50 mM ammonium bicarbonate (pH ~8) containing 3% w/v DOC (see Recipes). Scrape walls and pipette up and down.
Note: Samples can be stored at -80 °C if necessary, and then thawed at 4 °C prior to use.
Pressure cycling technology (PCT)-aided trypsin digestion
Turn on the Barocycler and the air compressor and set the Barocycler water bath to 35–37 °C.
Add 500 μL of milliQ water to the lyophilized TPCK-treated trypsin for a 1 mg/mL concentration and vortex.
Note: Resuspended trypsin can be stored at -20 °C for future use.
Add 10 μL of trypsin (10 μg) to each sample (250 μg of protein) (at a w/w ratio of 1:25 trypsin:protein) and vortex at a medium speed (setting 4 on a scale of 0–8) for 3~5 s.
Transfer 138 μL of each sample to individually labeled microtubes using gel tips.
Note: Use care when transferring the sample into the microtubes and aim to prevent bubbles.
Cap microtubes with the microcap tool.
Place microtubes into the Barocycler cartridge (Figure 2).
Place the Barocycler cartridge into the PCT Barocycler instrument.
Press PRECHARGE three times to prime the lines.
Run the Barocycler for protein digestion for 75 cycles, 60 s per pressure cycle (50 s high pressure, 10 s ambient pressure), with a pressure around 25.5 psi (Figure 3).
After completion of the Barocycler run, remove the Barocycler cartridge with the magnetic rod.
Remove caps from microtubes with the microcap tool.
Add 10 μL of trypsin to each sample (ratio of 1:25 w/w trypsin:protein) using a gel tip.
Repeat PCT digestion steps as described in steps E5–E8.
After completion of the Barocycler run, remove the Barocycler cartridge with the magnetic rod.
Remove microtubes from the cartridge, remove caps with the microcap tool, and place the PCT tubes in the microtube rack.
Note: PCT enhances trypsin digestion by promoting efficient and rapid protein breakdown through mechanical disruption, which accelerates the enzymatic process. Moreover, its controlled pressure cycles minimize sample variability, ensuring more reliable digestion results. In the absence of PCT, users can conduct the traditional trypsin digestion protocol, as recommended by the manufacturers, typically involving incubation with trypsin for 4–16 h at 37 °C.
Figure 2. Microtube cartridges where up to 16 samples can be run on the pressure cycling technology (PCT) Barocycler for trypsin digestion
Figure 3. Pressure cycling technology (PCT) Barocycler instrumentation utilized for trypsin digestion of samples. The cycling methods can be programmed prior to use.
Desalting and precipitation of DOC
Prepare 5% formic acid in a 50:50 ratio of ACN to water and add 15 μL to 0.5 mL microcentrifuge tubes (see Recipes).
Caution: Wear protective equipment and work under a vented fume hood, as formic acid and/or acetonitrile are harmful via all routes of exposure and flammable.
Add 135 μL of the respective samples using a gel tip to 0.5 mL microcentrifuge tubes containing 15 μL of 5% formic acid in a 50:50 ratio of ACN to water and vortex to desalt and precipitate the DOC.
Note: A white precipitate will immediately form.
Centrifuge samples at 10,000× g for 5 min at 10 °C to remove precipitated DOC.
Transfer sample supernatants into new 0.5 mL microcentrifuge tubes and centrifuge samples at 10,000× g for 5 min at 10 °C .
Transfer 100 μL of supernatant into an amber glass vial containing a bottom spring glass insert.
Samples are ready to be run for LC-MS/MS proteomics analyses.
Note: Samples can also be stored in microcentrifuge tubes, frozen at -80 °C, thawed at 4 , and pipetted into HLPC inserts/glass vials on the day of the LC-MS/MS run.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Puopolo et al. (2024). Exploring immunoregulatory properties of a phenolic-enriched maple syrup extract through integrated proteomics and in vitro assays. Food Funct. (Figures 2 and 3) [15].
Puopolo et al. (2023). Uncovering the anti-inflammatory mechanisms of phenolic-enriched maple syrup extract in lipopolysaccharide-induced peritonitis in mice: insights from data-independent acquisition proteomics analysis. Food Funct. (Figures 4 and 6) [16].
General notes and troubleshooting
General notes
Avoid multiple freeze/thaw cycles to ensure protein is not degraded.
This protocol is also applicable to cell lysate samples and has been validated in the following manuscripts:
Li et al. (2024). Anti-Ferroptotic Effect of Cannabidiol in Human Skin Keratinocytes Characterized by Data-Independent Acquisition-Based Proteomics. J Nat Prod. (Figures 3 and 4).
Li et al. (2023). Cannflavins A and B with Anti-Ferroptosis, Anti-Glycation, and Antioxidant Activities Protect Human Keratinocytes in a Cell Death Model with Erastin and Reactive Carbonyl Species. Nutrients. (Figure 2).
This protocol is also applicable to other animal tissue samples (such as rat brain) with minor sample preparation modifications and has been validated in the following manuscripts:
Schrader et al. (2024). Longitudinal markers of cerebral amyloid angiopathy and related inflammation in rTg-DI rats. Sci Rep. DOI: 10.1038/s41598-024-59013-7. (Figures 3–6) [17].
Schrader et al. (2021). Distinct brain regional proteome changes in the rTg‐DI rat model of cerebral amyloid angiopathy. J Neurochem. DOI: 10.1111/jnc.15463. (Figures 1 and 2) [18].
Use extreme care not to disrupt the protein pellet during the removal of solvents or wash steps.
Gel tips should be used to ensure the protein pellet is not disrupted and when adding the samples to and from the PCT microtubes.
After sample preparation, samples can be run with any mass spectrometry and analysis method of the user’s choice.
Troubleshooting
Problem 1: Protein yield is low.
Possible cause 1: The sample underwent multiple freeze/thaw cycles.
Solution 1: Aim to avoid unnecessary freeze/thaw cycles as this can degrade protein, leading to lower identifiable proteins.
Possible cause 2: Incompatibility of the homogenization buffer with the sample.
Solution 2: Switching the homogenization buffer to a more suitable buffer, such as RIPA buffer, is effective for challenging proteins like nuclear or mitochondrial proteins. For example, 1 mL of RIPA buffer can replace the same volume of the current homogenization buffer (urea and TEAB) during initial homogenate preparation.
Acknowledgments
This protocol was validated in Puopolo et al. [15,16]. The authors acknowledge the RI-INBRE core facility at the University of Rhode Island, supported by Grant P20GM103430 from the National Center for Research Resources (NCRR), a component of the National Institutes of Health (NIH). Chang Liu, Ph.D., was supported by the Pilot Projects Program at the CardioPulmonary Vascular Biology (CPVB) Center for Biomedical Research Excellence (COBRE) and by a Pilot Grant from the College of Pharmacy, University of Rhode Island.
Competing interests
The authors have no competing interests to disclose.
Ethical considerations
This protocol was conducted in strict accordance with the University of Rhode Island’s Institutional Animal Care and Use Committee (IACUC) under protocol number AN1819-011 and the guidelines outlined in the “Animal Welfare Regulations” (AWRs) by the United States Department of Agriculture (USDA), the “Guide for the Care and Use of Laboratory Animals,” 8th Edition, by the National Institutes of Health (NIH) and the “Public Health Service Policy on Humane Care and Use of Laboratory Animals” (PHS Policy).
References
Duong, V. A. and Lee, H. (2023). Bottom-Up Proteomics: Advancements in Sample Preparation. Int J Mol Sci. 24(6): 5350. https://doi.org/10.3390/ijms24065350
Varnavides, G., Madern, M., Anrather, D., Hartl, N., Reiter, W. and Hartl, M. (2022). In Search of a Universal Method: A Comparative Survey of Bottom-Up Proteomics Sample Preparation Methods. J Proteome Res. 21(10): 2397–2411. https://doi.org/10.1021/acs.jproteome.2c00265
Milkovska-Stamenova, S., Wölk, M. and Hoffmann, R. (2021). Evaluation of Sample Preparation Strategies for Human Milk and Plasma Proteomics. Molecules. 26(22): 6816. https://doi.org/10.3390/molecules26226816
Snashall, C. M., Sutton, C. W., Faro, L. L., Ceresa, C., Ploeg, R. and Shaheed, S. u. (2023). Comparison of in-gel and in-solution proteolysis in the proteome profiling of organ perfusion solutions. Clin Proteomics. 20(1): 51. https://doi.org/10.1186/s12014-023-09440-x
Alfonso-Garrido, J., Garcia-Calvo, E. and Luque-Garcia, J. L. (2015). Sample preparation strategies for improving the identification of membrane proteins by mass spectrometry. Anal Bioanal Chem. 407(17): 4893–4905. https://doi.org/10.1007/s00216-015-8732-0
Shahinuzzaman, A. D. A., Chakrabarty, J. K., Fang, Z., Smith, D., Kamal, A. H. M. and Chowdhury, S. M. (2020). Improved in‐solution trypsin digestion method for methanol–chloroform precipitated cellular proteomics sample. J Sep Sci. 43(11): 2125–2132. https://doi.org/10.1002/jssc.201901273
Ludwig, K. R., Schroll, M. M. and Hummon, A. B. (2018). Comparison of In-Solution, FASP, and S-Trap Based Digestion Methods for Bottom-Up Proteomic Studies. J Proteome Res. 17(7): 2480–2490. https://doi.org/10.1021/acs.jproteome.8b00235
Wang, F., Veth, T., Kuipers, M., Altelaar, M. and Stecker, K. E. (2023). Optimized Suspension Trapping Method for Phosphoproteomics Sample Preparation. Anal Chem. 95(25): 9471–9479. https://doi.org/10.1021/acs.analchem.3c00324
Weke, K., Kote, S., Faktor, J., Al Shboul, S., Uwugiaren, N., Brennan, P. M., Goodlett, D. R., Hupp, T. R. and Dapic, I. (2022). DIA-MS proteome analysis of formalin-fixed paraffin-embedded glioblastoma tissues. Anal Chim Acta. 1204: 339695. https://doi.org/10.1016/j.aca.2022.339695
Nakayasu, E. S., Nicora, C. D., Sims, A. C., Burnum-Johnson, K. E., Kim, Y. M., Kyle, J. E., Matzke, M. M., Shukla, A. K., Chu, R. K., Schepmoes, A. A., et al. (2016). MPLEx: a Robust and Universal Protocol for Single-Sample Integrative Proteomic, Metabolomic, and Lipidomic Analyses. mSystems. 1(3): e00043–16. https://doi.org/10.1128/msystems.00043-16
Wessel, D. and Flügge, U. (1984). A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal Biochem. 138(1): 141–143. https://doi.org/10.1016/0003-2697(84)90782-6
Pirog, A., Faktor, J., Urban-Wojciuk, Z., Kote, S., Chruściel, E., Arcimowicz, Å., Marek-Trzonkowska, N., Vojtesek, B., Hupp, T. R., Al Shboul, S., et al. (2021). Comparison of different digestion methods for proteomic analysis of isolated cells and FFPE tissue samples. Talanta. 233: 122568. https://doi.org/10.1016/j.talanta.2021.122568
Li, H., Puopolo, T., Seeram, N. P., Liu, C. and Ma, H. (2024). Anti-Ferroptotic Effect of Cannabidiol in Human Skin Keratinocytes Characterized by Data-Independent Acquisition-Based Proteomics. J Nat Prod. 87(5): 1493–1499. https://doi.org/10.1021/acs.jnatprod.3c00759
Li, H., Deng, N., Puopolo, T., Jiang, X., Seeram, N. P., Liu, C. and Ma, H. (2023). Cannflavins A and B with Anti-Ferroptosis, Anti-Glycation, and Antioxidant Activities Protect Human Keratinocytes in a Cell Death Model with Erastin and Reactive Carbonyl Species. Nutrients. 15(21): 4565. https://doi.org/10.3390/nu15214565
Puopolo, T., Chen, Y., Ma, H., Liu, C. and Seeram, N. P. (2024). Exploring immunoregulatory properties of a phenolic-enriched maple syrup extract through integrated proteomics and in vitro assays. Food Funct. 15(1): 172–182. https://doi.org/10.1039/d3fo04026g
Puopolo, T., Li, H., Ma, H., Schrader, J. M., Liu, C. and Seeram, N. P. (2023). Uncovering the anti-inflammatory mechanisms of phenolic-enriched maple syrup extract in lipopolysaccharide-induced peritonitis in mice: insights from data-independent acquisition proteomics analysis. Food Funct. 14(14): 6690–6706. https://doi.org/10.1039/d3fo01386c
Schrader, J. M., Xu, F., Agostinucci, K. J., DaSilva, N. A. and Van Nostrand, W. E. (2024). Longitudinal markers of cerebral amyloid angiopathy and related inflammation in rTg-DI rats. Sci Rep. 14(1): 8441. https://doi.org/10.1038/s41598-024-59013-7
Schrader, J. M., Xu, F. and Van Nostrand, W. E. (2021). Distinct brain regional proteome changes in the rTg‐DI rat model of cerebral amyloid angiopathy. J Neurochem. 159(2): 273–291. https://doi.org/10.1111/jnc.15463
Article Information
Publication history
Received: May 20, 2024
Accepted: Jul 17, 2024
Available online: Aug 2, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
Systems Biology > Proteomics > Whole organism
Biochemistry > Protein > Isolation and purification
Molecular Biology > Protein > Detection
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Laser-Assisted Microdissection and High-Throughput RNA Sequencing of the Arabidopsis Gynoecium Medial and Lateral Domains
VL Valentín Luna-García
SF Stefan de Folter
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5056 Views: 568
Reviewed by: Samik BhattacharyaShengze Yao Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant Physiology May 2024
Abstract
For obtaining insights into gene networks during plant reproductive development, having transcriptomes of specific cells from developmental stages as starting points is very useful. During development, there is a balance between cell proliferation and differentiation, and many cell and tissue types are formed. While there is a wealth of transcriptome data available, it is mostly at the organ level and not at specific cell or tissue type level. Therefore, methods to isolate specific cell and tissue types are needed. One method is fluorescent activated cell sorting (FACS), but it has limitations such as requiring marker lines and protoplasting. Recently, single-cell/nuclei isolation methods have been developed; however, a minimum amount of genetic information (marker genes) is needed to annotate/predict the resulting cell clusters in these experiments. Another technique that has been known for some time is laser-assisted microdissection (LAM), where specific cells are microdissected and collected using a laser mounted on a microscope platform. This technique has advantages over the others because no fluorescent marker lines must be made, no marker genes must be known, and no protoplasting must be done. The LAM technique consists in tissue fixation, tissue embedding and sectioning using a microtome, microdissection and collection of the cells of interest on the microscope, and finally RNA extraction, library preparation, and RNA sequencing. In this protocol, we implement the use of normal slides instead of the membrane slides commonly used for LAM. We applied this protocol to obtain the transcriptomes of specific tissues during the development of the gynoecium of Arabidopsis.
Key features
• Laser-assisted microdissection (LAM) allows the isolation of specific cells or tissues.
• Normal slides can be used for LAM.
• It allows the identification of the transcriptional profiles of specific tissues of the Arabidopsis gynoecium.
Keywords: Laser-assisted microdissection LAM Specific cells and tissues Gynoecium Arabidopsis Transcriptomes RNA-Seq
Graphical overview
Laser-assisted microdissection (LAM) procedure of gynoecium tissues in Arabidopsis
Background
Multicellular organisms, such as animals and plants, are made up of many cells, which are grouped together to form different tissues and organs. Proper development requires complex molecular regulation, ranging from changes in gene expression and signaling events to hormonal activity. Despite the efforts made to understand these mechanisms that regulate development, the great morphological complexity of tissues or organs represents a limitation to creating tissue-specific expression profiles [1–4]. However, in recent years, several techniques have been successfully developed and implemented to isolate and characterize the transcriptomic profiles of specific cell populations, such as fluorescent activated cell sorting (FACS), single-cell isolation, single-nuclei isolation, and laser-assisted microdissection (LAM) [5]. LAM is a microscopy-based technique that has established itself as a good technique for isolating and collecting specific cell populations from their tissue context [6–11]. The LAM technique is distinguished from others for not requiring the generation of marker lines or protoplasts or previous information on a large number of marker genes; it only requires knowing the morphology of the tissues to be collected. These characteristics have allowed expanding LAM implementation to different organs of different plant species such as tomato fruit [12, 13], rice [14], California poppy [15], and Arabidopsis [16–18]. Here, we describe a protocol, built on previous work by Chávez Montes et al., 2016 [10], to perform LAM and RNA-seq of four tissue types from the Arabidopsis gynoecium: carpel marginal meristem (CMM) and septum (SEP), both tissues of the medial domain of the gynoecium, and the presumptive carpel wall (PC) and valves (VV), two tissues that make up the lateral domain of the gynoecium [19]. The aim was to identify the transcriptomic profiles that regulate their development. In the implementation of this protocol, the use of slides without the membrane was chosen instead of the commercially available slides (MembraneSlides) for LAM, which was better for the collection of the tissues of interest. Furthermore, we briefly describe the data analysis and gene expression visualization. This protocol should also be applicable for other plant species.
Materials and reagents
Biological materials
Arabidopsis thaliana plants (inflorescence tissue)
Reagents
Absolute ethyl alcohol for molecular biology (ethanol) (Karal, catalog number: 2003)
Glacial acetic acid (Fermont, catalog number: 03015)
Histo-Clear II (Electron Microscopy Sciences, catalog number: 64111-04)
Paraplast Plus® (paraffin) (Sigma-Aldrich, catalog number: P3683)
Methyl alcohol (Karal, catalog number: 2010)
Direct-zol RNA Microprep (Zymo Research, catalog number: R2063)
Solutions
Fixation solution (see Recipes)
Note: We do not recommend using formaldehyde (FDA; paraformaldehyde) as the fixation solution, because a cross-link reversal step would be needed for RNA extraction. Some discussion on fixation solutions is given in Chávez Montes et al. [10].
Recipes
Fixation solution
Reagent Final concentration Quantity or Volume
Ethanol (absolute) 75% (v/v) 3 mL
Glacial acetic acid 25% (v/v) 1 mL
Total n/a 4 mL
Laboratory supplies
Sterile falcon tubes
Beakers
Dissecting forceps
Steel inclusion molds 15 mm × 15 mm (Citotest)
Embedding cassettes without lid, color white (Simport Scientific, catalog number: M480-2)
Microscope slides 26 × 76 mm, thickness of 1 mm (LAUKA)
Low-profile disposable blades (Leica, model: 819)
Slide rack and glass staining dish with lid (LaboLan, catalog number: 29600001XL)
TubeCollector 2×200 CM II (Zeiss, catalog number: 415101-2000-410)
AdhesiveCap 200 clear (Zeiss, catalog number: 415190-9191-000)
MembraneSlide 1.0 OPEN (Zeiss, catalog number: 415101-4401-000)
Equipment
Vacuum pump (Gast, model: DOA-P704-AA)
Vacuum desiccator
Mini-rotator, 2–80 rpm w/disk & clamps (Glas-Col, catalog number: 099A MR1512)
Convection incubator (Binder, model: BD 23)
Paraffin dispenser (Leica, model: EG 1120)
Microtome (Leica, model: RM2035)
Slide warmer (Lab-Line Instruments, model: 26005)
PALM MicroBeam IV microscope with motorized RoboMover and CapMover (Zeiss, series number: 1214000156)
NanoDrop (optional)
Covaris S2 ultrasonicator (optional)
Qubit 4 fluorometer (optional)
Bioanalyzer 2100 (optional)
Software and datasets
PALM Robo software (Zeiss, v4)
Optional: R studio for RNA-seq data analysis
Procedure
Embedding inflorescence tissues in paraffin
Tissue fixation
Day 1. Collect inflorescence tissue (use dissecting forceps) and place the tissue directly into the fixation solution [ethanol: glacial acetic acid 3:1 (v/v)] on ice.
Notes:
i. Prepare the fixation solution fresh and keep cold.
ii. As a guideline, for 15–20 Arabidopsis inflorescences in a 50 mL Falcon tube or beaker, 10 mL of each solution (for this and the next steps) is needed. The tissue should always be covered with solution.
iii. Tissues of different genotypes or treatments should be maintained separated and always labeled.
Caution: RNA integrity should be conserved. Work clean and use gloves.
Apply vacuum for 15 min using a vacuum desiccator.
Notes:
i. The open Falcon tube or beaker is on ice in the vacuum desiccator.
ii. In our work, we use the maximum force of the vacuum pump, which is around -0.6 bar.
Replace the fixation solution and repeat the vacuum step for 15 min.
Leave the samples in fixation solution overnight at 4 °C.
Notes:
i. The tissue should sink during the vacuum steps. If not, repeat vacuum step.
ii. It is most practical to use a 50 mL Falcon tube(s) with cap from this point onward.
Dehydration
Day 2. Dehydrate the tissue samples using five steps of ethanol concentrations (70%, 80%, 90%, 100%, 100%). For each step, incubate the samples for 1 h at 4 °C. Constant agitation of 10 rpm may be used.
Leave samples in 100% ethanol at 4 °C overnight.
Pause point: Samples may be stored at 4 °C for more days if needed.
Histo-Clear II incubation
Day 3. Incubate the samples in a series of three steps with ethanol:Histo-Clear II (v/v) solutions [3:1, 1:1, 1:3 (v/v)]. For each step, incubate the samples for 1 h.
Note: Histo-Clear II is a replacement for the use of xylene, which is toxic.
Place the samples in 100% Histo-Clear II for 1 h.
Remove half of the Histo-Clear II volume by decanting and replace this volume with paraffin wax pellets (Paraplast Plus®). Leave overnight at room temperature.
Note: We recommend transferring the samples and solution to a small beaker(s), although Falcon tubes also work.
Paraffin incubation
Day 4. Place the beaker(s) with samples from the previous step in an incubator at 58 °C for 15–20 min (or until the Histo-Clear and paraffin mixture becomes liquid). During this step, all paraffin wax pellets will melt.
Notes:
i. We recommend the use of a convection incubator. Do not leave the door open too much time to avoid a drop in temperature. The use of a bigger incubator helps to maintain the temperature.
ii. The temperature of melting the paraffin is crucial and must be maintained according to the brand of paraffin in use.
Once the mixture is homogenized, decant a quarter of the total volume and replace this same volume with liquid paraffin at 58 °C. Incubate for 1 h at 58 °C in the incubator.
Note: For a correct homogenization of liquid paraffin, the paraffin dispenser should be turned on between 1 and 2 days prior to use; this depends on the type of paraffin dispenser and the amount of paraffin it contains. If no paraffin dispenser is available, Falcon tubes with melted paraffin stored at 58 °C may be used; these can be in the same incubator.
After 1 h, decant half of the total volume, replace this same volume with liquid paraffin at 58 °C, and place immediately back in the incubator at 58 °C for 1 h.
After 1 h, decant three-quarters of the total volume and replace this same volume with liquid paraffin at 58 °C. Incubate for 1 h at 58 °C in the incubator.
Note: From this step onward, the mixture is likely to start solidifying faster than in the previous steps; therefore, the work should be done as fast as possible to avoid solidification.
In the fourth wash, decant the whole mixture and replace it with liquid paraffin. Incubate again for 1 h at 58 °C in the incubator. Additional time may be needed depending on whether the paraffin is completely liquid; if not, incubate longer.
Repeat step A4e once or twice. If the smell of Histo-Clear II is no longer perceived, it is not necessary to continue with paraffin replacements.
After paraffin replacement steps, leave the samples in paraffin and incubate at 58 °C overnight in the incubator.
Embedding the tissue in paraffin blocks
Day 5. Prepare the paraffin blocks with the tissue (Figure 1). First, remove the beaker with inflorescences from the incubator and transfer the tissue with liquid paraffin to a hot plate (~58 °C), to maintain the liquid state of the paraffin.
Note: Some paraffin dispensers come coupled with a plate, but if you do not have this model, you can use any other hot plate. Practical tip: The plate can be covered with aluminum foil for easier cleaning afterward.
Place the steel inclusion molds also on top of the hot plate (~58 °C) to warm them; this helps for better handling of the melted paraffin during the generation of the blocks.
Next, add a small amount of liquid paraffin to the well of the inclusion mold. Subsequently, place the inflorescences (from 1 to 4 inflorescences per block) in the position you want to obtain the histological section (transverse or longitudinal). Then, place the plastic embedding cassette on top of the mold and fill the mold completely with liquid paraffin, let it cool, and then let it solidify on ice. See Figure 1 for various parts of the preparation of a paraffin block.
Store the block(s) at 4 °C until use.
Critical: Maintain the blocks free of RNase.
Figure 1. Overview of the complete laser-assisted microdissection (LAM) protocol. The protocol starts with plant tissue fixation, followed by tissue embedding in paraffin blocks, specific tissue/cell collection with the microscope, and finally RNA extraction, library preparation, and RNA sequencing. Taken from Luna-García et al. [19] (Supplementary Figure S1) with permission.
Histological sections
Day 6. Prepare the histological sections from the paraffin blocks with the tissue of interest using a microtome. In our experience, a thickness from 8 to 12 microns works well for the microdissection of Arabidopsis gynoecium tissues (Figure 1).
Notes:
The thickness of the histological sections can vary according to the tissue and the needs of each project.
Previous experience with a microtome is strongly advised.
Observe the cross sections with the help of an optical microscope; use only those sections in which the tissue of interest is present. Once the histological sections to be used have been identified, place them on a slide(s) as described in the next step.
Note: Slides with a membrane (typically used) to maintain cell integrity during LAM can be used, but for the development of this protocol with Arabidopsis inflorescences, slides without membrane were used since they allowed a better manipulation during microdissection.
Place slides on a slide warmer at ~42 °C. Add pure methanol to each slide surface. Place/float the selected paraffin sections (ribbons of paraffin sections; Figure 1) on the methanol, remove the excess methanol, and let the slides dry for ~45–60 min on the slide warmer. This will allow the fixation of the histological sections to the slides.
Notes:
The slides with fixed sections can be used directly in the next step of deparaffinization and LAM; alternatively, the non-deparaffinized slides can be stored for 1–2 days maximum at 4 °C.
We did not evaluate that but, when working with slides stored for longer times, the amount of extracted RNA from the LAM tissue decreased.
We use, as others do, pure methanol on the slide surface instead of water to minimize RNase activity.
Caution: Methanol is highly toxic.
Laser-assisted microdissection (LAM) of medial and lateral gynoecia tissues
Day 7. Deparaffinize the slides by placing them in a slide rack in a glass staining dish with 100% Histo-Clear II for 15 min (Figure 1). Subsequently, let the slides dry at room temperature for ~20–30 min. Afterward, start with the microdissection of the tissues of interest as described in the next steps.
Critical: Remember to keep everything free of RNase.
Laser-microdissection of tissue of interest
Turn on the PALM microscope (Figure 2A) and open the PALM Robo v4 software (Figure 3).
To place the slide(s) on the PALM stage (Figure 2B, C), click on button A (Load position) in the software (Figure 3A); the stage will move. Place slide(s) and click again on button A to let the stage return to the working position.
To locate the tissue of interest, select the appropriate objective by clicking on button B in the software (Figure 3A) where an objective, with which the PALM is equipped, can be selected (5×, 10×, 20×, 40×, 63×). With the help of the correct objective, locate the tissues of interest. Use button C to help focus (Autofocus; Figure 3A).
Note: In our work, depending on the gynoecium stage/size, we used either the 10× or the 20× objective.
To select the tissue to be isolated by LAM, first select the tissue to isolate with the drawing tool in the software by clicking on button D (Freehand; Figure 3A). The drawing tool can be manipulated with the mouse or with the special pencil on the touch-sensitive monitor. Start selecting the tissue to isolate by drawing with the pen a closed circle/region of interest on the screen (Figure 3B, indicated with a red line); the laser will later cut over this drawn line. Next, to mark the catapulting of the selected tissue, click on button E (Dot; Figure 3A) and, with the pencil, mark by dots (or single dot) inside the previously drawn region to be catapulted (Figure 3B, indicated with blue dots), toward the collecting tubes (Figure 2C, E). After this process, reactivate the selection option by clicking on button F (Pointer; Figure 3A).
Note: Many regions can be selected and collected by cutting and catapulting in series. In our work, per gynoecium, we collected the medial and lateral domains (in two different tubes); subsequently, we repeated it for the next gynoecium, and so on. In our hands, selecting regions in many gynoecia and then trying to collect all selected tissues did not work well, resulting, at a certain moment, in not properly cutting (not cutting completely through the section; probably loss of focus) or not accurately.
To collect the selected tissue, place the TubeCollector 2×200 CM II on the microscope CapMover with the collecting tube(s) placed (Figure 2D, E). Click on button G (Cap Check; Figure 3A); the collecting tube will travel from its initial position to the position a few millimeters above the slide with the sample to be collected. To isolate and catapult the tissue of interest, activate the laser by clicking the button H (Figure 3A) and then let the sequence of cutting and catapulting actions run by clicking the button I (Start Cutting Laser; Figure 3B, Video 1).
Notes:
i. Before starting the process of cutting and catapulting the tissues of interest, test the laser energy and thickness to find the ideal working parameters by modifying the parameters of the part of the software indicated by the letter J (Energy and Focus; Figure 3A). Do this by trial and error prior to a real experiment. In our experiment, as a guideline, we used the following settings: laser cut energy 50–70; speed 10–50; LPC energy 60–75. Cut Focus and LPC focus change when the focus of the microscope changes, so we did not adjust them manually. The number of cycles of laser cutting is normally one. The z-focus parameter was not changed manually.
ii. Use a single collection tube to collect a single tissue type of interest; you can perform this step as many times as necessary using the same tube to collect enough biological material for a good amount of RNA.
iii. For our experiment, for one tissue type (for one RNA-seq sample), it took three days to collect the tissue from 20–30 slides with inflorescence tissue (i.e., different gynoecia stages were present, and only specific stages were selected for our experiment).
iv. In general, microdissected material of around 80 gynoecia (sections of around 24 paraffin blocks with three or four embedded inflorescences) was pooled to result in one tissue sample (one replicate).
Figure 2. PALM microscope. A. PALM microscope, with which the laser-microdissection of the tissues of interest is performed. B, C. Area of the stage, where the slide with the transverse sections (previously deparaffinized), as well as the TubeCollector 2×200 CM II, are placed. D, E. TubeCollector 2×200 CM II and AdhesiveCap 200 clear tubes.
Video 1. Laser-assisted microdissection (LAM) of medial and lateral gynoecium tissues
Figure 3. PALM software. A. PALM Robo v4 software and various parameters used for microdissection of tissues of interest. B. The area to be cut by the laser is in red and the amount of spot energy to be applied to catapult the tissues of interest into the collecting tubes is in blue. Letters in (A) indicate the important buttons in the menu and order of control for performing LAM, as written in section C of the protocol.
Once the collection of the tissue of interest is completed, the collection tubes are placed in liquid nitrogen and stored at -80 °C until further use (RNA extraction).
Note: For our experiment, 12 h per day of LAM was performed, resulting in the collection of two tissue types (one from the medial and one from the lateral domain), collected in the 2 × 200 µL tubes in the TubeCollector. This process was repeated twice, so in total 18 h (6 h/tissue × 3 days) was needed for one sample of one tissue type.
To finalize the PALM work session, the reverse workflow is followed; close PALM software and close Windows software to turn off the computer, followed by turning off the microscope.
RNA Extraction and library preparation
After sufficient LAM of the tissue(s) of interest, perform RNA extraction. For this protocol, we used the Direct-zol RNA Microprep kit, which allows yields of ~100–200 ng of total RNA in a volume of 8–10 µL.
Note: For our experiment, we had three tubes with collected tissue that made up one library/RNA-seq sample. Tubes were retrieved from -80 °C storage and placed in liquid nitrogen. When RNA extraction started, tubes were taken out of the liquid nitrogen; 100 µL of Trizol was added to each tube (to the cap of the tube and the tube itself), followed by pooling the solution of the three tubes to one new Eppendorf tube. Subsequently, the instructions of the kit were followed. We did not perform any grinding of the samples.
Note: During the RNA extraction procedure, a DNase I treatment is performed on the column to remove genomic DNA.
Note: We use a NanoDrop spectrophotometer to check RNA quality and quantity.
For cDNA preparation, different kits are available. For cDNA synthesis, we use the SMART-Seq v4 Ultra Low Input RNA Kit for Sequencing (Takara Bio).
Notes:
For our experiment, we used a Covaris S2 ultrasonicator to shear the cDNA into around 100 bp fragments.
For our experiment, we analyzed the cDNA quality and quantity with a Qubit 4 fluorometer and a Bioanalyzer 2100.
For sequence library preparation, different kits are available. For library preparations, we use the NEBNext Ultra II DNA Library Prep Kit for Illumina (New England Biolabs) and the NEBNext Multiplex Oligos for Illumina (Index Primers Set 1; New England Biolabs).
Note: The sequence libraries normally can be prepared as part of the RNA sequence service, which will save time. For our experiment, we did it in-house to avoid sending low quantities of RNA and country-related complicated logistics.
RNA sequencing
Send samples/libraries to an RNA sequencing service. The data used in our publication was generated using a HiSeq2000 Illumina resulting in paired-end libraries with an average of 43 million reads (150 bp paired-end) after filtering adapters and low-quality reads.
RNA-seq data analysis can be requested as part of the sequencing service or can be done in-house. We refer to our publication or other literature available for information on data analysis.
Note: Expertise is required for data analysis when starting with raw reads.
Data analysis
Bioinformatic analysis may be solicited as part of the RNA sequence service. If so, lists with genes and their respective expression values in transcripts per million (TPM) are obtained. Depending on the experiment, one or more lists are provided with differentially expressed genes (DEGs) given in the log-transformed of the expression difference. Often, a cutoff is used, for instance, a log-transformed fold change (logFC) > 1 and an FDR < 0.05, although a more severe cutoff can also be used (e.g., logFC > 2). This depends on the aim. When using a cutoff of logFC > 2, a shorter list of DEGs will be obtained.
As with any RNA-seq experiment, nowadays, it is important to perform this in triplicate, meaning using biological replicates. A principal component analysis (PCA) can be performed to observe the reproducibility of the experiment. Furthermore, gene ontology (GO) analysis can be performed, among others.
When opting for in-house data analysis, some skills are necessary, and it is handy to know Linux and R. For a description of the data analyses, see our article Luna-García et al. [19]. In short, the quality of raw sequencing reads was analyzed using FASTQC v0.11.5 [20], and overrepresented sequences, low-quality reads, and adapters were removed using trimmomatic v0.39 [21] and cutadapt v2.8 [22]. The filtered sequencing reads (Phred Quality Score > 30) were mapped to the Arabidopsis thaliana reference transcriptome (Araport11) using Kallisto v0.46.1 [23]. Read counts and TPMs were generated using the R package tximport v1.0.3 and the lengthScaledTPM method [24]. For DEG analysis, use the files with extension.h5 (Hierarchical Data Format 5), generated in alignment with Kallisto, processed in RStudio with R packages tximport v1.12.3 [24] and edgeR v3.26.8 [25], following part of the workflow by Chen and collaborators [26].
Validation of protocol
This protocol has been used and validated in the following research article:
Luna-García et al. [19]. A high-resolution gene expression map of the medial and lateral domains of the gynoecium of Arabidopsis. Plant Physiology.
As presented in Luna-García et al. [19] (in Supplementary Figure S2), in Figure 4 here, the result of the PCA of our data is presented, where the replicates group nicely together, indicating the reproducibility of the experiment. Details on the RNA quality and the raw reads for each library can be found in Luna-García et al. [19] (in Supplementary Table S1).
Subsequently, possible data validation methods include searching public data to see if gene expression is similar/identical, for instance, in other RNA-seq data, RT-qPCR data of specific tissues, or published in situ hybridization results. In our article, we compared our data with 18 published in situ hybridization results of well-known transcription factors, which nicely confirmed our data [19] (in Supplementary Table S22). We presented a heatmap of these 18 well-known transcription factors, demonstrating the results and specificity of the data [19] (in Supplementary Table S22) and, in this protocol, Figure 4B.
Lastly, we would like to mention that data availability is very important. Besides presenting annotated (Excel) tables as part of a research article, depositing data in known databases such as the European Nucleotide Archive (ENA) is essential (data of Luna-García et al. [19] is stored in ENA under the accession number PRJEB65130). Furthermore, data from our work in collaboration with the Arabidopsis eFP Browser platform [27] is fully searchable in this publicly available tool (https://bar.utoronto.ca/efp_arabidopsis/cgi-bin/efpWeb.cgi?dataSource=Gynoecium); Figure 4C gives an example for how this looks like in the eFP browser for two genes.
Figure 4. Examples of data analyses approaches. A. Results of the principal component analysis (PCA) of the article by Luna-García et al. [19]. Four different tissue types in triplicate have been isolated by LAM, followed by RNA-seq, where the replicates nicely group together. CMM, carpel margin meristem; PC, presumptive carpel wall; SEP, septum; VV, valves. B. Heatmap of the expression values of 18 well-known transcription factors. Values expressed in Z-score. Yellow means higher and blue means lower expression. These genes are also identified as DEGs in the tissues. C. eFP browser views of expression profiles for FILAMENTOUS FLOWER (FIL), a DEG of the lateral domain of the gynoecium, and SPATULA (SPT), a DEG of the medial domain of the gynoecium; these views exemplify the visualization of our data available in the Arabidopsis eFP Browser [27]. Red means high expression, and yellow means low expression. Modified from Luna-García et al. [19] (from Supplementary Figure S2 and Figure 4) with permission.
General notes and troubleshooting
General notes
Work RNase-free during all wet lab experiments. In our experiments, we use gloves; before starting work, we clean the table, pipettes, and centrifuge with ethanol, use sterilized pipette tips, lab coat, and general hygiene. Furthermore, during RNA extraction, use ice to keep samples cold. The aim is to isolate RNA to analyze the transcriptome of the tissue of interest.
Obtain previous experience with the microtome and all histological steps.
Obtain previous experience with the LAM microscope.
Consider that a LAM protocol is time-consuming.
The software manual can be downloaded for more detailed information on its use (PALM RoboSoftware 4.5 Quick Guide).
This protocol should also be applicable to other species. Verify if tissue fixation time should be increased or decreased because this is species-specific. Thicker tissue requires increased fixation time.
Troubleshooting
Problem 1: During paraffine block sectioning with the microtome, the tissue falls out.
Possible cause: Tissue is not well fixed.
Solution: Collect new tissue for fixation and prepare new paraffin blocks.
Problem 2: RNA extraction results in no or low RNA quantity.
Possible cause(s): RNA is degraded during slide preparation or problem during RNA extraction.
Solution(s): Work free of RNase during the complete protocol and divide the protocol into steps to find out where the problem of RNA degradation is.
Problem 3: Laser-assisted cutting or catapulting is not working well.
Possible cause(s): Incorrect settings in the PALM software (e.g., focus, energy, and speed).
Solution: Adjust settings in the PALM software.
Problem 4: Data analysis does not show reproducibility (e.g., in the PCA).
Possible cause(s): Different plant/tissue stages used, or lack of tissue-specificity collection during LAM.
Solution(s): Here, the possibilities may be limited. Depending on what is observed, a possible solution could be to not consider one replicate, or in the worst scenario, it probably means that you have to repeat the LAM and RNA-seq experiments.
Acknowledgments
V.L.G. was supported by the Consejo Nacional de Humanidades, Ciencias y Tecnologías (CONAHCYT, Mexico) with a PhD fellowship (487657). S.d.F. acknowledges UGA-Langebio intramural funds for the acquisition of the PALM microscope. The work in the S.d.F. laboratory was financed by the CONAHCYT grants CB-2012-177739, FC-2015-2/1061 and CB-2017-2018-A1-S-10126. S.d.F. is grateful for the participation in the European Union project H2020-MSCA-RISE-2020 EVOfruland (101007738). The protocol is modified from Chávez Montes et al. [10] and used in Luna-García et al. [19].
Competing interests
The authors declare no competing interests.
References
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Article Information
Publication history
Received: May 9, 2024
Accepted: Jul 22, 2024
Available online: Aug 2, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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An Imaging-Based Assay to Measure the Location of PD-1 at the Immune Synapse for Testing the Binding Efficacy of Anti-PD-1 and Anti-PD-L1 Antibodies
JZ Justin C. Zhong
SL Shalom Lerrer
AM Adam Mor
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5057 Views: 567
Reviewed by: Emilie BesnardPaurvi ShindeKerui Huang
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Original Research Article:
The authors used this protocol in Molecular Therapy Oncology Sep 2024
Abstract
PD-1 is an immune checkpoint on T cells. Antibodies to PD-1 or its ligand PD-L1 are gaining popularity as a leading immunotherapy approach. In the US, 40% of all cancer patients will be treated with anti-PD-1 or anti-PD-L1 antibodies but, unfortunately, only 30% will respond, and many will develop immune-related adverse events. There are nine FDA-approved anti-PD-1/PD-L1 antibodies, and approximately 100 are in different stages of clinical development. It is a clinical challenge to choose the correct antibody for a given patient, and this is critical in advanced malignancies, which often do not permit a second-line intervention. To resolve that, an in vitro assay to compare the performance of the different anti-PD-1/PD-L1 antibodies is not only a critical tool for research purposes but also a possible tool for personalized medicine. There are some assays describing the binding affinity and function of anti-PD-1/PD-L1 antibodies. However, a significant limitation of existing assays is that they need to consider the location of PD-1 in the immune synapse, the interface between the T cell and tumor cells, and, therefore, ignore a critical component in its biology. To address this, we developed and validated an imaging-based assay to quantify and compare the ability of different anti-PD-1/PD-L1 antibodies to remove PD-1 from the immune synapse. We correlated that with the same antibodies' ability to increase cytokine secretion from the targeted cells. The strong correlation between PD-1 location and its function in vitro and in vivo within the antibody treatment setting validates this assay's usability, which is easily recordable and straightforward.
Key features
• Live-cell imaging quantifies and compares how anti-PD-1 and anti-PD-L1 antibodies disrupt PD-1 localization, causing the removal of PD-1 during immune synapse formation.
• Hao et al. [1] validated the protocol, and the findings were extended to a live confocal microscopy method.
• It requires a Zeiss LSM 900 confocal microscope and appropriate imaging software and is optimized for the latest version of Zen Blue.
• Anti-PD-1 antibodies are commonly used in cancer therapies, and this protocol optimizes the analysis of their effectiveness.
Keywords: PD-1 PD-L1 B cells Confocal microscopy Immunotherapy T cells Cancer Immune checkpoints Immune synapse
Graphical overview
Background
PD-1, or programmed cell death protein 1, is a protein found on the surface of T cells. PD-1 acts as a brake on T cells, preventing them from overreacting and attacking healthy tissues. This is important for maintaining immune tolerance and preventing autoimmune diseases. However, cancer cells can sometimes exploit the PD-1 pathway to evade immune attacks. They can do this by expressing high levels of PD-L1 (programmed cell death ligand 1), the ligand for PD-1 that inhibits T-cell activity, and in some cases also PD-L2, a higher-affinity ligand for PD-1 [2].
This is where PD-1 inhibitors come in. These drugs block the interaction between PD-1 and PD-L1, allowing T cells to recognize and destroy cancer cells [3,4]. PD-1 and PD-L1 inhibitors have revolutionized cancer treatment, and they are now used to treat a variety of cancers including melanoma, lung cancer, and head and neck cancer. Unlike traditional chemotherapy, which often has short-lived effects, anti-PD-1/PD-L1 antibodies can induce long-lasting tumor shrinkage and even complete remission in some patients. Compared to chemotherapy, anti-PD-1/PD-L1 antibodies have a more favorable side effect profile. While they can cause immune-related adverse events, these are generally less severe and more manageable than the toxicities associated with chemotherapy. Therefore, anti-PD-1/PD-L1 antibodies have become the standard of care for many cancer types, and their use is rapidly expanding to new applications. This provides more treatment options for patients with advanced or aggressive cancers who may not have had many choices before. Unfortunately, not all patients respond to anti-PD-1 therapy [5]. The reasons for this are complex and not fully understood. Still, it highlights the need for further research to identify biomarkers and in vitro assays that can predict who will benefit most from this treatment.
There are currently nine FDA-approved anti-PD-1/PD-L1 antibodies [6], each with its specific indications and potential side effects: pembrolizumab (Keytruda), nivolumab (Opdivo), cemiplimab (Libtayo), atezolizumab (Tecentriq), avelumab (Bavencio), durvalumab (Imfinzi), dostarlimab (Jemperli), toripalimab (Loqtorzi), and tislelizumab (Tevimbra). It is important to note that this is a partial list, and the number of anti-PD-1 and PD-L1 antibodies in clinical trials continually evolves. Still, over 100 different anti-PD-1 and PD-L1 antibodies are currently in various stages of clinical development, according to the ClinicalTrials.gov database [7]. This number encompasses multiple anti-PD-1/PD-L1 agents, including monoclonal antibodies, bispecific antibodies, and antibody-drug conjugates (ADCs).
With so many different anti-PD-1/PD-L1 antibodies in development, there are several reasons why in vitro assays are crucial for studying them. In vitro assays offer a safe and relatively inexpensive way to test the properties of these antibodies before moving to complex and expensive animal or human studies. In vitro assays provide a controlled environment to study antibodies' specific mechanisms of action [8]. They can help understand how these antibodies interact with PD-1/PD-L1, T, and cancer cells, ultimately leading to better drugs and treatment strategies. In vitro assays can also identify biomarkers that predict which patients are more likely to respond to specific anti-PD-1/PD-L1 therapy [9]. This personalized approach can help optimize treatment for individual patients and avoid unnecessary side effects for those unlikely to benefit. In vitro assays help compare different formulations or combinations of antibodies, allowing the optimization of their efficacy and safety before clinical trials.
Promega developed an assay to measure the ability of anti-PD-1/PD-L1 antibodies to block immune checkpoint signals [10]. The assay consists of two genetically engineered cell lines, PD-1 effector cells and PD-L1 aAPC/CHO-K1 cells. When cocultured, the PD-1/PD-L1 interaction inhibits T-cell receptor (TCR)-mediated luminescence. When the PD-1/PD-L1 interaction is disrupted, TCR activation induces luminescence via activation of the NFAT pathway, which can be detected by adding bio-glo reagent and quantitation with a luminometer. Another report described a flow cytometry assay to evaluate the T-cell binding status of the anti-PD-1 antibodies using a single drop of peripheral blood [11]. Another group established a PD-1/PD-L1 blockade assay based on surface plasmon resonance (SPR) biosensors [12]. This assay immobilizes human PD-1 on a sensor chip, where the binding kinetics of PD-L1 to PD-1 and the blockade rates of PD-1 inhibitors are determined. Compared to other techniques, such as PD-1/PD-L1 pair ELISA, this method offers real-time and label-free detection with advantages including shorter experimental runs and more minor sample quantity requirements. We and others have published in vitro assays used to study the effects of anti-PD-1 antibodies on T-cell proliferation, cytotoxicity (to assess the killing ability of T cells activated by PD-1 blockade against cancer cell lines), and cytokine secretion [13], providing insights into the immune response triggered by PD-1. While in vitro assays are valuable tools, they have limitations. They cannot fully replicate the complex interactions in the human body, and validating findings with in vivo studies and clinical trials is crucial.
The formation of a synapse between a T cell and a tumor cell is a crucial initiator of the immune response, with interactions between the major histocompatibility complex (MHC) and TCR being essential for T-cell activation and killing [14]. Synapse formation causes the clustering of proteins in different regions of the contact area, called supramolecular activation complexes (SMACs). SMACs can be further separated into different subregions: the central SMAC (cSMAC), the peripheral SMAC (pSMAC), and the distal-SMAC (dSMAC), where the cSMAC contains the proteins and checkpoint receptors of interest for this assay [15]. PD-1 and PD-L1 are a part of the group of molecules that centralize to the cSMAC during the formation of the synapse [16]. Interactions between PD-1 and PD-L1 activate the phosphatase SHP2, causing suppression of T-cell activity and overall immune response by preventing stable T-cell interactions from forming. PD-L1 and PD-1 can also develop microclusters within the cSMAC, contributing to their suppressive function [17]. The localization of PD-1 in the synapse when bound or unbound by antibodies allows it to be a prime target for studying the efficacy of different antibodies. Specifically, antibody binding mislocalizes PD-1 and prevents interaction with PD-L1 [1]. The exclusion of antibody-bound PD-1 can be considered size-based, as the immune synapse is observed to exclude molecules above a specific size limit [18]. Removing PD-1 from the synapse prevents PD-1/PD-L1 clustering within the cSMAC and interactions with PD-L1 for inhibitory function. As such, antibodies with different characteristics may affect the localization of PD-1 in the immune synapse to different degrees.
Previously, we reported an in vitro assay to study PD-1 signaling in primary human T cells [13], based on the phospho-flow cytometry method to determine how PD-1 ligation alters the levels of CD3ζ phosphorylation on Tyr142, which can be easily applied to other proximal signaling proteins. We also reported a plate-bound assay to examine the long-term consequences of PD-1 ligation, such as cytokine production and T-cell proliferation. We also described an in vitro superantigen-based assay to evaluate T-cell responses to therapeutic agents targeting the PD-1/PD-L1 axis and immune synapse formation in the presence of PD-1 engagement [13]. Based on this, we developed an imaging-based assay, described here. This protocol utilizes PD-1 GFP-expressing Jurkat T cells and PD-L1-/PD-L2 mCherry-expressing Raji B cells, yielding a model to study the function and location of anti-PD-1/PD-L1/PD-L2 antibodies in real time. This study design requires a Zeiss LSM 900 confocal microscope and an updated version of Zen Blue software, at least version 3.3. Compared with other methods of quantifying PD-1 antibody strength, such as flow cytometry, this method allows for both visual and numerical analysis of synapse formation between B and T cells, simultaneously providing qualitative and quantitative data. Furthermore, it enables live-cell imaging, captures temporal data, and gives insights into the dynamics of these interactions over time. This protocol was validated by Hao et al., 2024.
Materials and reagents
Biological materials
Jurkat T cells (ATCC, catalog number: TIB-152)
Raji B cells (ATCC, catalog number: CCL-86)
Staphylococcus enterotoxin E (SEE) ET404 1 mg (Toxin Technology, catalog number: NC1467973)
Reagents
RPMI 1640 (Thermo Fisher Scientific, GibcoTM, catalog number: 31800)
Opti-MEM (Thermo Fisher Scientific, catalog number: 11058021)
Heat-inactivated fetal bovine serum (FBS) (Thermo Fisher Scientific, GibcoTM, catalog number: 10082147)
Penicillin/streptomycin (P/S) (Thermo Fisher Scientific, GibcoTM, catalog number: 15140)
Human CD3/CD28/CD2 T-cell activator (StemCell Technologies, ImmunoCultTM, catalog number: 10970) (optional; see protocol)
10× Poly-L-Lysine (PLL) solution (0.1% w/v in H2O) (Sigma-Aldrich, Millipore, catalog number: P8920)
Trypan Blue stain (0.4%) (Thermo Fisher Scientific, InvitrogenTM, catalog number: T10282)
Molecular biology grade water (Corning, catalog number: 46-000-CI)
pHR-PD-1-GFP vector and pHR-PD-L1-mCherry vector (AddGene, catalog number: 180819) [19]
Solutions
1× Poly-L-Lysine solution (see Recipes)
RPMI (w/ FBS) media (see Recipes)
Staphylococcus enterotoxin E (SEE) working stock (see Recipes)
Recipes
1× Poly-L-Lysine solution
Reagent Final concentration Amount
10× PLL [0.1 % (w/v) in H2O] 1× 100 µL
ddH2O n/a 900 µL
Total n/a 1,000 µL
RPMI (w/ FBS) media
Reagent Final concentration Amount
RPMI 1640 n/a 500 mL
FBS 10% 55 mL
P/S 1% 5.5 mL
Total n/a 560.5 mL
Staphylococcus enterotoxin E (SEE) working stock (stored at -20 °C)
Reagent Final concentration Amount
Staphylococcus enterotoxin E 1 mg/mL 20 µg
ddH2O n/a 20 µL
Total n/a 20 µL
Laboratory supplies
15 mL and 50 mL conical tube (Falcon, catalog numbers: 38009, 38010)
12-well cell culture plate (Thermo Fisher Scientific, catalog number: 150200)
Untreated cell culture flask with vent cap (Thermo Fisher Scientific, CorningTM, catalog number: 431463)
MatTek Plates (50 mm dish, No. 0 coverslip, 14 mm glass diameter, uncoated) (MatTek, catalog number: P50G-0-14-F)
Single-channel pipette, Pipetman 0.2–2 µL, 1–10 µL, 2–20 µL, 20–200 µL, 100–1,000 µL (Gilson, catalog numbers: F144054M, F144055M, F144058M, F144059M)
Filtered pipette tips (USA Scientific)
1.5 mL microcentrifuge tubes (Fisher Scientific, catalog number: 05-408-129)
Immersol autofluorescence-free immersion oil 518 F (Zeiss, catalog number: 444960-0000-000)
Aluminum foil (Amazon brand)
Cell counting chamber slides (CountessTM, catalog number: C10228)
KOVATM GlassticTM slide 10 with grids (Fisher Scientific, catalog number: 22-270141)
Cell culture multiwell plate 24 wells (Grenier Bio-One, catalog number: 662160)
Equipment
Confocal microscope (Zeiss, model: LSM 900)
Heracell 150i CO2 incubator (37 °C, 5% CO2) (Thermo Fisher Scientific, catalog number: 50116048)
Digital miniblock heater (VWR, catalog number: 460-0334)
Automated cell counter (InvitrogenTM, CountessTM, catalog number: AMQAX2000)
FisherbrandTM manual differential counter (Fisher Scientific, catalog number: 13-684-141)
5425 centrifuge (Eppendorf, catalog number: 5405000646)
5810R centrifuge (Eppendorf, catalog number: 022625501)
Software and datasets
Image J (Version 1.54)
Zen (Version 3.3 Blue Edition)
Procedure
Preparation of T and B cells
Note: For optimal imaging, the assay should be done with a fluorescent PD-1-expressing T-cell line (such as Jurkat T cells expressing pHR-PD-1-GFP) and a fluorescent PD-L1-expressing antigen-presenting cell line (such as Raji B cells expressing pHR-PD-L1-mCherry). However, primary T cells can also be used following transfection/viral transduction with a fluorescent PD-1 expression vector in combination with Raji cells [20]. Fluorescent T and B cell lines can be generated as described in Lerrer et al. [21].
Day 1
Note: If activation via CD3/CD28/CD2 ImmunoCult T-cell activator for T cells is needed, the steps delineated for day 1 and below for day 2 must be done separately. Otherwise, the steps for day 1 and day 2 can be combined into a single day. See Figure 1.
Figure 1. Timeline of the cell preparation protocol. Timelines for the protocol A) with T-cell activation or B) without T-cell activation are given. Steps shown on the same tick mark are to be completed in parallel. Steps should be completed in the order shown and according to the protocol written.
Maintain T and B cells in T75 flasks with 12–15 mL of RPMI (w/ FBS) media (see Recipes) at the appropriate optimal growing concentrations.
~2 × 105 to 2 × 106 cells/mL are optimal growing concentrations for Jurkat.
~3 × 105 to 2 × 106 cells/mL are optimal growing concentrations for Raji.
Collect around 5 mL from flasks and ensure that any clumps are broken up with pipetting.
Count viable T cells manually or using an automated cell counter with Trypan Blue.
Take 10 µL of cells and 10 µL of Trypan Blue.
Mix with a pipette.
If counting using an automated cell counter:
i. Add 10 µL of the mixture to a cell counting chamber slide.
ii. Insert the slide into an automated cell counter, with the side containing the cell/Trypan mixture inserted first.
iii. Focus the machine, press scan, and record cell viability as reported.
If counting manually:
i. Take 10 µL of the mixture and add it to a single chamber of the KOVATM GlassticTM slide.
ii. Place the slide under a microscope and count the number of viable cells per grid using an appropriate magnification to visualize cells in each box.
1) Viable cells will appear shiny or bright, whereas nonviable or dead cells will be permeable to the Trypan Blue dye and appear darker.
iii. Calculate the number of viable cells by converting 10 viable cells/box = 2 million viable cells/mL.
1) Count multiple boxes, convert, and then average the resulting viable cell numbers together to get the most accurate estimate of total cell viability.
Calculate the appropriate volume needed for 106 viable cells (or more, if desired) and centrifuge the cells at 500× g for 5 min in a microcentrifuge tube.
If needed, centrifuge in a 15 mL or 50 mL tube at 500× g, and then resuspend and transfer to a microcentrifuge tube.
Remove the supernatant and leave around 50 µL of media.
Add fresh RPMI (w/ FBS) media to a final concentration of 106 cells per milliliter (950 µL of media if 106 cells were used in step A5).
Move isolated 106 cells (total volume 1 mL) to one well of a 24-well cell culture plate for overnight culture.
Add 12.5 µL of CD3/CD28/CD2 ImmunoCult T-cell activator for T cells per milliliter and incubate overnight (not necessary for PD-1-expressing Jurkat cells) at 37 °C in a tissue culture incubator.
Day 2
Note: Both day 1 and day 2 steps can be done on the same day if overnight T-cell activation is not necessary, e.g., when using PD-1-expressing Jurkat cells. See Figure 1.
Repeat steps 2–6 from Day 1 for B cells or perform in parallel for B and T cells if overnight T-cell activation was not performed.
Collect around 5 mL from flasks and ensure that any clumps are broken up with pipetting.
Count viable cells manually or using an automated cell counter with Trypan Blue, as described in the steps for day 1.
Calculate the appropriate volume needed for 106 viable cells (or more, if desired) and centrifuge them in a microcentrifuge tube at 500× g for 5 min.
If needed, centrifuge in a 15 mL or 50 mL tube at 500× g, and then resuspend and transfer to a microcentrifuge tube.
Remove the supernatant and leave around 50 µL of media.
Add Opti-MEM media to a final concentration of 106 cells per milliliter (950 µL of media if 106 cells were used in step 1d).
If T-cell activation is required, change the media with the T-cell activator solution to fresh media.
Collect cells from the well into a microcentrifuge tube.
Spin cells down at 500× g for 5 min.
If desired, this spin can be done at the same time as the spin for step 1.
Remove supernatant, leaving up to 50 µL above the cells, and resuspend in 1 mL Opti-MEM media.
Add 5 µL of Staphylococcus enterotoxin E (SEE) working stock (see Recipes) for each 1 mL of B cells and incubate at 37 °C in a tissue culture incubator for 30–60 min in the microcentrifuge tube.
Add the desired amount of anti-PD-1/PD-L1 antibodies to the appropriate tube and incubate at 37 °C in a tissue culture incubator for 30–60 min in the microcentrifuge tubes:
If an anti-PD-1 antibody is to be tested, add the antibody to the T-cell tube.
i. Add the anti-PD-1 antibody in parallel with the Staphylococcus enterotoxin E (SEE) so that both T and B cells will incubate for 30–60 min.
If an anti-PD-L1 antibody is to be tested, add the antibody to the B-cell tube.
i. Add the anti-PD-L1 antibody in parallel with the Staphylococcus enterotoxin E (SEE) so that both will incubate with the B cells for 30–60 min.
ii. Keep the T-cell tube incubating at 37 °C during this process.
During T-cell and B-cell incubation, coat MatTek plates with 200 µL of 1× Poly-L-Lysine (PLL) (see Recipes) solution for 30–45 min at room temperature or 37 °C in a tissue culture incubator.
Remove PLL from the plate via vacuum before plating cells.
Note: No wash is needed after removing the PLL.
Add 100 µL (105 cells) of treated B cells to a MatTek dish and let them settle for 5 min at room temperature.
Note: This step does not have to be done in a sterile hood.
Add 100 µL (105 cells) of T cells on top of the previously plated B cells.
Note: Ensure that T cells are added dropwise from the air, gently and evenly over the plate, to help maximize cell dispersion and minimize air bubble formation.
Let the combined solution settle for at least 10 min at room temperature before imaging.
Store microcentrifuge tubes with remaining treated B and T cells in a mini block heater at 37 °C, covered with tin foil to prevent photobleaching.
Imaging of T and B cells synapse formation
Note: Before proceeding with this protocol portion, ensure that the microscope is turned on and calibrated and that Zen Blue software is functional.
Load MatTek plate with cell solution (200 µL total) onto the microscope stage and center above 10× objective for initial observations of cell settling in the plate.
Note: Be gentle with the transfer and ensure that the cell solution is not disturbed from the center of the plate.
Locate the area of interest on the MatTek plate using the 10× objective and eyepiece to ensure that cells have fully settled in.
Remove the MatTek plate from the microscope stage and change to 63× objective with a small amount of oil on the lens.
Note: To prevent damage to the lens, ensure it is at the lowest Z-level before reloading the MatTek plate.
Adjust the objective height to ensure cells are in focus.
Note: The oil must touch the plate and the objective lens for clear images (Figure 2).
Figure 2. Oil is applied to the 63× objective lens, and contact is formed with the MatTek plate. Areas of interest are highlighted using white circles. A) The microscope objective is at the lowest Z-position, with nothing placed on the lens. B) The objective is at the lowest Z-position with a small amount of Immersol autofluorescence immersion oil applied to the lens. C) The MatTek plate is centered above the lens without contact between the plate, oil, and lens. D) MatTek plate centered above the lens with contact between the plate, oil, and lens formed by adjusting using the focus knobs.
Select wavelengths in the fluorescence setting to be imaged.
Adjust gain, intensity, and wavelength absorbance until the background is sufficiently low, but PD-1 and PD-L1 fluorescent signals can be imaged.
Press Capture to take an image for further analysis when settings are acceptable.
The colocalization of both fluorescent signals will evidence synapse formation in a “straight line” between B and T cells (Figure 3).
The number of synapses formed for each antibody tested can be used to visually analyze synapse abundance and quality.
Count the number of synapses formed per X of pairs of T and B cells in contact with one another and compare the number of synapses counted between antibody treatment groups and the control group.
Note: Synapses should be counted only when GFP-PD-1 is enriched in the contact area.
Figure 3. Conjugates formation between Jurkat T cells and Raji B cells. Jurkat T cells expressing GFP-PD-1 were cocultured with Raji B cells expressing mCherry-PD-L1 and pulsed with SEE. The cells on the left panel (A) were imaged after 20 min of coculturing, while the cells on the right (B) were imaged after treatment with anti-PD-1 antibodies. Without anti-PD-1 antibodies, GFP-PD-1 is enriched in the synapse between the Raji and the Jurkat cells (yellow arrows). Scale bar in 20 μm.
Quantification of synapse intensity using microscopic colocalization analysis
Note: A proper version of Zen Blue is recommended for this step. Other software, such as AIM or Image J, can also be utilized, but these steps are outlined explicitly for Zen Blue. For best results, use a .czi file. See Figure 4.
Figure 4. Colocalization analysis using ZEN Blue software. Areas of note are highlighted in white (colocalization tab, threshold adjustment bars, quadrant numbers, region of interest indicator, colocalization coefficients, region of interest tool, Costes auto threshold button, table option). For the example image, the colocalization coefficient to be recorded is 0.725, and the weighted colocalization coefficient to be recorded is 0.767, indicating that 72.5% of GFP-PD-1 signal colocalizes with mCherry-PD-L2 without taking into account pixel intensity, and 76% of GFP-PD-1 signal colocalizes with mCherry-PD-L2 when taking into account pixel intensity. A) Click on the colocalization option on the sidebar. B) Select the region of interest (ROI) using the shape tools. C) Appropriately position the ROI over the synapse and ensure it remains selected. D1) Either manually adjust the bars on the cytofluorogram, or D2) Click the Costes button to auto-set the threshold bars with the selected region of interest using the mouse. E) Click the table option to view the values for the localization coefficients. F) Unweighted and weighted colocalization coefficient values will be listed from left to right (Colocalization Coefficient 1, Colocalization Coefficient 2, W Colocalization Coefficient 1, W Colocalization Coefficient 2). G) Colocalization coefficient values for the region of interest (synapse) will be displayed in the table.
After capturing the images, restart Zen Blue, select image processing, and open the images to be quantified in the image processing suite.
Select colocalization on the sidebar.
Markdown channels associated with PD-1 and PD-L1 fluorescence in the colocalization tab (e.g., GFP and mCherry).
Note: Channels will usually be automatically labeled according to how they were captured during the image acquisition step.
Utilize the region of interest (ROI) tool to mark areas where contact between T and B cells is observed.
Notes:
ROIs should not be limited to GFP-PD-1-enriched contact areas, especially in antibody treatment groups.
This step can also be excluded if a whole image analysis is desired, in which case the rest of the protocol can be conducted similarly without an ROI.
To ensure reproducibility, utilize the Costes auto threshold option by clicking the Costes button. Alternatively, if desired, select the crosshair option to set proper quadrants for use in colocalization analysis. Quadrants are labeled on the cytofluorogram.
Quadrant 4 will contain pixels with low-intensity levels for both channels.
Quadrants 1 and 2 will have high-intensity pixels in one of the channels.
Quadrant 3 will contain pixels with high intensity in both channels.
Cytofluorogram threshold bar placements can be determined using single fluorescence control samples to determine the optimal boundaries to eliminate background signals.
Check the table option to view values for colocalization coefficients and compare between groups with higher colocalization coefficients indicative of more pixels with signals from both channels.
The unweighted colocalization coefficient values (labeled Colocalization Coefficient 1 and Colocalization Coefficient 2) for the region of interest can be utilized to analyze colocalization. Values for weighted colocalization coefficients will range from 0 to 1, with 0 indicating no colocalization and 1 indicating complete colocalization.
i. Unweighted colocalization coefficients are derived from Mander’s coefficients as described in Manders et al. [22].
ii. Weighted colocalization coefficient values (labeled as W Colocalization Coefficient 1 and W Colocalization Coefficient 2 on Zen Blue) consider pixel intensity in their calculation but otherwise use the same methodology as unweighted colocalization coefficient values.
iii. Either coefficient can be utilized at one’s discretion.
1) Colocalization Coefficient 1 (or W Colocalization Coefficient 1) displays the percentage of the cytofluorogram horizontal axis channel that overlaps with the vertical axis channel.
2) Colocalization Coefficient 2 (or W Colocalization Coefficient 2) displays the percentage of the cytofluorogram vertical axis channel that overlaps with the horizontal axis channel.
3) Record the colocalization coefficient (or W Colocalization Coefficient) that correlates with the channel for PD-1.
a) If PD-1 is the horizontal axis channel, use Colocalization Coefficient 1 (or W Colocalization Coefficient 1).
b) If PD-1 is the vertical axis channel, use Colocalization Coefficient 2 (or W Colocalization Coefficient 2)
Higher colocalization coefficient values mean increased colocalization of PD-1 and PD-L1 in the synapse, which indicates poorer antibody efficiency due to the lessened exclusion of PD-1 from the synapse.
Note: Colocalization coefficients can be plotted in GraphPad Prism to analyze quantified synapse intensity values between antibody treatment groups.
Multiple images for each antibody are used to ensure the colocalization coefficient derived is accurate.
Alternative ImageJ quantification of synapse intensity using microscopic colocalization analysis
Note: The JACoP plugin can be utilized as an alternative method for quantifying colocalization. Image processing can be optimally done using a Czi or TIFF file. If not already integrated, the JACoP plugin must be downloaded to the plugin folder in ImageJ. A region of interest cannot easily be specified using the JACoP plugin, so specific synapses must be preemptively isolated for analysis before using ImageJ. Please see the ImageJ website for more details regarding the usage of JACoP.
Following capturing images, open the image to be quantified using ImageJ.
Select Hyperstack for Stack viewing.
Select Default for Color options and check the Autoscale box.
Select Split channels to split into separate windows.
Note: This will separate the PD-L1 and PD-1 fluorescent channels, allowing the JACoP plugin to work correctly.
Select the Plugins tab and click JACoP.
On the JACoP menu, select M1 and M2 coefficients, which will give Mander’s coefficients as calculated in Manders et al. [22].
M1 will give the percentage of channel 1 that overlaps with channel 2.
M2 will give the percentage of channel 2 that overlaps with channel 1.
Record the Mander’s coefficient that correlates with the channel for PD-1.
For example, if PD-1 is labeled as channel 1, look at M1.
Click the threshold tab and adjust thresholds for each channel.
The area utilized for the colocalization analysis will appear in red when the threshold is adjusted.
Once thresholding is satisfactory, click analyze.
Results for M1 and M2 will appear in a separate window.
M1 and M2 can be interpreted in the same manner as described for Zen colocalization analysis.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Hao et al. [1]. Exclusion of PD-1 from the immune synapse: A novel strategy to modulate T cell function. Molecular Therapy: Oncology (Figures 1–3, S1).
General notes and troubleshooting
This protocol outlines the process for Zen Blue software analysis of confocal microscopy images, which is commonly utilized in conjunction with the Zeiss confocal microscope. Image analysis can also be conducted in other programs, such as ImageJ (the most recent version is preferred), but the steps taken to quantify colocalization will differ. The rest of the steps involving plating and imaging remain the same. Confocal microscopes, such as the Leica SP8, can also be utilized. In this case, the analysis would need to be done using the JACoP plugin for ImageJ.
The amount of SEE added to B cells for incubation can be increased as needed to help form immune synapses. SEE concentration can vary, with the minimum recommended volume added being 0.5 µL. It was found that 5 µL of SEE was necessary to optimize this protocol, but the amount can be altered as needed.
Utilizing the CD3/CD28/CD2 ImmunoCult T-cell activator does not harm imaging quality when using a cell line such as PD-1-expressing Jurkat T cells to enhance results further. Still, it is unnecessary, as adding the activator only sometimes leads to significant improvements in synapse formation and image.
Optimal synapse formation begins after cells settle for at least 10 min. Synapses are mostly readily noted by a bright line of PD-1 GFP signal in the contact area between Raji and Jurkat cells.
Following the 10-min settling time, there is an approximately 40–50 min window for capturing images. Synapses will begin to form after this settling time, but the optimal imaging window for synapses will be 20–60 min after coculture. Cells have been observed to dissociate 1 h after being cocultured together (Figure 5).
Figure 5. Image capture timeline and synapse formation. Synapses can be observed to begin forming at 10 min after coculture. The optimal image window for more developed synapses is 20–60 min after coculture. Cells are observed to dissociate at times greater than 60 min after coculture or greater than 50 min after initial synapse formation.
Acknowledgments
Grants from the NIH AI125640, AI150597, and AI175498 supported this work.
Competing interests
The authors report no competing interests.
Ethical considerations
No animal or human subjects were used in this protocol.
References
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Promega. (2019). PD-1/PD-L1 blockade bioassay. https://www.promega.com/products/reporter-bioassays/immune-checkpoint-bioassays/pd1_pdl1-blockade-bioassays/?catNum=J1250
Naito, Y., Osa, A., Masuhiro, K., Hirai, T., Koyama, S. and Kumanogoh, A. (2020). Monitoring PD-1-Blocking Antibodies Bound to T Cells Derived from a Drop of Peripheral Blood. J Visualized Exp. doi.org/10.3791/60608.
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Alarcón, B., Mestre, D. and Martínez-Martín, N. (2011). The immunological synapse: a cause or consequence of T-cell receptor triggering? Immunology. 133(4): 420–425.
Zinselmeyer, B. H., Heydari, S., Sacristán, C., Nayak, D., Cammer, M., Herz, J., Cheng, X., Davis, S. J., Dustin, M. L., McGavern, D. B., et al. (2013). PD-1 promotes immune exhaustion by inducing antiviral T cell motility paralysis. J Exp Med. 210(4): 757–774.
Yokosuka, T., Takamatsu, M., Kobayashi-Imanishi, W., Hashimoto-Tane, A., Azuma, M. and Saito, T. (2012). Programmed cell death 1 forms negative costimulatory microclusters that directly inhibit T cell receptor signaling by recruiting phosphatase SHP2. J Exp Med. 209(6): 1201–1217.
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Article Information
Publication history
Received: Apr 12, 2024
Accepted: Jul 14, 2024
Available online: Aug 8, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Immunology > Antibody analysis > Antibody-antigen interaction
Cell Biology > Cell imaging > Confocal microscopy
Immunology > Immunotherapy
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Evaluating Mechanisms of Soil Microbiome Suppression of Striga Infection in Sorghum
TT Tamera Taylor
JD Jiregna Daksa
MS Mahdere Z. Shimels
DE Desalegn W. Etalo
BT Benjamin Thiombiano
AW Aimee Walmsey
AC Alexander J. Chen
HB Harro J. Bouwmeester
JR Jos M. Raaijmakers
SB Siobhan M. Brady
DK Dorota Kawa
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5058 Views: 515
Reviewed by: Xiaofei LiangSteven RunoKerui Huang
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Original Research Article:
The authors used this protocol in Cell Reports Apr 2024
Abstract
The root parasitic weed Striga hermonthica has a devastating effect on sorghum and other cereal crops in Sub-Saharan Africa. Available Striga management strategies are rarely sufficient or not widely accessible or affordable. Identification of soil- or plant-associated microorganisms that interfere in the Striga infection cycle holds potential for development of complementary biological control measures. Such inoculants should be preferably based on microbes native to the regions of their application. We developed a method to assess microbiome-based soil suppressiveness to Striga with a minimal amount of field-collected soil. We previously used this method to identify the mechanisms of microbe-mediated suppression of Striga infection and to test individual microbial strains. Here, we present protocols to assess the functional potential of the soil microbiome and individual bacterial taxa that adversely affect Striga parasitism in sorghum via three major known suppression mechanisms. These methods can be further extended to other Striga hosts and other root parasitic weeds.
Key features
• This protocol provides a detailed description of the methods used in Kawa et al. [1].
• This protocol is optimized to assess soil suppressiveness to Striga infection by using natural field-collected soil and the same soil sterilized by gamma-radiation.
• This protocol is optimized to test bacterial (and not fungal) isolates.
• This protocol can be easily extended to other host–parasite–microbiome systems.
Keywords: Striga hermonthica Suppressive soils Microbiome Sorghum Aerenchyma Suberin Haustorium-inducing factors
Background
Sorghum (Sorghum bicolor) is among the five most important crops in the world and a staple food and forage cereal in Sub-Saharan Africa [2]. Sorghum is grown in diverse agroecological zones, predominantly by small-hold farmers [3]. Most sorghum cultivars are suited to arid, nutrient-depleted soils; yet, sorghum productivity is challenged by pathogens. The parasitic weed Striga hermonthica causes substantial sorghum yield losses, affecting approximately 60% of Sub-Saharan African farmlands [4].
Striga is a root parasite that infects cereal crop species, including sorghum, rice, and pearl millet [5]. Striga’s life cycle is dependent on specific compounds exuded from the host roots into the surrounding soil. Striga germinates upon the perception of strigolactones, while haustorium-inducing factors (HIFs) initiate the development of a haustorium, a specialized organ that allows Striga to attach to and penetrate the root tissue [6–8]. Upon reaching the host vasculature, Striga connects its own xylem vessels with its host vasculature [7], enabling the parasite to withdraw water and nutrients from its host, thereby compromising host fitness and productivity [9].
Striga management includes manual weed removal, chemical control methods, and breeding for host resistance [10]. Chemical application poses a challenge in rain-fed agricultural systems, while commercial sorghum varieties show only partial resistance and are not always accessible or suited to the agricultural practices of small-holder farmers [11]. Recently, the functional potential of the soil microbiome to suppress Striga infection has been described. These include microbial isolates that are pathogenic to Striga [12] and those that degrade HIFs or that induce physical barriers to Striga parasitism in host roots [1]. Further development of microbial-based agricultural solutions against Striga will require screening candidate microbes native to soils in Striga-infested regions.
Here, we present a set of protocols to assess the contribution of soil microbiome to Striga infection levels and individual soil-borne bacterial isolates to Striga suppression-associated mechanisms. The soil plug assay enables testing Striga suppressiveness of multiple soils collected from agricultural fields. Striga resistance modes are typically described as pre-attachment resistance [reduced germination and (pre)haustorium formation], and post-attachment resistance (when parasite fails to penetrate the root tissue and/or establish the vascular connection with the host). To distinguish which stage of Striga infection the soil microbiome affects, we recommend extracting and testing host root exudates in an in vitro Striga germination assay and a haustorium formation assay. Preparation of host root cross-sections and their histological staining allows quantification of aerenchyma and suberization, both associated with microbe-mediated suppression of post-attachment stages of Striga infection [1]. To test which isolates affect haustorium formation and induce changes in host root cellular anatomy, we present an in vitro haustorium induction assay and a method of sorghum inoculation in sand, respectively. The presented protocols can be easily adapted and applied to multiple sorghum cultivars and other Striga hosts.
Materials and reagents
Biological materials
Sorghum bicolor seeds. We recommend using Striga-susceptible sorghum varieties, e.g., Shanqui Red, and the Striga-resistant cultivar SRN39 as positive and negative controls for Striga germination, respectively [13,14] [Shanqui Red: PI 656025, SRN39: PI 656027, available from Germplasm Resource Information Network (GRIN-Global)]
Striga hermonthica seeds (note that working with Striga in certain locations/countries requires permits (e.g., Federal Noxious Weed Permit issued by APHIS in US) and must be performed in contained-type facilities (e.g., https://crf.ucdavis.edu)
Field-collected soil of interest
Bacterial isolates of interest
Reagents
Commercial household bleach [Clorox containing 8.25% (v/v) sodium hypochlorite (NaOCl)]
Captan 50% wettable powder (active ingredient: N-Trichloromethylthio-4-cyclohexene-1,2-dicarboximide, 48.9%) (Arysta Life Science, CAS number: 133-06-2)
Tween-20 (Sigma-Aldrich, CAS number: 9005-64-5)
Calcium nitrate (Sigma-Aldrich, CAS number: 13477-34-4)
Potassium nitrate (Spectrum, CAS number: 7757-79-1)
Potassium phosphate (Sigma, CAS number: 7778-77-0)
Magnesium sulphate (Research Products International, CAS number: 7487-88-9)
EDTA (Ethylenediaminetetraacetic acid) ferric salt (Sigma, CAS number: 18154-32-0)
Boric acid (Fisher Chemical, CAS number: 10043-35-3)
Manganese chloride (J.T. Baker, CAS number: 13446-34-9)
Zinc sulphate (Sigma-Aldrich, CAS number: 7446-20-0)
Copper sulphate (Spectrum, CAS number: 7758-99-8)
Sodium molybdate (Sigma-Aldrich, CAS number: 10102-40-6)
Sodium chloride (Fisher Chemical, CAS number: 7647-14-5)
BactoTM yeast extract (Thermo Fisher, catalog number: 212750)
N-acetylglucosamine (Sigma-Aldrich, CAS number: 7512-17-6)
Tryptic soy broth (TSB), Bacto soybean-casein digest medium (Difco, catalog number: 211825)
Bactoagar (agar bacteriological) (Difco, catalog number: 214530)
Agarose (VWR, CAS number: 9012-36-6)
GR-24rac (StrigoLab, CAS number: 76974-79-3)
DMBQ (2,6-methoxy-1,4-benzoquinone), 97% (Sigma-Aldrich, CAS number: 530-55-2)
Fluorol yellow 088 (Santa Cruz Biotech., CAS number: 81-37-8)
Aniline blue (Fisher Chemical, CAS number: 28631-66-5)
Toluidine blue O (J.T. Baker, CAS number: 92-31-9)
Lactic acid (Acros Organics, catalog number: 189870010, CAS number: 79-33-4)
Syringic acid, 95% (Sigma-Aldrich, CAS number: 530-57-4)
Vanillic acid, 98% (Alfa Aesar, CAS number: 121-34-6)
Formalin (37% formaldehyde) (Fisher Chemical, CAS number: 50-00-0)
95% ethanol (Koptec, CAS number: 64-17-5)
Glacial acetic acid (Fisher Chemical, CAS number: 64-19-7)
Glycerol (Fisher Chemical, CAS number: 56-81-5)
DMSO (dimethyl sulfoxide) (Sigma-Aldrich, CAS number: 67-68-5)
Sodium chloride (Fisher Chemical, CAS number: 7647-14-5)
Methanol (Sigma-Aldrich, CAS number: 67-56-1)
Solutions
Half-strength modified Hoagland media (see Recipes)
Bacterial growth media (see Recipes)
Formalin-Aceto-Alcohol (FAA) solution (see Recipes)
Recipes
Half-strength modified Hoagland media
Stock solution Reagent
g/L stock
Volume (mL) stock per 1 L of final solution
0.5 M calcium nitrate Ca(NO3)2·4H2O 118.08 5
1.0 M potassium nitrate KNO3 101.11 2.5
0.1 M potassium phosphate KH2PO4 13.609 0.5
0.5 M magnesium sulphate MgSO4·7H2O 123.24 2
98.6 mM EDTA ferric salt C10H12FeN2NaO8·3H2O 41.52 1
Micronutrient stock: 1
46.3 mM boric acid H3BO3 2.86
9.1 mM manganese chloride MnCl2·4H2O 1.81
0.77 mM zinc sulphate ZnSO4·7H2O 0.22
0.32 mM copper sulphate CuSo4·5H2O 0.08
0.52 mM sodium molybdate Na2MoO4·2H2O 0.126
Store at 4 °C.
Bacterial growth media
Reagent Final concentration g/L final solution
Sodium chloride 0.5% 5
Potassium dihydrogen phosphate 0.1% 1
BactoTM yeast extract 0.01% 0.1
N-acetylglucosamine 2 mM 0.44
Store at room temperature.
Formalin-Aceto-Alcohol (FAA) solution
Reagent Final concentration mL/L final solution
Formalin (37% formaldehyde) 10% 100
95% ethanol 50% 500
Glacial acetic acid 5% 50
Store at room temperature.
Laboratory supplies
4 mm mesh
50 mL conical tubes
Whatman qualitative filter paper, Grade 1, 90 mm diameter (Sigma-Aldrich)
Sterile plastic Petri plate, 100 mm diameter
Parafilm
Aluminum foil
Cones, Depot tree pots 40 cm (Greenhouse Megastore)
Gauze pads
Rubber bands
Ethanol 200 proof (KOPTEC, CAS number: 64-17-5)
High-purity filtered sand, effective size 0.45–0.55 mm (Covia)
Forceps
Pycnometers, 50 mL (KLM BioScientific, Borosilicate 3.3 Glass)
Transparent tray
Whatman glass microfiber filters, grade GF/A, 13 mm diameter discs (Sigma-Aldrich)
Equipment
Laminar flow cabinet (Labconco, Purifier Biological Safety Cabinet, model: 3440009 LS)
Hot stirrer plate (VWR, model: VMS-C7)
Stereomicroscope (Nikon, model: SMZ 1500)
Balance (Sartorius, Explorer Pro, model: E0114)
Orbital shaker (GeneMate, model: Orbital Shaker Variable)
Centrifuge for 50 mL tubes (Eppendorf, model: 5810 R)
Vibratome (Leica, model: VT1000 S)
Fluorescent confocal microscope (Carl Zeiss, model: LSM 700)
Vacuum chamber (SP Bel-Art)
Microbiological incubator shaker (Innova, model: 4400 Incubator Shaker)
Autoclave (Tuttnauer, model 5596SP-1V)
Spectrophotometer (Eppendorf, model: BioPhotometer plus)
Software and datasets
ImageJ version 1.53g (https://imagej.net/ij/download.html) with Cell Counter plug-in
Procedure
Sorghum seed surface sterilization and germination
Note: Always try to use sorghum seeds that are similar in size.
Place seeds in a 50 mL conical tube.
Add up to 50 mL of freshly prepared sterilizing solution (30% commercial bleach, 0.2% Tween-20, v/v).
Gently agitate seeds on an orbital shaker in sterilizing solution for 20 min.
Discard the sterilizing solution and seeds that float on the surface of the solution.
Wash seeds with sterile water for 2 min, five times.
Optional: If fungal contamination occurs, agitate seeds overnight in 5% (w/v) Captan slurry followed by five washes in sterile water.
Place seeds on sterile Petri dishes containing two Whatman filter papers moistened with 5 mL of sterile water.
Seal plates with parafilm.
Incubate plates in the dark at 30 °C for the duration indicated per experiment.
Striga seed sterilization and preconditioning
Estimate the density of Striga seeds.
Weigh out a small number of dry seeds on a Whatman filter paper in a Petri dish.
Record mass.
Count the total number of seeds using a stereomicroscope.
Calculate the number of seeds per milligram.
Repeat steps B1a–d at least three times and calculate an average.
Note: We recommend conducting this once for each new batch of Striga seeds.
Wash Striga seeds in 10% (v/v) bleach and 0.02% (v/v) Tween-20 for 10 min.
Discard the solution and wash the seeds five times in sterile water.
Precondition Striga seeds by incubation at 30 °C in the dark. The chemicals and protocols needed for preconditioning are dependent on the experiment and are outlined in their respective sections.
Optional: The number of Striga seeds for each experiment is indicated as the number of germinable seeds. For each newly collected Striga seed batch, germinability should be assessed, as it is highly variable.
Calculate Striga germinability:
Sterilize a specific amount (number or weight) of Striga seeds as determined in step B1.
Disperse seeds onto a Petri dish with two sterilized Whatman filter papers moistened with 5 mL of sterile water.
Precondition at 30 °C in the dark for 10–14 days.
Transfer seeds to a new Petri dish with two sterile Whatman filter papers moistened with 5 mL of 1 ppm GR24rac.
Seal with parafilm and return to 30 °C in a dark incubator.
After three days, count the total number of seeds and number of germinated seeds in the viewing area using a stereomicroscope.
Calculate the germination rate for each Petri dish.
GR% = (Ngs/Nts) × 100
Ngs: Total number of germinated seeds per Petri dish
Nts: Total number of seeds per Petri dish
Use these estimates to treat each experiment with the desired amount of germinable Striga seeds using the following formula:
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Calculate the total amount of sand needed (you can use 1/10 of required sand amount for preconditioning and mix with the remaining 9/10 on the day of planting).
Mix sand with sterile water to reach approximately 16% (v/w) moisture level.
Add Striga seeds to wet sand and mix thoroughly.
Incubate sand–Striga mix at 30 °C in the dark for 10–14 days.
Prepare the sand used as a negative control in the same manner, without adding the Striga seeds.
Soil plug assay for soil suppressiveness
C1. Soil sterilization
Air-dry the soil at room temperature for 4–7 days.
Sieve the soil through a 4 mm mesh.
Sterilize the soil by gamma irradiation with an 8 kGy dose.
Notes:
To attribute the Striga suppressive effect solely to its microbiome, it should be ensured that the physicochemical properties of gamma-sterilized and natural (non-sterilized) soil remain comparable.
Assess bacterial and fungal diversity to ensure which is depleted.
C2. Soil plug preparation
Mix each soil batch (sterilized and non-sterilized) with sterile water to reach 5% of moisture level (w/v).
Cut a hole at the bottom of a 50 mL conical tube to provide drainage of excess liquid.
Wrap the tubes in aluminum foil or other comparable material to block light. Do not cover the hole at the bottom.
Fill the tubes with soil.
With a 1 mL pipette tip, make a hole in the soil for a germinated sorghum seedling.
Using forceps, gently place a 3-day-old sorghum seedling in the soil.
Place sorghum in the greenhouse or grow in a chamber at 28 °C during the day (11 h) and 25 °C at night (13 h), with a light intensity of 450 μmol/m2/s and 70% relative humidity.
Apply 3 mL of sterile deionized water to each soil plug every second day.
Grow sorghum in the soil plug for 10 days.
C3. Cone preparation
Sterilize 3,000 germinable Striga seeds per plant as determined in step B5.
Autoclave the cones prior to planting.
Place a gauze pad to cover the hole at the bottom of each cone and secure it with a rubber band.
Fill each cone with 350 mL of fresh (not preconditioned) sand (Figure 1A).
Top up with 350 mL of preconditioned sand without (control) or with Striga seeds.
Within the 50 mL conical cone, make a hole to accommodate the soil plug.
Transfer the 10-day-old sorghum seedling together with the soil plug to the cone (Figure 1A).
Cover the soil plug surface with a thin layer of fresh sand.
Place sorghum in the greenhouse or growth chamber at 28 °C during the day (11 h) and 25 °C at night (13 h) with a light intensity of 450 μmol/m2/s and 70% relative humidity.
Apply 50 mL of sterile half-strength modified Hoagland solution to each plant on days 0, 7, and 14. Apply 50 mL of sterile deionized water to each plant on days 1, 4, 10, 13, and 17.
Quantify Striga infection 14 and/or 21 days after transfer to cones.
Figure 1. Growth setup to assess soil suppressiveness to Striga in soil plug cone assay (A) and screening for bacterial isolates associated with Striga-suppressive phenotypes (B)
Quantification of Striga infection
Gently remove the plant from the cone.
Shake the plant to remove as much of the sand and soil from the roots as possible. Collect all the soil and sand.
Gently swirl the roots in a tray filled with water to wash off the remaining substrate. Do not rub the roots, since Striga might get de-attached from the host.
Gently dry the roots with a paper towel.
Cut the root system from the shoot.
Record the root’s fresh weight.
Place the root system in a transparent tray filled with water.
Use a stereomicroscope with 10× magnification to examine the roots for sites of Striga attachment and penetration. Use forceps to spread the roots and systematically look for the presence of Striga. Refer to Figures 2D and 2E as to how to distinguish attached from penetrated Striga.
Screen the sand and soil collected in step D2 for any attached Striga that may have fallen off the root, called “de-attached” Striga (see Figure 2C for the developed, penetrated Striga that de-attached from sorghum root).
Sum the number of penetrated Striga on the roots and the number of Striga rescued from the sand (“de-attached Striga”) to obtain the total penetration number.
Sum the number of total penetrated Striga and attached Striga to obtain the total Striga count.
The total Striga count and the total number of attachments should be normalized by the fresh weight of the root scored in step D6.
Figure 2. Striga developmental stages of interest. (A) Striga germination and (B) haustorium development of in vitro experiments. (C) Striga may break off at the haustorial connection during harvest even when well-developed. (D) Both attachment and penetrated Striga have developed haustoria that are connected to the sorghum root, but the penetrated Striga shows a further development of leaf lobes. Yellow arrowheads: developing leaf lobes in young, penetrated Striga. (E) Older penetrated Striga may begin developing green leaf tissue.
In vitro Striga germination and haustorium formation assay
E1. Root exudate collection
Note: Exudates are collected from a sorghum plant grown in sterilized soil and a plant grown in non-sterilized soil.
Place a 1 L beaker under the cone.
Pour 300 mL of sterile water into the cone.
Collect 100 mL of the flowthrough and store refrigerated until use.
E2. In vitro Striga germination and haustorium formation assay
Surface sterilize the required amount of Striga seeds as described in section B.
Place a sterile glass fiber filter paper in a 94 mm diameter polystyrene Petri dish and add 3 mL of sterile deionized water.
Spread the sterilized Striga seeds on the Petri dishes with wet filter paper. Use approximately 200 seeds per plate.
Seal the Petri dishes with parafilm.
Precondition Striga seeds for 6–8 days at 30 °C in the dark.
After preconditioning, dry Striga seeds in the Petri dish in the laminar flow hood.
Transfer approximately 30 Striga seeds per well to a 6-well plate.
Add 50–200 times diluted root exudates (as needed to get approximately 40%–50% of Striga seeds to germinate) to the Striga seeds in three technical replicates (exudate collected from an individual plant divided into triplicates) for the germination bioassay. For the haustorium formation bioassay, use undiluted root exudate in triplicates (in the case of the undiluted root exudate, 1 ppm GR24rac is added first to induce uniform germination).
Use 1 ppm GR24rac and 100 mM DMBQ in three technical replicates as a positive control for the Striga seed germination and haustorium formation assays, respectively.
Incubate the 6-well plate in the dark at 30 °C for two days.
Count the number of germinated seeds, seeds that developed haustorium, and the total number of seeds per well. Refer to Figure 2A and 2B for examples of these stages.
Calculate the Striga germination rate (GR):
GR% = (Ngs/Nts) × 100
Ngs: Number of germinated seeds per well
Nts: Total number of seeds per well
Calculate the haustorium formation rate (HFR):
HFR% = (NHs/Ngs) × 100
NHs: Total number of haustorium per well
Ngs: Number of germinated seeds per well
Striga germination induced by 1 ppm GR24rac is highly dependent on the Striga seed batch, and this germination rate can vary seasonally. In our experiments, 1 ppm GR24rac induces, on average, germination of 60% of Striga seeds.
In vitro assay to test the potential of individual bacterial strains to reduce Striga haustorium formation
F1. Bacterial inoculum preparation
Streak the bacterial strain from the glycerol stock onto a plate with a 1/10 strength TSB agar media (1.5% (w/v) agar).
Incubate plates at 26 °C for 24–48 h in the dark.
Prepare liquid bacterial growth media by dissolving the ingredients in Recipe 2 into deionized water.
Autoclave the liquid bacterial growth media for 30 min at 25 °C.
F2. Striga haustorium formation assay
Surface-sterilize the required amount of Striga seeds as described in steps B1–B4.
Place two sterile glass fiber filter papers in a 60 mm Petri dish and add 3 mL of sterile deionized water.
Place four 13 mm sterile glass fiber filter papers in each Petri dish.
Add the desired volume of sterile deionized water to surface-sterilized Striga seeds to bring the number of Striga seeds to between 80 and 100 per disc.
Pipette 100 µL of the Striga–water mix prepared in section E2 to each 13 mm disc.
Seal the Petri dishes with parafilm.
Incubate the plates with the Striga seeds for 11 days at 30 °C in the dark.
At day 12, add 100 µL of water containing a final concentration of 1 µM GR24rac in DMSO to each disc.
Seal the Petri dishes with parafilm and incubate at 28 °C for 48 h.
At day 12, inoculate sterile liquid bacterial growth media with a single colony of the isolate of interest (as described in section F1) and incubate for 24 h at 25 °C with shaking at 200 rpm.
Measure OD600 using a spectrophotometer.
Centrifuge the liquid culture at 20,000 rcf for 5 min.
Wash the collected cells twice with 10 mL of sterile 0.9% NaCl.
Adjust the OD600 to 0.15 by resuspending the cells in 0.9% NaCl.
Transfer 18 μL of the bacterial suspension to 332 μL of the bacterial growth media supplemented with HIFs: 100 µM syringic or 50 µM vanillic acid, dissolved in methanol. Prepare a negative control with the same volume of methanol as used to dissolve the HIFs.
Incubate the culture for another 24 h at 25 °C with shaking at 200 rpm.
Add 50 μL of the cell-free culture filtrate to pre-germinated Striga seeds.
Seal the plates with parafilm and incubate them at 25 °C for 48 h in the dark.
Take pictures of each disc (filter paper with haustoria) under a dissecting microscope.
F3. Quantification of haustorium formation rate
Open ImageJ and load the disc (filter paper with haustoria) image file.
Go to Plugins and click on Cell Counter.
Click on Initialize.
Select Type 1 and add a mark on each non-germinated Striga seed.
Select Type 2 and add a mark on each germinated Striga seed.
Select Type 3 and add a mark on each Striga seed that developed haustorium.
Click on Save Markers.
Note the counts in the Excel file.
Calculate the haustorium formation rate (HFR) as described in section E2.
Note: The Striga haustorium formation rate should be approximately 70% for 100 µM syringic acid and 30% for 50 µM vanillic acid.
Assay to test the potential of individual bacterial strains to induce root cellular phenotypes associated with Striga suppression
Note: Each plant is inoculated with 107 CFU/g sand.
G1. Bacterial inoculum preparation
Streak the bacterial strain from the glycerol stock onto a plate with a 1/10th strength TSB agar media (1.5% w/v agar).
Incubate plates at 26 °C for 24–48 h in the dark.
Inoculate 5 mL of liquid 1/10th TSB with a single colony.
Incubate liquid cultures for 24–48 h at 26 °C with shaking (200 rpm). Culture time depends on the number of cells required.
Calculate the number of cells required per the following formula:
Number of cells required = 107 (CFU/g soil) × 50 g sand × number of plants to test
Calculate the required volume of the inoculum per the following formula:
Volume of inoculum required = number of cells required/(CFU/mL of bacterial culture)
Harvest bacteria by centrifugation of the required inoculum volume at 7,000 rcf for 20 min.
Resuspend bacterial pellet in sterile half-strength modified Hoagland solution (5 mL per plant).
G2. Sorghum inoculation
Cut a hole at the bottom of the 50 mL conical tube to provide drainage of excess liquid.
Wrap the tubes in aluminum foil. Do not cover the hole at the bottom.
Fill the tubes with sand (approximately 50 g of sand per tube)
Apply 5 mL of sterile half-strength modified Hoagland solution to each tube.
With a 1 mL pipette tip, make a hole for the sorghum seedling.
Pick up a 2-day-old sorghum seedling and, while transferring it to the sand, apply 5 mL of bacterial inoculum (step 8 in section G1) directly onto the sorghum root.
Bury the sorghum root and seed with sand.
Place the tube with sorghum in the greenhouse or growth chamber at 28 °C during the day (11 h) and 25 °C at night (13 h) with a light intensity of 450 μmol/m2/s and 70% relative humidity.
Apply 5 mL of sterile water every second day.
Quantification of root cellular anatomy
H1. Root harvesting
Gently remove a plant from the tube or a cone.
Tap the root to remove the attached sand and soil.
Wash the root system with water.
Cut a 1–1.5 cm segment of the root from the part of interest within the root system.
Note: We recommend harvesting fragments from the tip and middle section of the crown and seminal root from plants grown in the cone system. For plants grown in 50 mL tubes, harvest individual 1 cm fragments starting 3 cm from the primary root tip in the shootward direction. Avoid root regions that are wavy or damaged. Remove lateral roots from your fragments of interest.
H2. Root tissue embedding
Prepare a 5% agarose gel (m/v with water). We recommend melting the agarose in the autoclave. Keep the agarose stirring on a hot plate to prevent it from solidifying.
Prepare glass vials, each filled with 10 mL of FAA solution.
Pour 5% of the liquid agarose into the Eppendorf tube (Figure 3A, B).
Using forceps, insert the root into the Eppendorf tube containing agarose. Aim to place the root as straight as possible (Figure 3C).
Let the agarose solidify.
Cut off the bottom of the Eppendorf tube with a hot scalpel (Figure 3D).
Open the Eppendorf tube and, with forceps, push the agar “plug” through the hole at the bottom (Figure 3E).
Transfer the agar plug to the glass vial containing FAA (make sure the agar plug is completely covered with solution).
H3. Root tissue fixation and rehydration
Place the open glass vial with the submerged agar plug in a vacuum chamber and vacuum infiltrate for 10 min.
Remove the vial from the vacuum chamber, close it, and leave the agar plugs in the FAA solution overnight at room temperature.
Remove FAA from the glass vials.
Add 70% ethanol to the vial (make sure the agar plug is covered in solution) and incubate for 30 min.
Replace 70% ethanol with 50% ethanol and incubate for 30 min.
Replace 50% ethanol with 30% ethanol and incubate for 30 min.
Replace 30% ethanol with 10% ethanol and incubate for 30 min.
Replace 10% ethanol with sterile deionized water.
Store agar plugs in 4 °C.
H4. Root tissue sectioning
Cut a 0.5–0.75 cm fragment from the embedded sample with the scalpel, making sure the cut is perpendicular to the root. The root needs to be cut perpendicularly so that it will be 100% vertical when mounted on the specimen disc (Figure 3).
Figure 3. Root embedding process. (A, B) Melted agarose is poured into an Eppendorf tube; (C) then, a section of root is placed into the agarose while still warm. (D) After the agarose solidifies, the bottom is cut off with a hot scalpel, and (E) the agarose plug is pushed out. (F) This plug is cut down further to obtain a straight, perpendicular root portion that is suitable for sectioning on a vibratome.
Mount the agar plug on the specimen disc (provided with the vibratome) with glue (we recommend using Cyanoacrylate glue).
Fill the vibratome buffer tray with ice and pour water over it.
Attach the specimen disk with the sample to the buffer tray.
Place the knife securely into the knife holder and attach it to the vibratome. We recommend wearing cut-proof gloves during this step.
Set the required knife amplitude and speed. We recommend starting with amplitude 8 and speed 8 and adjusting it depending on the tissue structure.
Set the section thickness to 300 μm. If your tissue keeps popping out from the agar, increase the section thickness.
Make the first section and discard it.
Make the next section and capture it gently with the fine-tipped paintbrush.
Observe the section under a 10× magnification stereomicroscope. Discard sections that are blurry or damaged.
Place the section in the 12-well plate filled with water and store it on ice.
Prepare 3–4 sections per sample for each staining type.
H5. Suberin quantification
Suberin staining and image acquisition
Replace the water in the 12-well plate with 0.01% Fluorol yellow (in lactic acid, w/v).
Incubate at room temperature for 30 min in the dark with gentle agitation.
Remove the Fluorol yellow solution.
Wash sections for 5 min with deionized water. Repeat three times.
Add 0.5% aniline blue solution (in water, w/v).
Incubate at room temperature for 30 min with gentle agitation.
Wash sections for 10 min with deionized water. Repeat four times.
Mount sections on microscopy slides with 50% glycerol (v/v).
Image sections with a confocal laser scanning microscope with an excitation wavelength of 488 nm. The gain should be adjusted manually. (Image sections where you expect the highest signal at first to set the gain.)
Save file as a raw image file (.czi file in case of Zeiss 700).
Suberin content quantification
Open ImageJ and load root cross-section raw image file.
Use the Freehand selections tool to outline endodermal cells with representative fluorescent signals.
Use the Measure tool under Analyze to record the Mean Gray Value.
Repeat steps H5.2b–c for 3–4 representative cells.
Average the Mean Gray Value for quantified cells to express mean fluorescent signal.
Developmental patterns of suberin quantification
Note: To assess the influence of microbes on the developmental status of suberization, one needs to first assess the distance from the root tip where the patchy and fully suberized zone starts for a given growth condition and genotype.
Open the cross-section image in the image processing software.
Mark the regions where the signal is weaker due to the technical challenges of obtaining sorghum root sections that can be visualized in a single plane. These regions can be recognized by following the changes in the background fluorescent signal from the vasculature. Exclude the regions with less fluorescence in the vasculature and adjacent endodermal cells.
Count the number of suberized cells (excluding the regions marked in the above step) (Figure 4A, B).
Count the total number of endodermal cells.
Calculate the proportion of suberized cells in the endodermis.
Calculate the proportion of plants with a fully suberized endodermis.
H6. Aerenchyma quantification from sections
Note: Fluorol yellow sections can be used in the brightfield microscope to quantify aerenchyma. If using the same sections as for suberin quantification, proceed to step H6.1e.
Image acquisition
Replace the water in the well plate with sections with 0.1% toluidine blue (m/v in deionized water).
Incubate at room temperature for 5 min with gentle agitation.
Remove the toluidine blue solution.
Wash the sections for 1 min with deionized water. Repeat five times.
Image sections with a brightfield microscope and save the image in .tiff format.
Aerenchyma quantification
Open ImageJ and load root cross-section .tiff file.
Use the Freehand selections tool to outline the aerenchyma (Figure 4C).
Use the Measure tool under Analyze to record the area of each aerenchyma lacuna outlined.
Use the Add Noise tool under Process to mark already recorded areas.
Use the Freehand selections tool to outline and record the total root cross-section area (TRA).
Save the Results report as a .csv file with a name that indicates the image analyzed.
Calculate the total aerenchyma area (TAA) as a sum of individual aerenchyma lacunae quantified in step 2c.
Calculate aerenchyma proportion per formula:
Aerenchyma proportion = TAA/TRA
We typically observe aerenchyma formed in 30% of the root area (for crown roots of 4-week-old sorghum grown in a soil-plug system).
Figure 4. Example quantification images. (A) The endodermis may be partially or (B) fully suberized. (C) Aerenchyma lacuna outlines are traced to measure total aerenchyma content. White triangles: not suberized endodermal cells; black triangles: aerenchyma lacuna.
Aerenchyma quantification—porosity assay for plants grown in 50 mL tubes
Fill in pycnometers with water and weigh them (Pw).
Gently remove the plant from the tube.
Gently tap the root system to remove sand.
Wash the root system in water, trying to remove as much sand as possible without squeezing the root.
Very gently dry the root system with a paper towel.
Weigh the root system (R).
Transfer the root system to the pycnometer, refill the pycnometer with water, and weigh (Pr).
Place the open pycnometer with the root system in the vacuum chamber and vacuum infiltrate until the last air bubbles can be seen. A longer time is required for larger root systems. Shake the pycnometers a few times during infiltration to aid in the release of air bubbles.
Weigh the pycnometer with the root and water after vacuum infiltration (Pv).
Calculate the porosity per the following formula:
Root system porosity = (Pv - Pr)/ (Pw + R - Pr)
An average root porosity of 8.4% in mock (sterile media) treatment is typically observed for the entire root system of 10-day-old sorghum plant.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Kawa et al. [1]. The soil microbiome modulates the sorghum root metabolome and cellular traits with a concomitant reduction of Striga infection. Cell Reports. (Figure 1, panel C–E; Figure 3, panel B–G, Figure 4 panel A–H)
Acknowledgments
This research was supported by the Bill and Melinda Gates Foundation (Seattle, WA, USA) via grant OPP1082853 “RSM Systems Biology for Sorghum.” This protocol is used in: Cell Reports (2024), DOI: 10.1016/j.celrep.2024.113971 [1].
Competing interests
There are no conflicts of interest or competing interests.
References
Kawa, D., Thiombiano, B., Shimels, M. Z., Taylor, T., Walmsley, A., Vahldick, H. E., Rybka, D., Leite, M. F., Musa, Z., Bucksch, A., et al. (2024). The soil microbiome modulates the sorghum root metabolome and cellular traits with a concomitant reduction of Striga infection. Cell Rep. 43(4): 113971.
Paterson, A. H., Bowers, J. E., Bruggmann, R., Dubchak, I., Grimwood, J., Gundlach, H., Haberer, G., Hellsten, U., Mitros, T., Poliakov, A., et al. (2009). The Sorghum bicolor genome and the diversification of grasses. Nature. 457(7229): 551–556.
Spallek, T., Mutuku, M. and Shirasu, K. (2013). The genus Striga: a witch profile. Mol Plant Pathol. 14(9): 861–869.
Ejeta, G. and Gressel, J. (2007). Integrating New Technologies for Striga Control. World Scientific. ISBN: 978-981-270-708-6.
Runo, S. and Kuria, E. K. (2018). Habits of a highly successful cereal killer, Striga. PLoS Pathog. 14(1): e1006731.
Cui, S., Wada, S., Tobimatsu, Y., Takeda, Y., Saucet, S. B., Takano, T., Umezawa, T., Shirasu, K. and Yoshida, S. (2018). Host lignin composition affects haustorium induction in the parasitic plants Phtheirospermum japonicum and Striga hermonthica. New Phytol. 218(2): 710–723.
Yoshida, S., Cui, S., Ichihashi, Y. and Shirasu, K. (2016). The Haustorium, a Specialized Invasive Organ in Parasitic Plants. Annu Rev Plant Biol. 67(1): 643–667.
Bouwmeester, H., Li, C., Thiombiano, B., Rahimi, M. and Dong, L. (2021). Adaptation of the parasitic plant lifecycle: germination is controlled by essential host signaling molecules. Plant Physiol. 185(4): 1292–1308.
Graves, J. D., Press, M. C. and Stewart, G. R. (2006). A carbon balance model of the sorghum‐Striga hermonthica host‐parasite association. Plant Cell Environ 12(1): 101–107.
Goldwasser, Y. and Rodenburg, J. (2013). Integrated Agronomic Management of Parasitic Weed Seed Banks. In: Joel, D. M., Gressel, J. and Musselman, L. J. (Eds.). Parasitic Orobanchaceae. Springer Berlin Heidelberg. 393–413.
Jamil, M., Kountche, B. A. and Al-Babili, S. (2021). Current progress in Striga management. Plant Physiol. 185(4): 1339–1352.
Nzioki, H. S., Oyosi, F., Morris, C. E., Kaya, E., Pilgeram, A. L., Baker, C. S. and Sands, D. C. (2016). Striga Biocontrol on a Toothpick: A Readily Deployable and Inexpensive Method for Smallholder Farmers. Front Plant Sci. 7: e01121.
Gobena, D., Shimels, M., Rich, P. J., Ruyter-Spira, C., Bouwmeester, H., Kanuganti, S., Mengiste, T. and Ejeta, G. (2017). Mutation in sorghum LOW GERMINATION STIMULANT 1 alters strigolactones and causes Striga resistance. Proc Natl Acad Sci USA. 114(17): 4471–4476.
Kawa, D., Taylor, T., Thiombiano, B., Musa, Z., Vahldick, H. E., Walmsley, A., Bucksch, A., Bouwmeester, H. and Brady, S. M. (2021). Characterization of growth and development of sorghum genotypes with differential susceptibility to Striga hermonthica. J Exp Bot. 72(22): 7970–7983.
Article Information
Publication history
Received: May 4, 2024
Accepted: Jul 17, 2024
Available online: Aug 9, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Plant Science > Plant immunity > Host-microbe interactions
Microbiology > Microbe-host interactions
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5,059 | https://bio-protocol.org/en/bpdetail?id=5059&type=0 | # Bio-Protocol Content
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An Automated pre-Dilution Setup for Von Willebrand Factor Activity Assays
TS Tobias Schachinger
AH Ann-Katrin Holik
GS Gerald Schrenk
HG Herbert Gritsch
SH Stefan Hofbauer
PF Paul G. Furtmüller
PT Peter L. Turecek
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5059 Views: 341
Reviewed by: Anupama SinghNeha NandwaniNavnita Dutta
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Abstract
Accurate quantification of von Willebrand factor ristocetin cofactor activity (VWF:RCo) is critical for the diagnosis and classification of von Willebrand disease, the most common hereditary and acquired bleeding disorder in humans. Moreover, it is important to accurately assess the function of von Willebrand factor (VWF) concentrates within the pharmaceutical industry to provide consistent and high-quality biopharmaceuticals. Although the performance of VWF:RCo assay has been improved by using coagulation analyzers, which are specialized devices for blood and blood plasma samples, scientists still report a high degree of intra- and inter-assay variation in clinical laboratories. Moreover, high, manual sample dilutions are required for VWF:RCo determination of VWF concentrates within the pharmaceutical industry, which are a major source for assay imprecision. For the first time, we present a precise and accurate method to determine VWF:RCo, where all critical pipetting and mixing steps are automated. A pre-dilution setup was established on CyBio FeliX (Analytik-Jena) liquid handling system, and an adapted VWF:RCo method on BCS-XP analyzer (Siemens) is used. The automated pre-dilution method was executed on three different, most frequently used coagulation analyzers and compared to manual pre-dilutions performed by an experienced operator. Comparative sample testing revealed a similar assay precision (coefficient of variation = 5.9% automated, 3.1% manual pre-dilution) and no significant differences between the automated approach and manual dilutions of an expert in this method. While no outliers were generated with the automated procedure, the manual pre-dilution resulted in an error rate of 8.3%. Overall, this operator-independent protocol enables standardization and offers an efficient way of fully automating VWF activity assays, while maintaining the precision and accuracy of an expert analyst.
Key features
• Automated pre-dilution setup for von Willebrand factor concentrates of various natures.
• Combination of a liquid handling system (CyBio FeliX) with a coagulation analyzer (BCS-XP).
• Simplifies method transfer to other laboratories.
• Basic training for CyBio FeliX and BCS-XP is required.
Keywords: Automation Blood coagulation Ristocetin cofactor VWF:RCo Von Willebrand disease Von Willebrand factor
Graphical overview
VWF:RCo assay principle and measurement setup. Platelets (yellow ellipsoids) with negative surface charge (- - -) are treated with formaldehyde, which partly denatures the cell surface and thus stabilizes platelets for use as assay reagents. Stabilized platelets (dark-yellow-framed yellow ellipsoids) are then brought in contact with ristocetin A (chemical structure shown; black dots), which binds to the platelet surface and facilitates binding of VWF (green circles). The graphs show an example of quantitative determination of platelet agglutination by measurement of light transmission, where increasing amounts of VWF increase light transmission over time. The photo in the left-bottom corner shows the CyBio FeliX setup for VWF sample dilution and the photo in the right-bottom corner displays the BCS-XP system, which is used for VWF:RCo measurements.
Background
Von Willebrand factor (VWF) is the largest soluble plasma protein in all species with a blood clotting system; it has essential functions in blood coagulation, control of bleeding, and angiogenesis [1,2]. Shear stress in the bloodstream plays a crucial role in the functional characteristics of VWF. While, for example, high shear rates are needed for the binding of VWF to GPIbα, the shear rate needs to be below a critical level for effective binding to coagulation factor VIII (Figure 1) [3]. VWF plays an important role in the pathology of various diseases, including bacterial infections and cardiovascular conditions [4,5]. Another example where VWF is highly involved is in diseases resulting from inherited or immune-mediated deficiencies of ADAMTS13, a metalloprotease that serves as the natural regulator of VWF size and function through specific proteolysis [6,7]. Furthermore, most recently, an association between a VWF/ADAMTS13 imbalance and the prolonged presence of highly thrombogenic ultra-high-molecular-weight (UHMW) VWF multimers was identified in patients with severe COVID-19 [8,9]. As a consequence, the VWF:RCo assay is now widely performed for research and the differential diagnosis of diseases in which VWF and ADAMTS13 are involved in pathological mechanisms and/or serve as valid biomarkers guiding therapies. Deficiencies or defects of VWF result in von Willebrand disease (VWD), the most frequent inherited bleeding disorder with a prevalence of 1 in 1000 individuals. VWD is a heterogeneous disease with different genetic subtypes called type 1, type 2 (with subtypes 2A, 2B, 2M, and 2N), and type 3 [10,11].
Besides genetic analysis, differential diagnosis of VWD requires the determination of VWF activity by different functional assays, of which the von Willebrand factor Ristocetin cofactor activity (VWF:RCo) assay is the most important. Ristocetin A is an antibiotic effective against Gram-positive bacteria and mycobacteria. Its use was discontinued due to its association with thrombocytopenia. However, this effect is utilized to, at least partially, mimic primary hemostasis in vitro by facilitating the binding of VWF to GPIbα, leading to platelet agglutination [12]. A recently published survey among expert laboratories revealed that the VWF:RCo assay is the most commonly established VWF assay worldwide [13]. It is needed for the diagnosis of all VWD subtypes except for type 2N. While the other subtypes are, in part, characterized by lower VWF:RCo levels, type 2N typically exhibits normal VWF:RCo. Instead, this subtype involves reduced binding to coagulation factor VIII [14]. The VWF:RCo assay is not only the standard functional assay for the diagnosis of VWD but is required for potency assignment to VWF concentrates within the pharmaceutical industry [15–17]. The European Pharmacopoeia defined this assay and international VWF standard samples to determine the VWF activity of VWF-containing drug products. The assigned activity is then used to specify the dosage of these biopharmaceuticals [18]. Besides its broad utilization, the assay requires technical expertise, is labor intensive, and has a high degree of inter- and intra-assay variation [15]. Recently issued international clinical guidelines call for further research improving the VWF:RCo assay, which “has greater variability, resulting in the potential for misdiagnosis and/or misclassification” [14].
The performance of the VWF activity assays has already been improved and simplified for operators by automation on dedicated coagulation analyzers [19,20]. These devices are commonly used worldwide to determine VWF activity [17,21–23]. In contrast to diagnostic plasma samples, plasma-derived VWF (e.g., Immunate® and Biostate®) and recombinant VWF (rVWF) products (Vonvendi® or Veyvondi®) are highly concentrated. To obtain the concentration range prescribed by the European Pharmacopoeia for the measurement of VWF:RCo, these biopharmaceuticals need to be diluted, which is mainly performed manually [18].
Here, we demonstrate an approach to determine the VWF:RCo of VWF concentrates, where all critical pipetting and mixing steps are automated. We have established a pre-dilution setup for VWF concentrates on a CyBio FeliX system (Analytik-Jena). This liquid handling system (LHS) is versatile, compact, and affordable, making it interesting not only for the pharmaceutical industry but also for academia. Despite its benchtop size of only 650 mm × 450 mm × 700 mm, it can be employed for a wide range of applications. For example, it is used for DNA purification, transformation of bacterial cells, and purification of Aβ peptides for the diagnosis of Alzheimer's disease [24–26]. The combination of the LHS and the adapted method on BCS-XP provides a precise and accurate determination of VWF activity in samples of various natures of VWF concentrates, and VWF-containing samples, without requiring specific training for the VWF:RCo assay. This ensures accurate diagnosis and potency assignment of VWF as a drug substance and drug product. Moreover, this approach offers an operator-independent method, which may also reduce errors compared to manual dilutions. With fewer errors, fewer measurements need to be repeated, resulting in reduced hands-on time for operators. Furthermore, fewer errors result in reduced reagent consumption, thereby increasing cost efficiency. In addition, implementation of the automated pre-dilution steps simplifies method transfer to other laboratories and lays the foundation for the full automation of various VWF assays.
Figure 1. Schematic domain structure of von Willebrand factor according to Lenting et al. [1]. Binding domains of coagulation factor (FVIII), platelet glycoprotein Ib alpha (GPIbα), and different collagens are shown.
Materials and reagents
Reagents
Distilled water
Imidazole + 1% albumin buffer (Baxter AG Vienna, catalog number: 1501390)
BC von Willebrand reagent (Siemens, catalog number: OUBD37)
Equipment
CyBio FeliX (Analytic-Jena, catalog number: OL5015-100-24)
BCS-XP (Siemens)
Adapter 24 tubes, passive cooling function (Analytik-Jena, catalog number: 844-00136-0)
Waste box (Analytik-Jena, catalog number: 844-00430-0)
TipRack 96/1,000 µL (Analytik-Jena, catalog number: OL3317-11-140)
CyBio Robo tip tray 96/1,000 µL (Analytik-Jena, catalog number: OL3810-25-871)
12-channel adapter (Analytik-Jena, catalog number: OL3316-14-340)
Support; 37 mm height (Analytik-Jena, catalog number: OL3317-11-120)
Axygen 8-row v-bottom high-profile reservoir (VWF, catalog number: 47743-966)
Protective cap (Analytik-Jena, catalog number: OL3316-11-200)
96-deep-well plate (VWR, catalog number: 736-0607)
5 mL glass flasks (Siemens, catalog number: 10873438)
Magnetic stirrer (Siemens, catalog number: 10642244)
Pipettes and tips
1.5 mL tubes
Software and datasets
CyBio FeliX script (shown in Supplementary information 1)
BCS-XP tests (shown in Supplementary information 2)
Microsoft Excel
Procedure
The protocol consists of two major parts: the automated dilution using the liquid handling system CyBio FeliX and the VWF:RCo assay on the BCS-XP. The CyBio FeliX system dilutes the VWF samples to 1 IU/mL VWF:RCo. The diluted samples are then transferred manually to the BCS-XP, where the automated VWF:RCo assay is executed, and the agglutination of the platelets in the reagent is measured. Data analysis is performed in Microsoft Excel. For a better understanding of the general functioning of the CyBio FeliX and BCS-XP and how to work with these devices, example videos (not specific to this protocol) are provided in Supplementary Information 3.
Pre-dilution using CyBio FeliX
Bring the samples and the imidazole + 1% albumin buffer to room temperature (RT).
Turn on the CyBio FeliX system.
Put 1,000 µL tips in the 96/1,000 tip rack:
Complete rows A and B.
Row C: C5–C12.
The remaining wells stay empty.
Put the samples and empty 1.5 mL tubes (left open) in the 24-tube adapter (Table 1). The diluted samples are pipetted into the empty tubes in the same order as the undiluted samples (Table 1).
Table 1. Scheme of the 24-tube adapter for CyBio FeliX. The respective VWF sample, VWF standard sample, VWF control sample, and empty tubes are placed as shown in the table.
1 2 3 4 5 6
A Standard sample Sample 3 Empty tube Empty tube
B Control sample Sample 4 Empty tube Empty tube
C Sample 1 Sample 5 Empty tube Empty tube
D Sample 2 Sample 6 Empty tube Empty tube
Fill 12 mL of imidazole + 1% albumin buffer in well 1 of the 8-row v-bottom reservoir.
Equip the CyBio FeliX system:
Position 1: 96/1,000 tip rack
Position 3: Waste box
Position 4: 12-channel adapter
Position 6: 37 mm adapter for protective cap
Position 7: 24-tube adapter
Position 8: 96-deep-well plate
Position 9: 8-row v-bottom reservoir
Execute the VWF dilution script. (The script setup is shown in Supplementary Information 1.)
VWF:RCo measurement using BCS-XP
Bring the BC VWF:RCo reagent to RT, reconstitute in 4 mL of distilled water, put a stirrer in the flask, and place it, without cap, in a cool rack of the BCS-XP system.
Place a 5 mL flask filled with imidazole + 1% albumin buffer, without cap, in a RT rack of the BCS-XP system.
After the automated dilution, the samples are in the empty tubes in the same order as before. Place the samples (in the open tubes) in a sample rack of the BCS-XP system.
Execute the VWF:RCo tests for all samples, including standard and control samples (test setups are shown in Supplementary Information 2):
rVWF:RCo_0.67
rVWF:RCo_0.50
rVWF:RCo_0.35
rVWF:RCo_0.20
Transfer the raw data to a Microsoft Excel sheet (calculation is shown in Data analysis section).
Data analysis
The VWF:RCo is calculated by linear regression in Microsoft Excel. The measured agglutination time is logarithmically transformed. The calibration comprises four VWF dilutions of the standard sample (Figure 2). Additionally, four dilutions are measured from the samples. Only the agglutination times from sample dilutions that fall within the calibration line are utilized to calculate the VWF:RCo. Subsequently, the mean of these activities is calculated and used as the final result for each sample.
The raw data from the BCS-XP system is presented in seconds (agglutination time). Transfer the respective raw data into the yellow fields of the Microsoft Excel evaluation sheet and enter the dilution factor of the samples in the respective green field (an example of the evaluation sheet is shown in Figure 3, and an example Microsoft Excel evaluation sheet including formulas can be found in Supplementary information 4).
Figure 2. Example of the calibration line. Generated by plotting the known VWF:RCo against the corresponding measured agglutination time (logarithmically transformed). A linear regression analysis was performed.
Figure 3. Example of the Microsoft Excel evaluation sheet. Raw data are transferred to yellow fields and the dilution factor of the samples is input into the green fields. The grey fields are used for calculations. An example evaluation sheet, including the formulas, is provided in Supplementary Information 4.
Validation of protocol
Von Willebrand factor samples
The recombinant von Willebrand factor (rVWF) used in this study is co-expressed with recombinant coagulation factor VIII in Chinese hamster ovary cells. It is highly purified by several downstream processes and formulated in a protein-free buffer [27]. This rVWF shows remarkable similarity in many aspects to the structure and function of the native protein [28]. It was licensed by the US Food and Drug Administration (FDA) in 2015 and by the European Medical Agency (EMA) in 2018.
Statistical analysis
Minitab® 21.2 was used for statistical analysis. Significant differences between operators, pre-dilution procedures, and BCS-XP systems were calculated using one-way ANOVA with a confidence level of 95%. Anderson-Darling normality test was carried out. Data were tested for outliers using Tukey’s Fence test. All data showed normal distribution (p > 0.05) within groups. As stated in the corresponding table, outliers were not included in the calculations.
Comparison of manual dilutions by three operators
In total, 18 rVWF sample aliquots were analyzed. Each of the three operators measured six sample aliquots. The mean VWF:RCo of all samples across all three operators was 139.5 IU/mL and the coefficient of variation (CV) was 6.3%. The CVs of the measurements of the single operators ranged from 1.4% to 4.6% (Table 2). There was a significant (p < 0.001) difference between operator 3 compared to the other operators. The variations of the operators are shown in Figure 4.
Table 2. Comparison of three operators. rVWF sample aliquots were diluted to 1 IU/mL and VWF:RCo was measured on BCS-XP. Operator 3 showed a significant (p < 0.001) difference compared to the other operators. Statistical calculations were carried out using Minitab® 21.2. n = 1 with 6 technical replicates.
Sample aliquot Operator 1 [IU/mL] Operator 2 [IU/mL] Operator 3 [IU/mL]
1 133.9 128.0 148.8
2 136.0 126.1 152.6
3 133.3 129.9 149.8
4 135.4 140.5 151.8
5 138.4 139.5 147.1
6 136.4 131.6 152.6
Mean VWF:RCo [IU/mL] 135.6 132.6 150.5
CV [%] 1.4 4.6 1.5
Total mean VWF:RCo [IU/mL] 139.5
Total CV [%] 6.3
Figure 4. Boxplots of the VWF:RCo measured by three operators. Each operator manually diluted six rVWF sample aliquots to 1 IU/mL and measured the VWF:RCo on a BCS-XP system. The graph was created using Minitab® 21.2. n = 1 with 6 technical replicates for each operator.
Comparison of three BCS-XP systems
In total, six rVWF sample aliquots were analyzed. The CVs of the sample aliquots, which were split up and measured on each of the three devices, ranged from 0.4% to 3.1% (Table 3). There was no significant (p = 0.296) difference between the three instruments.
Table 3. Comparison of three BCS-XP systems. rVWF sample aliquots were manually diluted to 1 IU/mL VWF:RCo. Each aliquot was separated into three parts for measurement on each of the devices. No significant (p = 0.296) difference between the three devices was shown. Statistical calculations were carried out using Minitab® 21.2. n = 1 with 6 technical replicates on each of the three devices.
Sample aliquots BCS-XP 1 [IU/mL] BCS-XP 2 [IU/mL] BCS-XP 3 [IU/mL] Mean VWF:RCo [IU/mL] CV [%]
1 121.5 123.3 125.3 123.4 1.6
2 128.4 128.0 127.4 128.0 0.4
3 127.8 126.4 131.0 128.4 1.8
4 131.0 138.8 137.1 135.6 3.0
5 127.4 132.9 132.0 130.8 2.3
6 127.5 135.1 134.1 132.2 3.1
Pre-dilutions by CyBio FeliX compared with manual pre-dilutions on BCS-XP
The precision of the CyBio FeliX pre-dilution script compared with manually pre-diluted samples was evaluated by analyzing 36 rVWF sample aliquots each. The automated pre-dilution procedure resulted in a mean activity of 139.0 IU/mL (Table 4), and the manual pre-dilution showed a mean of 136.3 IU/mL (Table 5). The mean CV of the samples pre-diluted by the CyBio FeliX system was 5.9%, and the mean CV of manually pre-diluted samples was 3.1%. The CV within one run of the manually pre-diluted samples ranged from 1.2% to 3.4% (Table 5) and of the automated samples from 1.2% to 5.5% (Table 4). The variations of both pre-dilution procedures are shown in Figure 5. The manually pre-diluted samples exhibited three outliers, yielding an error rate of 8.3% (Table 5). There was no significant difference (p = 0.098) between an experienced operator, who executed this assay many times, and the automated dilution procedure. Thereby, the CyBio FeliX approach provides a method that is as precise and accurate as an expert without requiring specific training for this assay. While significant differences can occur between operators who are not specifically trained for this method (as shown in the respective section above), the automated dilution showed reproducible results. Furthermore, it is less error-prone, as indicated by the error rate of 8.3%.
Table 4. VWF:RCo [IU/mL] of rVWF sample aliquots on six days. The samples were pre-diluted on a CyBio FeliX liquid handling system to 1 IU/mL, and VWF:RCo was measured on BCS-XP. Statistical calculations were carried out using Minitab® 21.2. Data were tested for outliers using Tukey’s Fence test. No outliers were observed. n = 6 with 6 technical replicates on each of six days.
Sample aliquots Day 1 [IU/mL] Day 2 [IU/mL] Day 3 [IU/mL] Day 4 [IU/mL] Day 5 [IU/mL] Day 6 [IU/mL]
1 144.7 145.5 147.3 149.7 139.4 130.7
2 137.9 147.7 149.2 148.6 122.1 127.5
3 138.9 143.3 149.2 151.1 132.7 131.9
4 138.0 134.0 146.3 143.1 140.4 131.0
5 125.6 141.1 144.4 145.2 133.3 130.5
6 142.7 133.6 131.7 148.6 125.8 130.8
Mean VWF:RCo [IU/mL] 137.9 140.9 144.7 147.7 132.3 130.4
CV [%] 4.8 4.2 4.6 2.0 5.5 1.2
Total mean VWF:RCo [IU/mL] 139.0
Total CV [%] 5.9
Table 5. VWF:RCo [IU/mL] of rVWF sample aliquots on six days. The samples were pre-diluted manually by one operator to 1 IU/mL and VWF:RCo was measured on BCS-XP. Statistical calculations were carried out using Minitab® 21.2. Data were tested for outliers using Tukey’s Fence test. The outliers are marked in bold and were excluded from the calculations of mean and CV. The error rate was calculated by dividing the number of outliers by the total number of observations. n = 6 with 6 technical replicates on each of six days.
Sample aliquots Day 1 [IU/mL] Day 2 [IU/mL] Day 3 [IU/mL] Day 4 [IU/mL] Day 5 [IU/mL] Day 6 [IU/mL]
1 144.7 144.7 133.9 130.1 130.9 136.2
2 135.5 135.5 136.0 135.2 133.9 133.4
3 138.9 138.9 133.3 134.0 131.3 133.7
4 169.6 169.6 135.4 141.9 131.7 154.4
5 142.4 142.4 138.4 134.3 133.9 140.9
6 136.2 136.2 136.4 130.0 134.7 139.2
Mean VWF:RCo [IU/mL] 139.5 140.4 135.6 134.3 132.7 136.7
CV [%] 2.8 3.4 1.4 3.3 1.2 2.4
Total mean VWF:RCo [IU/mL] 136.3
Total CV [%] 3.1
Error rate [%] 8.3
Figure 5. Comparison of two rVWF pre-dilution procedures. rVWF sample aliquots (n = 36) were diluted by a CyBio FeliX liquid handling system and manually by one operator. VWF:RCo was measured on BCS-XP. There is no significant difference (p = 0.098) between manual dilutions and those performed by the CyBio FeliX system. The graph was created using Minitab® 21.2. n = 1 with 6 technical replicates on each of six days.
Pre-dilutions by CyBio FeliX compared with manual pre-dilutions on different coagulation analyzers
The performance of the pre-dilution script on the CyBio FeliX system was assessed on three different coagulation analyzers [BCS-XP (Siemens, Germany), Stago STA compact max (Stago, France), ACL TOP 500 (Instrumentation Laboratory, USA)]. In total, 36 rVWF sample aliquots were analyzed. On each of the devices, six sample aliquots pre-diluted manually as well as six sample aliquots pre-diluted by CyBio FeliX were measured. The mean VWF:RCo was calculated and normalized to the mean separately for both pre-dilution procedures on each device. The summary of the results is shown in Table 6. The two pre-dilution methods show similar variations on each device, as shown in Figure 6.
Table 6. Comparison of pre-dilutions by a CyBio FeliX liquid handling system with manual pre-dilutions of rVWF sample aliquots. The samples were diluted to 1 IU/mL and VWF:RCo was measured on three different coagulation analyzers: ACL TOP (Instrumentation Laboratory, USA), BCS-XP (Siemens, Germany), and Stago STA Compact Max (Stago, France). The VWF:RCo was normalized to the mean separately for both pre-dilution procedures on each device. Statistical calculations were carried out using Minitab® 21.2. n = 1 with 6 technical replicates on each of the three devices.
Coagulation analyzer Pre-dilution Normalized VWF:RCo [%] CV [%]
ACL TOP 500 Manual 92.0–104.7 4.8
CyBio FeliX 95.8–102.1 2.3
BCS-XP Manual 83.8–110.2 8.8
CyBio FeliX 89.9–108.8 7.4
Stago STA compact max Manual 83.5–115.5 11.5
CyBio FeliX 83.8–115.0 12.5
Figure 6. Comparison of two pre-dilution methods for measurement of VWF:RCo on three different devices. rVWF sample aliquots (n = 6) were pre-diluted manually and by a CyBio FeliX liquid handling system to 1 IU/mL. Three coagulation analyzers were used for the VWF:RCo measurements: ACL TOP (Instrumentation Laboratory, USA), BCS-XP (Siemens, Germany), and Stago STA Compact Max (Stago, France). The VWF:RCo was normalized to the mean separately for both pre-dilution procedures on each device. Normalized data and the graph were created using Minitab® 21.2. n = 1 with six technical replicates on each of the three devices.
General notes and troubleshooting
The CyBio FeliX script dilutes the VWF samples to 1 IU/mL VWF:RCo. This concentration demonstrated accurate and precise results using several coagulation analyzers [29,17]. In addition, the expected concentration of VWF in the blood is 1 IU/mL, which is routinely measured in the clinic using a variety of instruments [30]. In this study, a drug product of a specific supplier was used. Hence, when measuring other VWF samples, it should be noted that the dilution steps in the CyBio FeliX script may need to be adjusted accordingly. The main difference of the adapted method on BCS-XP, as compared to the pre-installed procedure, lies in the thorough nature of the measurement. Instead of only using four dilutions of the calibrator, this approach evaluates four dilutions from every sample. Furthermore, modifications were made to the programming of the BCS-XP system to ensure that measurements were exclusively obtained within the linear range of the assay. Consequently, the point-to-point calibration was replaced with a linear regression.
We included a comparison between the two dilution procedures on different coagulation analyzers in the validation section to emphasize that the automated dilution setup can be combined with devices other than BCS-XP. However, we still recommend using the CyBio FeliX system in combination with the adapted VWF:RCo assay on BCS-XP, as described in the protocol. This approach demonstrated reproducible results over six days. Further research is needed for a meaningful assessment of the other coagulation analyzers.
Compared with other more sophisticated LHS such as Tecan Fluent (Tecan, Switzerland), the CyBio FeliX is cheaper and more compact. Despite its smaller size and lower cost, it remains versatile, as described in the background section. Furthermore, the automated dilution setup creates the basis for the full automation of this assay and several other VWF analyses, such as VWF:Ag and VWF:GPIbR, on the CyBio FeliX system.
The CyBio FeliX system is robust and executes the dilution script without errors. However, it is important to double-check for human errors, such as providing too little buffer or too few tips, as the system does not track the liquid level or when there are not enough tips.
Also, there are no common errors related to the BCS-XP device itself. While this system recognizes insufficient volumes, it is still important to provide enough volume to avoid errors due to a disrupted workflow. In addition, ensure the magnetic stirrer is in the reagent flask, as the system will not detect its absence.
Further research is needed to validate the protocol for utilization with VWF-containing samples other than rVWF concentrates. While this protocol is most suitable for use in research labs, it paves the way for high-throughput applications in the pharmaceutical industry and clinical settings.
Acknowledgments
The authors would like to thank Elisabeth Pum and Christina Dirnberger-Elboraei for performing the experiments for the operator comparison.
Author Contributions: Conceptualization: Tobias Schachinger, Ann-Katrin Holik, Gerald Schrenk, and Herbert Gritsch; Methodology: Tobias Schachinger, Ann-Katrin Holik, Gerald Schrenk, and Herbert Gritsch; Writing—original draft preparation: Tobias Schachinger; Writing—review and editing: Ann-Katrin Holik, Gerald Schrenk, Herbert Gritsch, Peter Turecek, Paul G. Furtmüller, and Stefan Hofbauer; Supervision: Ann-Katrin Holik, Gerald Schrenk, Herbert Gritsch, Paul G. Furtmüller, and Stefan Hofbauer. All authors have read and agreed to the published version of the manuscript.
Competing interests
Baxalta Innovations GmbH (part of Takeda) provided funding for this research. Authors Tobias Schachinger, Ann-Katrin Holik, Gerald Schrenk, Herbert Gritsch, and Peter L. Turecek are employed at Baxalta Innovations GmbH, part of Takeda. Gerald Schrenk, Herbert Gritsch, and Peter L. Turecek hold stocks of Takeda. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
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Supplementary information
The following supporting information can be downloaded here:
Supplementary information 1: CyBio FeliX script
Supplementary information 2: BCS-XP tests
Supplementary information 3: Video demonstrations
Supplementary information 4: Example of the Microsoft Excel evaluation sheet including formulas
Article Information
Publication history
Received: Mar 21, 2024
Accepted: Jul 22, 2024
Available online: Aug 8, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Single-Molecule Sequencing of the C9orf72 Repeat Expansion in Patient iPSCs
YT Yu-Chih Tsai *
KB Katherine A. Brown *
MB Mylinh T. Bernardi *
JH John Harting
CC Claire D. Clelland
(*contributed equally to this work)
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5060 Views: 600
Reviewed by: Marion HoggSrinidhi Rao Sripathy RaoPrashanth N Suravajhala
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Original Research Article:
The authors used this protocol in Scientific Reports Jan 2024
Abstract
A hexanucleotide GGGGCC repeat expansion in the C9orf72 gene is the most frequent genetic cause of amyotrophic lateral sclerosis (ALS) and frontal temporal dementia (FTD). C9orf72 repeat expansions are currently identified with long-range PCR or Southern blot for clinical and research purposes, but these methods lack accuracy and sensitivity. The GC-rich and repetitive content of the region cannot be amplified by PCR, which leads traditional sequencing approaches to fail. We turned instead to PacBio single-molecule sequencing to detect and size the C9orf72 repeat expansion without amplification. We isolated high molecular weight genomic DNA from patient-derived iPSCs of varying repeat lengths and then excised the region containing the C9orf72 repeat expansion from naked DNA with a CRISPR/Cas9 system. We added adapters to the cut ends, capturing the target region for sequencing on PacBio’s Sequel, Sequel II, or Sequel IIe. This approach enriches the C9orf72 repeat region without amplification and allows the repeat expansion to be consistently and accurately sized, even for repeats in the thousands.
Key features
• This protocol is adapted from PacBio’s previous “no-amp targeted sequencing utilizing the CRISPR-Cas9 system.”
• Optimized for sizing C9orf72 repeat expansions in patient-derived iPSCs and applicable to DNA from any cell type, blood, or tissue.
• Requires high molecular weight naked DNA.
• Compatible with Sequel I and II but not Revio.
Keywords: C9orf72 Single-molecule sequencing No amplification Repeat expansion iPSCs
Graphical overview
Background
The most frequent genetic cause of amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD) is an intronic repeat expansion in the C9orf72 gene [1–3]. While non-diseased alleles typically have 10 or fewer GGGGCC repeats, expanded alleles can have thousands [1,2,4,5]. Accurately measuring the C9orf72 repeat expansion length applies to both clinical diagnosis and answering fundamental questions about the function of C9orf72 in health and disease. Traditional short-read sequencing methods fail to size the C9orf72 repeat region because amplification fails across the GC-rich region of the first intron of C9orf72, and short-read sequencing does not provide unique sequences adjacent to GGGGCC repeats to permit their alignment [5]. Instead, long-range PCR [6] and Southern blot [7] are used as clinical diagnostic and research tools to identify C9orf72 repeat expansions, but they are limited. Long-range PCR fails for repeats greater than 145 and, while Southern blot can identify long repeats, it requires a large amount of input DNA [5].
Single-molecule sequencing is an accurate, reliable, and sensitive method to size repeat lengths of varied sizes, showing success in sequencing the C9orf72 repeat expansion in both plasmid and tissue [5,8–10]. This amplification-free process employs a CRISPR/Cas9 system to excise and sequence the region containing the C9orf72 expansion in high-quality DNA from patient iPSCs (see Graphical overview). The excised genomic fragment is then sequenced from end to end, providing phased sequencing of the targeted region without the need for bioinformatic imputation of its sequence or structure. In addition to identifying and quantifying repeat expansions of various lengths, the sequencing data provides insights into the gene structure. Single-molecule sequencing can identify mosaicism within samples and help determine the stability of repeat lengths over time [5,8,10]. It can be used to assess cell line clonality, editing outcomes, and differences between each allele of a gene in a single individual [5,10]. This method provides additional information on the methylation of each allele within an individual or across a population [10].
Despite the technological advances afforded by single-molecule sequencing, there are a few limitations to employing this method in research and clinical settings. The first is cost. Single-molecule sequencing is expensive. The ability to multiplex samples, as we describe here, reduces the cost of this protocol. The second limitation is the variability in read depth with longer repeat expansions. As the size of repeat expansion increases, the number of reads on the expanded allele tends to decrease [5]. Long expansions can still be sized, albeit with reduced read depth. Over time, as technology improves and the number of reads for each run increases, this bias is likely to decrease.
We have optimized the protocol presented here for genomic DNA extracted from human induced pluripotent stem cell (iPSC) lines derived from C9-ALS/FTD patient samples. This protocol can also be applied to other sources of DNA such as any cultured cell type as well as donor blood and fresh or frozen tissue. This method can also be used to target other repeat expansions such as TCF4 and DPMK, for which this approach was initially reported [11,12], or other genomic regions of interest by targeting the Cas9/gRNA excision to the target region. This protocol is compatible with Sequel I and Sequel II. PacBio’s PureTargetTM Repeat Expansion Panel [13], which is an improved version of the NoAmp protocol, can be used additionally with Revio and quantifies 20 different repeat expansions for targeted single-molecule sequencing simultaneously and in the same individual.
Materials and reagents
Biological materials
5–25 μg of DNA from induced pluripotent stem cells (iPSCs) with C9orf72 repeat expansions (also compatible with DNA from blood or tissue)
Reagents
DPBS (Gibco, catalog number: 14190235)
ReLeSR (STEMCELL Technologies, catalog number: 05872)
Genomic DNA buffer set (QIAGEN, catalog number: 19060)
Proteinase K (Qiagen, catalog number: 19131)
QubitTM 1× dsDNA HS Assay kit (Thermo Fisher Scientific, catalog number: Q33230)
Agarose (Fisher Scientific, catalog number: BP164-500)
50× TAE buffer (Thermo Fisher Scientific, catalog number: B49)
SYBRTM Safe DNA gel stain (Thermo Fisher Scientific, catalog number: S33102)
6× gel loading dye (New England BioLabs, catalog number: B7024S)
Nuclease-free water, not DEPC treated (Ambion, catalog number: AM9937)
Ethanol, molecular biology grade
Isopropyl alcohol, molecular biology grade
Custom single-guide RNA (sgRNA 1) TTGGTATTTAGAAAGGTGGT (Synthego)
Custom single-guide RNA (sgRNA 2) GGAAGAAAGAATTGCAATTA (Synthego)
1× TE buffer (included with sgRNA orders from Synthego)
Custom barcoded adapter oligos (IDT)
Note: See Table 1 for a list of barcoded adapter oligos.
Table 1. Barcoded adapters
Barcoded adapter Sequence
Barcoded adapter 1 /5Phos/CGCACTCTGATATGTGATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATCACATATCAGAGTGCG
Barcoded adapter 2 /5Phos/CTCACAGTCTGTGTGTATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATACACACAGACTGTGAG
Barcoded adapter 3 /5Phos/CGCAGCGCTCGACTGTATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATACAGTCGAGCGCTGCG
Barcoded adapter 4 /5Phos/TCTGTCTCGCGTGTGTATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATACACACGCGAGACAGA
Barcoded adapter 5 /5Phos/CTCTGAGATAGCGCGTATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATACGCGCTATCTCAGAG
Barcoded adapter 6 /5Phos/ACACGCGATCTAGTGTATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATACACTAGATCGCGTGT
Barcoded adapter 7 /5Phos/ACGCGCGCGTAGTGAGATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATCTCACTACGCGCGCGT
Barcoded adapter 8 /5Phos/ACACACGTGTCATGCGATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATCGCATGACACGTGTGT
Barcoded adapter 9 /5Phos/ATACTATCTCTCTATGATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATCATAGAGAGATAGTAT
Barcoded adapter 10 /5Phos/CACAGTGAGCACGTGAATCTCTCTCTTTTCCTCCTCCTCCGTTGTTGTTGTTGAGAGAGATTCACGTGCTCACTGTG
Shrimp alkaline phosphatase (rSAP) (New England BioLabs, catalog number: M0371S or M0371L)
CutSmart® buffer 10× (New England BioLabs, catalog number: B7204S)
Exonuclease III (New England BioLabs, catalog number: M0206S or M0206L)
Cas9 nuclease, S. pyogenes (New England BioLabs, catalog number: M0386T or M0386M)
NEBufferTM 3.1, 10× (New England BioLabs, catalog number: B7203S)
Note: Included with Cas9 Nuclease, S. pyogenes from New England BioLabs.
T4 DNA ligase reaction buffer, 10× (New England BioLabs, catalog number: B0202S)
T4 DNA ligase, HC (Thermo Fisher Scientific, catalog number: EL0013)
SOLu-Trypsin (Sigma-Aldrich, catalog number: EMS0004)
1 kb DNA ladder (carrier DNA) (New England BioLabs, catalog number: N3232S or N3232L)
Recombinant ribonuclease inhibitor (Takara Bio, catalog number: 2313A or 2313B)
Pacific Biosciences® Binding Kits
For Sequel: Sequel Binding and Internal Control kit 3.0 (PacBio, catalog number: 101-626-600)
For Sequel II and IIe: Sequel II Binding kit 2.0 and Internal Control kit 1.0 (PacBio, catalog number: 101-842-900)
Pacific Biosciences® Sequencing Kits
For Sequel: Sequel Sequencing kit 3.0 (PacBio, catalog number: 101-597-900)
Note: Four reactions per kit.
For Sequel II and IIe: Sequel II Sequencing kit 2.0 (PacBio, catalog number: 101-820-200)
Note: Four reactions per kit.
Pacific Biosciences® SMRT® cells
For Sequel: SMRT® Cell 1M v3 tray or SMRT® Cell 1M v3 LR tray (PacBio, catalog number: 101-531-000 or 101-531-001)
For Sequel II and IIe: SMRT Cell 8M tray (PacBio, catalog number: 101-389-001)
AMPure® PB beads (PacBio, catalog number: 100-265-900)
Elution buffer (PacBio, catalog number: 101-633-500)
No-Amp Accessory kit (PacBio, catalog number: 101-788-900)
Sequencing primer v4
10× primer buffer v2
10× annealing buffer
SMRTbell® Enzyme Clean-up kit (PacBio, catalog number: 101-746-400)
Sample plate (PacBio, catalog number: 000-448-888)
Solutions
80% ethanol (see Recipes)
Recipes
80% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 80% 40 mL
Nuclease-free H2O n/a 10 mL
Total n/a 50 mL
Laboratory supplies
15 mL conical tubes (Falcon, catalog number: 352096)
50 mL conical tube (Falcon, catalog number: 1443222)
QIAGEN® genomic tip 100/G (QIAGEN, catalog number: 10243)
QubitTM assay tubes (Thermo Fisher Scientific, catalog number: Q32856)
1.5 mL DNA LoBind tubes (Eppendorf, catalog number: 0030108051)
0.2 mL PCR thermal cycling tubes (Genesee Scientific, catalog number: 27-125)
Equipment
Molecular biology pipettes, standard set
AvantiTM J-15R, IVD, refrigerated benchtop centrifuge (or equivalent) (Beckman Coulter, catalog number: B99517)
Platform shaker
QubitTM quantitation platform (Thermo Fisher Scientific, catalog number: Q33238)
Gel imaging system
Benchtop cooler
Microcentrifuge
Mini centrifuge
Vortex mixer
DynaMagTM-2 magnet (magnetic rack) (Thermo Fisher Scientific, catalog number: 12321D)
PCR thermal cycler
ThermoMixer C with heated lid (or equivalent) (Eppendorf, catalog number: 5382000023)
Sequel, Sequel II, or Sequel IIe Systems (Pacific Biosciences)
Software and datasets
SMRTLink (v12.0, 2023)
ccs (v7.0.0, https://github.com/PacificBiosciences/ccs)
lima (v2.7.1, 2023)
pbmm2 (v1.10.0, 2023)
primrose (v1.2.0, 2022, https://github.com/mattoslmp/primrose)
pb-CpG-tools (v2.3.2, 2023, https://github.com/PacificBiosciences/pb-CpG-tools)
Procedure
Genomic DNA extraction
Grow iPSCs in 2 wells of a 6-well plate to 80% confluency [14].
Aspirate the media and wash the cells once with 1 mL of DPBS.
Add 1 mL of ReLeSR to each well and incubate in the hood for 45 s.
Aspirate the ReLeSR and incubate at 37 °C for 3 min.
Resuspend the wells in 1 mL of DPBS each and transfer to a 15 mL conical tube.
Centrifuge at 300× g for 3 min.
Remove the supernatant.
Cell pellets may be immediately used for DNA extraction or can be stored at -20 °C.
To extract DNA from the cell pellet, follow the QIAGEN® Genomic DNA Handbook [15] for 100/G tips with the following modifications:
After adding 95 μL of Proteinase K, incubate at 55 °C for 90 min.
After precipitating with 3.5 mL of isopropyl alcohol, invert the tube four times and immediately centrifuge at 4,000× g for 20 min at 4 °C. Then, remove the supernatant.
After the ethanol wash, centrifuge at 4,000× g for 20 min at 4 °C and then remove the supernatant.
After the pellet air dries for 10 min, resuspend DNA in 50 μL of Buffer EB. At room temperature, dissolve the DNA in the elution buffer overnight on a shaker at 300 rpm.
Aliquot 3 μL of DNA for quality control and store the rest at -20 °C.
Evaluate DNA quality
Measure the concentration of your genomic DNA using the QubitTM 1× dsDNA HS Assay kit and associated protocol [16].
Note: Prior to measuring the concentrations, dilute genomic DNA 1:10.
It is recommended to evaluate sample quality with the Agilent FEMTO Pulse System, the Bio-Rad CHEF Mapper XA Pulsed Field Electrophoresis System, or the Sage Science Pippin Pulse Electrophoresis Power Supply. Genomic DNA samples should have fragment sizes ≥ 50 kb. Significant on-target reads can still be achieved with fragment sizes <50 kb by increasing the amount of input DNA, but better results may be achieved by collecting a fresh sample to achieve fragment sizes ≥ 50 kb.
Note: We skip this step. Instead, we evaluate DNA quality with a 1% agarose gel as described in steps B4–7 (Figure 1).
Prepare a 1% agarose gel with SYBRTM Safe DNA gel stain [17].
Add 2 μL of genomic DNA, 7 μL of water, and 1 μL of 6× gel loading dye to a PCR tube.
Pipette-mix gently and load the diluted DNA into the 1% gel.
Run the gel at 100 V for 1 h.
Image the gel and check for any smearing in your samples. Samples with a single band above 10 kb should be included. Samples that have smearing down the gel, even if they have the >10 kb band, indicate DNA shearing and should be excluded from the protocol. In the case of shearing, fresh genomic DNA samples should be collected.
Figure 1. 1% Agarose gel evaluation of genomic DNA quality. As a quick quality check, we run isolated DNA on a 1% agarose gel prior to initiating CRISPR isolation of the target region. We include only DNA that has high molecular weight (i.e., has no smearing). (A, B) Examples of genomic DNA samples extracted from C9orf72 patient iPSC lines and run on a 1% agarose gel to evaluate sample quality. (A) Samples 1, 2, 4, and 5 (A) and 6 (B) show no smearing and would be acceptable for use in the protocol. Samples 3 (A) and 7 (B) show significant smearing and therefore we recommend being excluded from this protocol. This is a quick quality check. Sample quality can be confirmed with higher accuracy by the Agilent FEMTO Pulse System, the Bio-Rad CHEF Mapper XA Pulsed Field Electrophoresis System, or the Sage Science Pippin Pulse Electrophoresis Power Supply.
Prepare reagents
Prior to starting the library preparation, prepare the PacBio barcoded adapters for multiplexing and the carrier DNA.
To prepare the PacBio barcoded adapters for multiplexing, resuspend the barcoded adapter oligos in nuclease-free water to 100 μM.
Add the following reagents, in order, to a PCR tube to make 20 μM stocks of barcoded adapters:
100 μM barcoded adapter oligo 10 μL
10× annealing buffer 5 μL
Nuclease-free water 35 μL
Place the 20 μM stocks in a thermal cycler and run the following protocol:
95 °C for 5 min
Decrease to 25 °C, ramping down at the maximum cooling rate
4 °C hold
Store stocks at -20 °C.
To prepare the carrier DNA (1 kb DNA ladder), add 5 μL of ladder to 45 μL of elution buffer in a PCR tube to dilute the carrier DNA to a concentration of 50 ng/μL.
Store at -20 °C.
Dephosphorylate the genomic DNA
In a PCR tube, dilute each genomic DNA sample to 5 μg with nuclease-free water for a total volume of 68 μL.
Note: When multiplexing five samples, 5 μg per sample is the optimal input. If using a different number of samples (1–10) for this protocol, the minimum total DNA input is 5 μg and the maximum total DNA input is 25 μg. Each multiplexed sample should have equimolar input amounts.
Add the following reagents, in order, to a LoBind microcentrifuge tube to make a master mix. Add 1 volume of each reagent per sample with 10% overage:
NEBuffer 8 μL
rSAP 4 μL
Add 12 μL of dephosphorylation master mix to each sample for a total volume of 80 μL.
Invert the tubes 20 times to mix.
Note: Do not vortex or flick the tube to avoid shearing the DNA.
Briefly spin down the tubes in a mini centrifuge.
Place in the thermal cycler and run the following protocol:
37 °C for 1 h
65 °C for 10 min
4 °C hold
Prepare the single-guide RNAs (sgRNAs)
Spin down lyophilized sgRNAs.
Resuspend sgRNAs in TE buffer to 50 μM.
Aliquot sgRNAs and store at -80 °C to avoid freeze-thawing.
Dilute a 50 μM aliquot of each sgRNA to 5 μM.
In a PCR tube, add 4 μL of sgRNA1 and 4 μL of sgRNA2 per sample to make a master mix of diluted sgRNAs. You will need 4 μL of each gRNA per sample with 10% overage.
Prepare the gRNA/Cas9 complex
Add the following reagents, in order, to a LoBind microcentrifuge tube to make a master mix. Add 1 volume of each reagent per sample with 10% overage:
Nuclease-free water 7 μL
NEBuffer 3.1 2 μL
Cas9 nuclease 2 μL
Diluted sgRNAs (from section E) 8 μL
Pipette-mix the reaction.
Briefly spin down the tube in a mini centrifuge.
Transfer the master mix to a PCR tube and place it in a thermal cycler.
Incubate at 37 °C for 10 min.
Place the gRNA/Cas9 complex on ice.
Cas9 digestion
To each tube with dephosphorylated DNA, add the following reagents, in order, for a total volume of 100 μL:
Recombinant ribonuclease inhibitor 1 μL
gRNA/Cas9 complex (from section F) 19 μL
Invert the tubes 20 times to mix.
Note: Do not vortex or flick the tube to avoid shearing the DNA.
Briefly spin down the tubes in a mini centrifuge.
Incubate at 37 °C for 1 h.
Briefly spin down the tubes in a mini centrifuge and then place on ice.
Purify Cas9-digested DNA
Transfer the samples to new LoBind microcentrifuge tubes.
Add 400 μL of elution buffer to each sample for a total volume of 500 μL.
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.45× volume of AMPure PB beads to each tube for a total volume of 725 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Invert the tubes 20 times to mix.
Note: Do not vortex or flick the tube to avoid shearing the DNA.
Briefly spin down the tubes in a mini centrifuge.
Incubate for 15 min at room temperature.
While the reaction is incubating, prepare a solution of 80% ethanol (see Recipes) in a 50 mL tube. Store in a tightly capped polypropylene tube for a maximum of three days.
After 15 min, briefly spin down the tubes in a mini centrifuge.
Place the tubes in the magnetic tube rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 5 min).
While the tubes are on the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and transfer to new LoBind microcentrifuge tubes.
Note: Save supernatant to repeat purification steps in case insufficient DNA is recovered at the end of the purification protocol.
Wash the beads with 1 mL of 80% ethanol by slowly adding it to the side opposite the beads.
Note: Leave the tubes on the magnetic rack during the ethanol washes. Do not invert the tubes.
Wait 30 s, then slowly pipette up the ethanol and discard.
Repeat the ethanol wash two more times.
Remove the tubes from the magnetic rack and briefly spin them down in a mini centrifuge.
Note: Both the beads and ethanol should be at the bottom of the tubes.
Return the tubes to the magnetic rack.
With a P20 pipette, slowly pipette up any remaining ethanol and discard.
Repeat steps H16–18 until no ethanol droplets remain.
Remove the tubes from the magnetic rack and add 31 μL of elution buffer to each tube.
Invert the tube 10 times to mix.
If the beads are stuck on the tube wall, leave the tube on the bench for 3 min and then invert 10 more times.
Repeat until most of the beads are resuspended in the elution buffer.
Incubate the tubes at room temperature for 10 min.
Briefly spin down the tubes in a mini centrifuge.
Return the tubes to the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tubes for at least 5 min.
Carefully pipette up the supernatant, transfer to new PCR tubes, and place on ice.
Discard the beads.
Add 1 μL of sample to 4 μL of elution buffer to dilute the DNA.
Use 1 μL of the diluted DNA to measure the concentration using the QubitTM 1× dsDNA HS Assay kit and associated protocol [16].
Ligate adapters
Add 1 μL of annealed barcoded adapter to each sample for a total volume of 31 μL.
Note: Use different barcodes for samples that will be pooled together.
Add the following reagents, in order, to a LoBind microcentrifuge tube to make a master mix. Add 1 volume of each reagent per sample with 10% overage:
T4 DNA ligase reaction buffer 5 μL
Nuclease-free water 12.5 μL
Add 17.5 μL of master mix to each sample for a total volume of 48.5 μL.
Invert the tube 20 times to mix.
Add 1.5 μL of T4 DNA ligase to each sample for a total volume of 50 μL.
Invert the tube 20 times to mix.
Note: Do not vortex or flick the tube to avoid shearing the DNA.
Briefly spin down the tubes in a mini centrifuge.
Place in the thermal cycler and run the following protocol:
16 °C for 2 h (turn heated lid off for this step)
65 °C for 10 min
4 °C hold overnight
Pool samples
Transfer samples to new LoBind microcentrifuge tubes.
In a microcentrifuge, centrifuge the tubes at 14,000× g for 5 min.
Note: Pay attention to the orientation of the tubes.
Carefully transfer the supernatant to a new LoBind microcentrifuge tube on ice by pipetting up the liquid from the inner-facing tube wall.
Notes:
Do not disturb the pellet on the outer-facing wall.
If multiplexing, pool samples in equimolar amounts at this step. A minimum of 5 μg and a maximum of 25 μg of DNA is recommended per pool/sequencing run.
Discard the tubes with the pellets.
Purify SMRTbell library
Add elution buffer to the pooled samples for a total volume of 500 μL.
Note: Additional elution buffer is not necessary if the sample volume is ≥500 μL.
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.45× volume of AMPure PB beads to the tube for a total volume of 725 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Vortex or flick the tube to mix the solution.
Briefly spin down the tube in a mini centrifuge.
Vortex the tube at 2,000 rpm for 10 min to bind the DNA to the beads.
Note: The solution should be homogenous.
Briefly spin down the tube in a mini centrifuge.
Place the tube into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 5 min).
While the tube is in the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and transfer to a new LoBind microcentrifuge tube.
Wash the beads with 1 mL of 80% ethanol by adding to the side opposite of the beads.
Note: Leave the tube on the magnetic rack during the ethanol washes. Do not invert the tube.
Wait for 30 s, slowly pipette up the 80% ethanol, and discard.
Repeat the ethanol wash two more times.
Remove the tube from the magnetic rack and briefly spin down the tube in a mini centrifuge.
Note: Both the beads and ethanol should be at the bottom of the tube.
Return the tube to the magnetic rack.
With a P20 pipette, slowly pipette up any remaining ethanol and discard.
Repeat steps K14–16 until no ethanol droplets remain.
Remove the tube from the magnetic rack and add 100 μL of elution buffer per 5 μg of input genomic DNA to the beads.
Pipette-mix until the solution is homogeneous.
Incubate at room temperature for 5 min.
Vortex the tube at 2,000 rpm for 1 min.
Briefly spin down the tubes in a mini centrifuge.
Place the tube back on the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tubes for at least 5 min.
Carefully pipette up the supernatant, transfer to a new LoBind microcentrifuge tube, and place on ice.
Safe pausing point: Store SMRTbell library at 4 °C overnight or at -20 °C for longer storage.
Discard the beads.
Nuclease treatment
Add the following reagents, in order, to the tube with the SMRTbell library. Add 1 volume of each reagent per 100 μL of purified DNA:
Nuclease-free water 67.2
CutSmart buffer 20 μL
Exonuclease III 4.8 μL
Enzyme A (Enzyme Cleanup kit) 4 μL
Enzyme B (Enzyme Cleanup kit) 1 μL
Enzyme C (Enzyme Cleanup kit) 1 μL
Enzyme D (Enzyme Cleanup kit) 2 μL
Invert the tube 20 times to mix.
Briefly spin down the tubes in a mini centrifuge.
Incubate at 37 °C for 2 h and then place the digested SMRTbell library on ice.
Add 9 μL of SOLu-Trypsin per 200 μL of digested SMRTbell library to the tube.
Invert the tube 20 times to mix.
Briefly spin down the tube in a mini centrifuge.
Incubate at 37 °C for 20 min and place the SMRTbell library on ice.
Purify the nuclease-treated SMRTbell library
Transfer the SMRTbell library to a new LoBind microcentrifuge tube.
Add elution buffer to the sample for a total volume of 500 μL.
Note: Additional elution buffer is not necessary if the sample volume is ≥500 μL.
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.45× volume of AMPure PB beads to the tube for a total volume of 725 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Vortex or flick the tube to mix the solution.
Briefly spin down the tubes in a mini centrifuge.
Vortex the tube at 2,000 rpm for 10 min to bind the DNA to the beads.
Note: The solution should be homogenous.
Briefly spin down the tubes in a mini centrifuge.
Place the tube into the magnetic tube rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 5 min).
While the tube is in the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and transfer to a new LoBind microcentrifuge tube.
Note: Save supernatant to repeat purification steps in case insufficient DNA is recovered at the end of the purification protocol.
Wash the beads with 1 mL of 80% ethanol by adding to the side opposite of the beads.
Note: Leave the tube on the magnetic rack during the ethanol washes. Do not invert the tube.
Wait for 30 s, slowly pipette up the 80% ethanol, and discard.
Repeat the ethanol wash two more times.
Remove the tube from the magnetic rack and briefly spin down the tube in a mini centrifuge.
Note: Both the beads and ethanol should be at the bottom of the tube.
Return the tube to the magnetic rack.
With a P20 pipette, slowly pipette up any remaining ethanol and discard.
Repeat steps M14–16 until no ethanol droplets remain.
Add 200 μL of elution buffer to the beads.
Pipette-mix until the solution is homogeneous.
Incubate at room temperature for 5 min.
Vortex the tube at 2,000 rpm for 1 min.
Briefly spin down the tubes in a mini centrifuge.
Place the tube back on the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tubes for at least 5 min.
Carefully pipette up the supernatant, transfer to a new LoBind microcentrifuge tube, and place on ice.
Discard the beads.
Purify the nuclease-treated SMRTbell library a second time
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.42× of AMPure PB beads to the tube from the first round of purification for a total volume of 284 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Vortex or flick the tube to mix the solution.
Briefly spin down the tube in a mini centrifuge.
Vortex the tube at 2,000 rpm for 10 min to bind the DNA to the beads.
Note: The solution should be homogenous.
Briefly spin down the tube in a mini centrifuge.
Place the tube into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 5 min).
While the tube is in the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and transfer to a new LoBind microcentrifuge tube.
Wash the beads with 1 mL of 80% ethanol by adding to the side opposite of the beads.
Note: Leave the tube on the magnetic rack during the ethanol washes. Do not invert the tube.
Wait for 30 s, slowly pipette up the 80% ethanol, and discard.
Repeat the ethanol wash two more times.
Remove the tube from the magnetic rack and briefly spin down the tube in a mini centrifuge.
Note: Both the beads and ethanol should be at the bottom of the tube.
Return the tube to the magnetic rack.
With a P20 pipette, slowly pipette up any remaining ethanol and discard.
Repeat steps N13–15 until no ethanol droplets remain.
Add 6.3 μL of elution buffer to the beads.
Pipette-mix until the solution is homogeneous.
Incubate at room temperature for 5 min.
Vortex the tube at 2,000 rpm for 1 min.
Briefly spin down the tube in a mini centrifuge.
Place the tube back on the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tubes for at least 5 min.
Carefully pipette up the supernatant, transfer to a new LoBind microcentrifuge tube, and place on ice.
Safe pausing point: Store SMRTbell library at 4 °C overnight or at -20 °C for longer storage.
If sending samples to a sequencing core, ship on dry ice for next-day delivery during this safe pausing point. The sequencing core should complete the rest of the protocol prior to sequencing.
Discard the beads.
Primer annealing
Add 1 μL of sequencing primer v4 stock to 29 μL of elution buffer.
Add the following reagents, in order, to a PCR tube:
10× primer buffer v2 36 μL
Diluted sequencing primer v4 18 μL
Pipette or flick the tube to mix the solution.
Briefly spin down the tubes in a mini centrifuge.
Place in the thermal cycler and run the following protocol:
80 °C for 2 min
4 °C hold
Transfer to a new LoBind microcentrifuge tube and place on ice.
Note: Store remaining conditioned sequencing primer at -20 °C for a maximum of 30 days.
Add the following reagents, in order, to a new PCR tube for a total volume of 9 μL:
Conditioned sequencing primer v4 (from step O6) 2.7 μL
SMRTbell library (from section L) 6.3 μL
Pipette or flick the tube to mix the solution.
Briefly spin down the tubes in a mini centrifuge.
Place in the thermal cycler and run the following protocol:
20 °C for 1 h
4 °C hold
Polymerase binding
For Sequel
Add 1 μL of Sequel DNA Polymerase 3.0 to 29 μL of Sequel Binding Buffer and place on ice.
Note: Diluted Sequel DNA Polymerase 3.0 must be used immediately; discard any excess.
Add the following reagents, in order, to the PCR tube with the primer-annealed SMRTbell library for a total volume of 13.5 μL:
Sequel Binding Buffer 1.5 μL
DTT 1.5 μL
Sequel dNTP 1.5 μL
Pipette or flick the tube to mix the solution.
Add 1.5 μL of Diluted Sequel DNA Polymerase 3.0 for a total volume of 15 μL.
Pipette or flick the tube to mix the solution.
Briefly spin down the tubes in a mini centrifuge.
Place the sample complex in the thermal cycler and run the following protocol:
30 °C for 4 h
4 °C hold (or hold on ice) until ready for the purification step
For Sequel II and IIe
Add 1 μL of Sequel II DNA Polymerase 2.0 to 29 μL of Sequel Binding Buffer and place on ice.
Note: Diluted Sequel II DNA Polymerase 2.0 must be used immediately; discard excess.
Add the following reagents, in order, to the PCR tube with the primer-annealed SMRTbell library for a total volume of 13.5 μL:
Sequel Binding Buffer 1.5 μL
DTT 1.5 μL
Sequel dNTP 1.5 μL
Pipette or flick the tube to mix the solution.
Add 1.5 μL of diluted Sequel II DNA Polymerase 2.0 for a total volume of 15 μL.
Pipette or flick the tube to mix the solution.
Briefly spin down the tubes in a mini centrifuge.
Place the sample complex in the thermal cycler and run the following protocol:
30 °C for 4 h
4 °C hold (or hold on ice) until ready for the purification step
Purify of SMRTbell complex
For Sequel
Add 35 μL of Sequel Complex Dilution Buffer to a new LoBind microcentrifuge tube.
Add the total volume (15 μL) of the sample complex to the tube for a total volume of 50 μL.
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.6× of AMPure PB beads to the diluted sample complex for a total volume of 80 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Pipette or flick the tube to mix the solution.
Incubate the tube at room temperature for 5 min.
Place the tube into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 2 min).
While the tube is in the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and discard.
Briefly spin down the tube in a mini centrifuge.
Place the tube back into the magnetic rack.
Remove any remaining supernatant and discard.
Resuspend the beads in 81 μL of Sequel Complex Dilution Buffer immediately.
Pipette or flick the tube to mix the solution.
Incubate at room temperature for 15 min.
Place the tube into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 1 min).
Carefully pipette up the supernatant, transfer to a new LoBind microcentrifuge tube, and place on ice.
On ice, dilute the Sequel DNA Internal Control Complex 3.0 10,000-fold by performing two 100-fold serial dilutions in Sequel Complex Dilution Buffer.
Add 3 μL of the diluted DNA internal control complex and 1 μL of carrier DNA to the tube with the eluted sample for a total volume of 85 μL.
Transfer the sample to a sample plate.
Cover the plate and store at 4 °C or on ice until sequencing.
For Sequel II and IIe
Add 35 μL Sequel Complex Dilution Buffer to a new LoBind microcentrifuge tube.
Add the total volume (15 μL) of the sample complex to the tube for a total volume of 50 μL.
Mix the AMPure PB beads until a homogeneous solution forms.
Add 0.6× of AMPure PB beads to the diluted sample complex for a total volume of 80 μL.
Note: The AMPure PB beads will be viscous; pipette slowly to ensure precise volume.
Pipette or flick the tube to mix the solution.
Incubate the tube at room temperature for 5 min.
Place the tube into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 2 min).
While the tube is in the magnetic rack, slowly remove the supernatant without disturbing the bead pellet and discard.
Briefly spin down the tube in a mini centrifuge.
Place the tube back into the magnetic rack.
Remove any remaining supernatant and discard.
Resuspend the beads in 109.6 μL of Sequel Complex Dilution Buffer immediately.
Pipette or flick the tube to mix the solution.
Incubate at room temperature for 15 min.
Place the tube back into the magnetic rack.
Allow the beads to separate from the liquid and collect on the side of the tube. Wait until the solution appears clear (at least 1 min).
Carefully pipette up the supernatant, transfer to a new LoBind microcentrifuge tube, and place on ice.
On ice, dilute the Sequel II DNA Internal Control Complex 1.0 10,000-fold by performing two 100-fold serial dilutions in Sequel Complex Dilution Buffer.
Add 4 μL of the diluted DNA internal control complex and 1.4 μL of carrier DNA to the tube with the eluted sample for a total volume of 115 μL.
Transfer the sample to a sample plate.
Cover the plate and keep at 4 °C or on ice until sequencing.
Sequencing
For Sequel
Sequence plate on the Sequel system with the following settings:
SMRT Link Run Design > Advanced Options > Immobilization Time > 4 h.
No pre-extension.
Movie collection time: 20 h.
Generate HiFi reads > IN SMRT LINK.
For Sequel II
Sequence on the Sequel II system with the following settings:
SMRT Link Run Design > Advanced Options > Immobilization Time > 4 h.
No pre-extension.
Movie collection time: 20 h.
Generate HiFi reads > IN SMRT LINK.
For Sequel IIe
Sequence on the Sequel IIe system with the following settings:
SMRT Link Run Design > Advanced Options > Immobilization Time > 4 h.
No pre-extension.
Movie collection time: 20 h.
Generate HiFi reads > ON INSTRUMENT.
Data analysis
Sequencing data is processed using SMRTLink according to the SMRTLink User Guide [18] and as described here. High-quality single-molecule circular consensus sequences (CCS or HiFi reads) are generated from the raw data employing ccs. To ensure the integrity of the data, only sequences achieving a minimum of three passes and a mean read accuracy of QV20 are retained.
For Sequel and Sequel II
Process raw subreads with the following steps and settings:
SMRT Link Analysis > Data Utility > Circular Consensus Sequencing > Advanced Options > MinPasses > 3.
Min QV: 20.
For Sequel IIe: CCS are generated on instrument.
Subsequently, the reads are demultiplexed using lima and aligned to the reference genome with pbmm2.
For Sequel and Sequel II
Demultiplex samples with the following settings:
SMRT Link Analyses > Data Utility > Demultiplex > same on both sides.
Align HiFi reads for each sample to the reference with the following settings:
SMRT Link Analysis > One Analysis Per Data Set – Identical Parameters > HiFi Mapping > Reference > GrCH38_no_alts.
For Sequel IIe
Demultiplex samples with the following settings:
SMRT Link Analyses > Data Utility > Demultiplex > same on both sides.
Align HiFi reads for each sample to the reference with the following settings:
SMRT Link Analysis > One Analysis Per Data Set – Identical Parameters > HiFi Mapping > Reference > GrCH38_no_alts.
Single-molecule sequencing reads can be assembled into waterfall plots for visualization of the target region (Figure 2).
Python extractRegion.py [mapped sample] [ref] [coords] > python waterfall.py [motifs].
Figure 2. Example of PacBio Sequel II sequencing output. Waterfall plots of single-molecule sequencing reads. Each horizontal line represents one molecule sequenced, anchored at a non-repeat region 5' to the C9orf72 repeat expansion. The X-axis shows the nucleotide position compared to the anchor. The Y-axis shows the CCS read count. The grey color represents changes in the sequence other than GGGGCC repeat, which are likely to be a sequencing error. (A) NoAmp sequencing of patient iPSC DNA with two repeats on the normal allele (tall blue tail reaching 300 CCS reads) and approximately 250 repeats on the expanded allele. (B) NoAmp sequencing of patient iPSC DNA showing repeat lengths from 2 to 250 representing mosaicism in this line.
The program primrose can be used to predict 5-methylcytosine signatures in HiFi reads using a convolutional neural network applied to multi-pass kinetics data for each read. Methylation data can be further characterized using pb-CpG-tools, which rely on tags generated by primrose for calculating the probability of 5mC methylation. Supplemental Figure 12 of Sachdev et al. [10] shows an example of methylation data obtained using this protocol.
Validation of protocol
This protocol has been used and validated in the following research articles:
Salomonsson et al. [5]. Validated assays for the quantification of C9orf72 human pathology. Sci Rep. 14: 828.
Sachdev et al. [10]. Reversal of C9orf72 mutation-induced transcriptional dysregulation and pathology in cultured human neurons by allele-specific excision. Proc Natl Acad Sci USA. 121(17): e2307814121
General notes and troubleshooting
General notes
Room temperature refers to 20–22 °C.
We have validated multiplexes up to 10 samples but recommend multiplexing a maximum of 5 samples for optimal sequencing coverage.
Before using genomic DNA, allow samples to reach room temperature and invert the tube 20 times to mix.
Use a benchtop cooler to keep all enzymes at -20 °C during the protocol.
Fresh 80% ethanol solutions (stored for up to three days) must be used for all wash steps.
For all enzymatic reactions, use a thermocycler or heat block with a heated lid set to 10 °C above the incubation temperature whenever possible.
sgRNAs and reagents used in sections E, F, and G should be kept on ice at all times.
Let AMPure PB beads and elution buffer reach room temperature prior to use.
The purification steps are necessary to eliminate any unwanted DNA fragments and failed ligation products. The second purification uses a smaller ratio of beads to sample volume (0.42× instead of 0.45×) than the first step, further selecting for larger fragments to enrich on-target reads.
Do not over-dry the AMPure PB beads during purification.
Unless there is a safe pausing point noted, immediately proceed to the next step of the protocol.
Troubleshooting
If genomic DNA samples show smearing (indicating DNA shearing) when run on a 1% gel, collect new samples. Avoiding the vortexing of genomic DNA samples and freeze-thaw cycles can prevent smearing.
If genomic DNA samples have fragment sizes <50 kb, the starting input amount can be increased to generate more reads. If fragments are significantly degraded (fragment sizes <20 kb), collect new samples. Avoiding the vortexing of genomic DNA samples and freeze-thaw cycles can prevent short fragment sizes.
Acknowledgments
C.D.C. is supported by NIH/NINDS K08-NS112330KO8, U19NS132303, Alzheimer’s Association AACSF-17-531484, UCSF CTSI TL1 Fellowship 5TL1TR001871-04, Bright Focus Foundation A20201490F, Carol and Gene Ludwig Award for Early Career Research, Larry H. Hillblom Fellowship, Shupin Fellowship from the UCSF Neurology Endowment, Weill Institute for Neurosciences and UCSF Memory & Aging Center. C.D.C. is grateful to Nancy Sakamoto, Joe DiSabato, the Wolfen Family Foundation and Kathleen D. Mayhew and Mayhew family for their generous support. This protocol was adapted from PacBio’s previous “no-amp targeted sequencing utilizing the CRISPR-Cas9 system.”
Competing interests
J.H. and Y-C.T. are full-time employees at Pacific Biosciences (PacBio) and J.H. holds stock in PacBio. C.D.C. is a founder, with equity, in Ciznor Co., a CNS therapeutics company. The other authors have no conflicts.
Ethical considerations
This protocol uses de-identified patient samples only. It was developed under IRB and approved by the UCSF Human Gamete, Embryo, and Stem Cell Research (GESCR) Committee.
References
DeJesus-Hernandez, M., Mackenzie, I. R., Boeve, B. F., Boxer, A. L., Baker, M., Rutherford, N. J., Nicholson, A. M., Finch, N. A., Flynn, H., Adamson, J., et al. (2011). Expanded GGGGCC Hexanucleotide Repeat in Noncoding Region of C9ORF72 Causes Chromosome 9p-Linked FTD and ALS. Neuron. 72(2): 245–256.
Renton, A. E., Majounie, E., Waite, A., Simón-Sánchez, J., Rollinson, S., Gibbs, J. R., Schymick, J. C., Laaksovirta, H., van Swieten, J. C., Myllykangas, L., et al. (2011). A Hexanucleotide Repeat Expansion in C9ORF72 Is the Cause of Chromosome 9p21-Linked ALS-FTD. Neuron. 72(2): 257–268.
Majounie, E., Renton, A. E., Mok, K., Dopper, E. G., Waite, A., Rollinson, S., Chiò, A., Restagno, G., Nicolaou, N., Simon-Sanchez, J., et al. (2012). Frequency of the C9orf72 hexanucleotide repeat expansion in patients with amyotrophic lateral sclerosis and frontotemporal dementia: a cross-sectional study. Lancet Neurol. 11(4): 323–330.
Iacoangeli, A., Al Khleifat, A., Jones, A. R., Sproviero, W., Shatunov, A., Opie-Martin, S., Morrison, K. E., Shaw, P. J., Shaw, C. E., et al. (2019). C9orf72 intermediate expansions of 24–30 repeats are associated with ALS. Acta Neuropathol Commun. 7(1): 115.
Salomonsson, S. E., Maltos, A. M., Gill, K., Aladesuyi Arogundade, O., Brown, K. A., Sachdev, A., Sckaff, M., Lam, K. J. K., Fisher, I. J., Chouhan, R. S., et al. (2024). Validated assays for the quantification of C9orf72 human pathology. Sci Rep. 14(1): 828.
Bram, E., Javanmardi, K., Nicholson, K., Culp, K., Thibert, J. R., Kemppainen, J., Le, V., Schlageter, A., Hadd, A., Latham, G. J., et al. (2018). Comprehensive genotyping of the C9orf72 hexanucleotide repeat region in 2095 ALS samples from the NINDS collection using a two-mode, long-read PCR assay. Amyotroph Lateral Scler Frontotemporal Degener. 20: 107–114.
Buchman, V. L., Cooper-Knock, J., Connor-Robson, N., Higginbottom, A., Kirby, J., Razinskaya, O. D., Ninkina, N. and Shaw, P. J. (2013). Simultaneous and independent detection of C9ORF72 alleles with low and high number of GGGGCC repeats using an optimised protocol of Southern blot hybridisation. Mol Neurodegener. 8(1): 12.
Ebbert, M. T. W., Farrugia, S. L., Sens, J. P., Jansen-West, K., Gendron, T. F., Prudencio, M., McLaughlin, I. J., Bowman, B., Seetin, M., DeJesus-Hernandez, M., et al. (2018). Long-read sequencing across the C9orf72 ‘GGGGCC’ repeat expansion: implications for clinical use and genetic discovery efforts in human disease. Mol Neurodegener. 13(1): 46.
DeJesus-Hernandez, M., Aleff, R. A., Jackson, J. L., Finch, N. A., Baker, M. C., Gendron, T. F., Murray, M. E., McLaughlin, I. J., Harting, J. R., Graff-Radford, N. R., et al. (2021). Long-read targeted sequencing uncovers clinicopathological associations for C9orf72-linked diseases. Brain 144(4): 1082–1088.
Sachdev, A., Gill, K., Sckaff, M., Birk, A. M., Aladesuyi Arogundade, O., Brown, K. A., Chouhan, R. S., Issagholian-Lewin, P. O., Patel, E., Watry, H. L., et al. (2024). Reversal of C9orf72 mutation-induced transcriptional dysregulation and pathology in cultured human neurons by allele-specific excision. Proc Natl Acad Sci USA. 121(17): e2307814121.
Hafford-Tear, N. J., Tsai, Y. C., Sadan, A. N., Sanchez-Pintado, B., Zarouchlioti, C., Maher, G. J., Liskova, P., Tuft, S. J., Hardcastle, A. J., Clark, T. A., et al. (2019). CRISPR/Cas9-targeted enrichment and long-read sequencing of the Fuchs endothelial corneal dystrophy–associated TCF4 triplet repeat. Genet Med. 21(9): 2092–2102.
Tsai, Y. C., de Pontual, L., Heiner, C., Stojkovic, T., Furling, D., Bassez, G., Gourdon, G. and Tomé, S. (2022). Identification of a CCG-Enriched Expanded Allele in Patients with Myotonic Dystrophy Type 1 Using Amplification-Free Long-Read Sequencing. J Mol Diagn. 24(11): 1143–1154.
PacBio. (2024). Generating PureTargetTM repeat expansion panel libraries.
Fisher, I. J., Salomonsson, S. and Clelland, C. D. (2023). iPSC Cell Culture Protocol. Protocols.io. dx.doi.org/10.17504/protocols.io.36wgqj2n3vk5/v1.
QIAGEN. (2015). QIAGEN® Genomic DNA Handbook.
Thermo Fisher Scientific. (2020). QubitTM 1X dsDNA HS Assay Kits User Guide.
Edvotek. (2020). Quick Guide: SYBR® Safe DNA Stain.
PacBio. (2023). SMRT® Link User Guide v12.0.
Article Information
Publication history
Received: May 13, 2024
Accepted: Jul 18, 2024
Available online: Aug 8, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Molecular Biology > DNA > DNA recombination
Neuroscience > Nervous system disorders
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Endothelin-1-Induced Persistent Ischemia in a Chicken Embryo Model
NK Neha Kumari *
RP Ravi Prakash *
AS Abu J. Siddiqui
AW Arshi Waseem
MK Mohsin A. Khan
SR Syed S. Raza
(*contributed equally to this work)
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5061 Views: 239
Reviewed by: Xiaokang WuIbrahim Alabri Anonymous reviewer(s)
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Abstract
Current ischemic models strive to replicate ischemia-mediated injury. However, they face challenges such as inadequate reproducibility, difficulties in translating rodent findings to humans, and ethical, financial, and practical constraints that limit the accuracy of extensive research. This study introduces a novel approach to inducing persistent ischemia in 3-day-old chicken embryos using endothelin-1. The protocol targets the right vitelline arteries, validated with Doppler blood flow imaging and molecular biology experiments. This innovative approach facilitates the exploration of oxidative stress, inflammatory responses, cellular death, and potential drug screening suitability utilizing a 3-day-old chicken embryo.
Key features
• This model enables the evaluation and investigation of the pathology related to persistent ischemia.
• This model allows for the assessment of parameters like oxidative stress, inflammation, and cellular death.
• This model enables quantification of molecular changes at the nucleic acid and protein levels.
• This model allows for the efficient screening of drugs and their targets.
Keywords: Ischemia Chicken embryo Endothelin-1 Doppler blood flow imaging
Graphical overview
Background
Ischemia, derived from the Greek words "to restrain" and "blood," refers to a critical situation in which insufficient blood supply to an organ or tissue causes nutritional, oxygen, and waste depletion, resulting in cellular and physiological problems [1,2]. It arises from various factors, including vascular obstruction, low blood pressure, or decreased circulation, and significantly contributes to multiple disorders [3–6].
Endothelin-1 (ET-1) is undeniably a powerful vasoactive peptide that is well-known for its capacity to cause blood vessel constriction, which in turn leads to a reduction in the amount of blood flow [7–9]. ET-1's vasoconstrictive properties make it useful in experimental studies to induce ischemia [10–12], helping to explore physiological and pathological responses. While ET-1-induced ischemia is commonly used in in vivo research, it has never been examined in an in ovo model. This study is the first to investigate its effect in ovo.
Doppler blood flow imaging, a crucial tool in ischemic research [13,14], assesses reduced blood flow in various organs like the heart [15], brain [16], and limbs [17]. For instance, in cardiac ischemia, it evaluates coronary artery blood flow [18,19], while in cerebral ischemia, it detects brain artery blood flow changes [20,21]. Furthermore, it detects abnormal blood flow patterns in the context of peripheral vascular and renal ischemia [22] to identify stenosis [23]. Given the utility of this instrument, we incorporated it into our experiment to validate our ischemic model. Additionally, we employed Southern blotting, western blotting, and ELISA for further validation of the model.
Materials and reagents
Biological materials
Fertilized White Leghorn chicken, provided by the Central Avian Research Institute in Bareilly, Uttar Pradesh, India, and reared in our on-campus poultry farm
Reagents
γ-H2AX, 1:500 (Fine Test, catalog number: FNab10441)
HIF-1α, 1:1,000 (Novus Biological, catalog number: NB100-449)
NOX2, 1:500 (Fine Test, catalog number: FNab05804)
IFN-γ, 1:1,000 (Abcam, catalog number: ab133566)
RIPK1, 1:500 (Cloud-Clone Corp., catalog number: MAE640Hu21)
Cleaved caspase-3, 1:500 (ImmunoTag, catalog number: ITT07022)
LC3 l/ll, 1:1,000 (Cell Signaling Technology, catalog number: 4108S)
GAPDH, 1:1,000 (Cloud-Clone Corp.,catalog number: MAB932Hu21)
Anti-mouse IgG, 1:3,000 (Cell Signaling Technology, catalog number: 7076S)
Anti-rabbit IgG, 1:10,000 (Jackson Immuno Research Laboratories, catalog number: 711-035-152)
ELISA Kit, TNF-α (Fine Test, catalog number: EH0302)
ELISA Kit, IL-1β (Fine Test, catalog number: ECH0040)
SDS-PAGE (Invitrogen, catalog number: A25977)
Endothelin-1 (ET-1), 1 pM (Sigma-Aldrich, catalog number: E7764)
N-Acetyl-L-Cysteine (NAC), 100 µM (Sigma-Aldrich, catalog number: A7250)
Solutions
70% ethanol (see Recipes)
1× phosphate buffer saline (PBS) (see Recipes)
Ringer’s solution (see Recipes)
ET-1 (see Recipes)
N-Acetyl-L-Cysteine (NAC) solution (see Recipes)
Recipes
70% ethanol (100 mL)
Combine 30 mL of sterile water with 70 mL of 200 proof/LR grade ethanol to make a final volume of 100 mL.
Reagent Final concentration Amount
Ethanol n/a 70 mL
Distilled Water n/a 30 mL
Total n/a 100 mL
1× phosphate buffer saline (PBS) (100 mL)
To make 100 mL of 1× PBS, combine all reagents with 70 mL of distilled water. Completely dissolve the reagents and adjust to the final volume. Autoclave the solution for 15 min at 121 °C. If needed, adjust the pH to 7.4 by adding a few drops of 0.1 N HCl or 0.1 N NaOH. Divide the solution into 10 mL aliquots and store them at room temperature (RT) in sterile 15 mL centrifuge tubes. Store at RT for up to 12 months.
Reagent Final concentration Amount
Na2HPO4·7H2O 5.37 mM 0.144 g
NaCl 136.8 mM 0.8 g
KCl 26.8 mM 0.2 g
KH2PO4 14.6 mM 0.2 g
H2O n/a 100 mL
Total n/a 100 mL
Ringer’s solution (100 mL)
Mix NaCl, CaCl2, and KCl in 70 mL of sterile distilled water. Adjust the pH to 7.4. Let it dissolve completely and autoclave. Then, filter through a 0.22 µm filter and aliquot into single-use amounts (approximately 10 mL). Store at 4 °C for up to 1–2 months.
Reagent Final concentration Amount
NaCl 123 mM 0.72 g
CaCl2 1.53 mM 0.017 g
KCl 4.96 mM 0.037 g
H2O n/a 100 mL
pH n/a 7.4
Total n/a 100 mL
ET-1
To make a 1 mM stock solution, dissolve 2.49 mg of ET-1 in 1 mL of Milli-Q water. Take 1 µL of the 1 mM ET-1 solution and mix it with 999 µL of Milli-Q water to make a 1 µM concentration solution. To achieve a 1 nM concentration, transfer 1 µL of the 1 µM solution into 999 µL of Milli-Q water. Lastly, dissolve 1 µL of the 1 nM solution in 999 µL of Milli-Q water to achieve a final concentration of 1 pM.
Reagent Final concentration Amount
ET-1 1 mM 2.49 mg
Milli-Q n/a 1 mL
Total 1 mM 1 mL
N-Acetyl-L-Cysteine (NAC) solution
Dissolve 1.63 mg of NAC in 1 mL of Ringer's solution to make a 10 mM stock concentration. Titrate the stock solution to 1 mM by adding 100 µL of the 10 mM NAC solution to 900 µL of Ringer's solution. Further dilute the resulting 1 mM concentration to 100 µM by combining 100 µL of the 1 mM NAC solution with 900 µL of Ringer's solution.
Reagent Final concentration Amount
NAC 10 mM 1.63 mg
Ringer's solution n/a 1 mL
Total n/a 1 mL
Laboratory supplies
Egg incubator (Gentek, catalog number: GL-100)
Pointed sharp curved scissors (Stoelting, catalog number: 52132-11)
Hamilton glass syringe (Hamilton, catalog number: 80087)
Needle 26G–26 × 1/2 (0.45 mm × 13 mm) (Dispovan, catalog number: 30722D
Needle 18G–18G × 1.5 (1.25 mm × 38 mm) (Romsons, catalog number: 13990)
Syringe (5 mL) (Dispovan, catalog number: 653054NE1)
Tube stand (1.5 mL) (TARSONS)
Ocular forceps (Stoelting, catalog number: 52100-52)
Ocular iris (Stoelting, catalog number: 52130-00)
Packaging tape (Sunrise)
Autoclaved tissue roll (Texam Technologies Pvt. Ltd.)
Overhead projection pens (OHP) Marker (Camlin)
Autoclaved bag (TARSONS, catalog number: 550022)
Petri dish (TARSONS, catalog number: 460050)
Egg rack
Sharp waste container (TARSONS, catalog number: 882210)
Centrifuge tube (1.5 mL) (TARSONS, catalog number: 500010X)
Beaker (100 mL) (GLASSCO, catalog number: 23.205.03)
Pasteur pipette (3 mL) (TARSONS, catalog number: 940050)
Equipment
New standardTM stereotaxic rat and mouse surgery apparatus (Stoelting, catalog number: 51500)
Laser Doppler blood flow meter (Moor instruments, catalog number: moorVMS-LDF1)
Stereo zoom microscope (Olympus, catalog number: SZ2-STU3)
Software and datasets
SPSS Statistics Software IBM 29.0
moorVMS-LDF1
GraphPad Prism 5 Software
Procedure
Inducing persistent ischemia
Day 1:
Using tissue paper wipes, clean the 0-day eggs with 70% ethanol. To clean, spray 1 mL of 70% ethanol onto the tissue wipe and then use the dampened wipe to clean the egg (Figures 1 and 2) (see General notes 1, 2).
Figure 1. Arrangement for the sterilization process. First, eggs were placed in the egg rack for the sterilization process. A 70% ethanol solution was used to disinfect the eggshell surfaces. A marker was used for labeling the eggs.
Figure 2. Sterilizing 0-day eggs using 70% ethanol for disinfection. This figure illustrates the process of sterilizing 0-day eggs using 70% ethanol for disinfection. The eggs were arranged in a rack, and 70% ethanol was used to disinfect eggshell surfaces. Eggs were wiped with tissue soaked in 70% ethanol, ensuring that the surface was adequately disinfected.
Use an OHP marker to write the current date and ID number on each egg (Figure 3) (see General notes 3, 4).
Figure 3. Marking eggs with the date. The left-side figure depicts the labeling of eggs on their upper side using a marker. The right-side figure shows the eggs with markings facing downward in the egg rack. This method helps to ensure that the marks remain legible and protected during handling and storage.
Place the egg in the egg incubator with the temperature and humidity maintained at 37 ± 1 °C and 60%–65%, respectively, for 24 h (Figure 4) (see General note 5).
Figure 4. Storing a 0-day egg in a 37 ± 1 °C egg incubator. This image shows an egg incubation setup. The inset images highlight the control panel and humidity meter. The control panel indicates temperature and humidity settings. The setup demonstrates the precision and care required while placing the eggs in the incubator to ensure successful hatching.
Day 2:
Apply a small piece of packaging tape to the egg's edge (approximately 10 mm in length and width) (Figure 5, 6A, 6B).
Figure 5. Requirements for albumin removal. This image depicts the essential tools and equipment required for the albumin removal process. Included in the setup are: a sharps waste container for the safe disposal of sharp instruments, a pointed sharp edge scissors used for precise cutting during the procedure, and a discarder for disposing of non-sharp waste. Additionally, we use an 18G needle to pierce the egg and facilitate albumin extraction, an egg rack to secure the egg during the process, packaging tape to tape the egg, and a 5 mL syringe to accurately measure and extract the albumin. These tools are critical to ensure safety and efficiency in the albumin removal process.
Figure 6. Steps in layering (A–D). (A) Retrieving eggs from the egg incubator set at 37 ± 1 °C to remove the albumin. (B) An illustration of packaging tape applied to the eggshell for albumin removal. (C) Demonstrating the creation of a small hole at the eggshell's edge using pointed scissors. (D) Depiction of the removal of the thin albumin layer from the egg.
Using pointed, sharp, curved scissors, cut a small hole in the eggshell's edge exactly where the packaging tape is located (Figure 6C).
Insert a 5 mL syringe at a 60–70° angle (see General note 6).
Slowly withdraw 5–6 mL of albumin by inserting the needle inside the eggshell and pulling up the required thin albumin (Figure 6D).
After albumin removal, dispose of the syringe in the syringe discarder (see General notes 7–9).
After removing the albumin, cover the needle-piercing area with packaging tape.
Incubate the eggs for 48 h at 37 ± 1 °C in the egg incubator (see General note 10).
Day 3:
After 72 h of incubation, the egg should be removed from the 37 ± 1 °C incubator, and the windowing procedure should be initiated.
Place transparent packaging tape on one side of the eggshell (Figure 7) (see General note 11).
Figure 7. Using packaging tape before creating the window. The depicted figure illustrates the application of transparent packaging or packaging tape onto the eggshell.
Use pointed, sharp, curved scissors to make a small hole in the eggshell at the desired windowing site (Figure 8A); then, carefully cut a circular opening.
Figure 8. Windowing procedure steps. Before beginning the windowing operation, make a hole in the middle of the eggshell on the packaging tape region (A). Remove the eggshells in a spherical shape, ensuring one end remains attached to the shell (B and C). (D) represents an enlarged image of an egg. Take note of the formation of well-organized vitelline arteries () Double-arrow represents the right vitelline artery; (▲) arrowhead represents the left vitelline artery; (*) star represents the site of ischemia; (#) hash represents the placement of the Doppler blood flow meter probe; (+) plus represents the heart; (←) arrow represents the eyeball (D).
Cut an oval-shaped window. This approach is generally known as "windowing" (Figures 8B and C).
Next, locate the right vitelline artery (RVA) using a stereo zoom surgical microscope (see General notes 12–16).
Just above the RVA, place the Doppler flow meter's probe. The probe should be 5 ± 1 mm away from the vasoconstriction area, i.e., toward the distal end of the RVA (Figure 9).
Figure 9. A typical image during ischemia. The figure depicts a Hamilton’s syringe attached to the stereotaxic setup for the controlled release of ET-1 onto the RVA. Black arrowheads indicate the probe.
Once the Doppler probe is fixed to the RVA through the probe holding stand, make sure that the probe wire transfers the laser ray directly onto the RVA. At this point, the Doppler flux readings display the blood flow readings (this is the phase of normoxia).
Next, treat the RVA with 1 µL of ET-1 at a concentration of 1 pM (diluted in Milli-Q, as described in Recipe 4) with the help of a Hamilton syringe (ET-1 was administered at 0 min only once over a period of 8 h) attached to a New standardTM stereotaxic rat and mouse apparatus directly onto the RVA, as shown in Figures 10 and 11A and B. In summary, place the egg within the egg holder and position it directly below the Hamilton syringe, attached to the stereotaxic arm through the packaging tape. The stereotaxic arm allows for unrestricted movement in both horizontal and vertical directions. To administer the ET-1, slowly lower the tip of the Hamilton syringe approximately 2 mm above the intended position of the RVA, and then gradually release 1 µL of ET-1 from the syringe (the site of ET-1 release should be considered the point of occlusion) (see General notes 17–23).
Figure 10. Tools used during the ET-1 treatment procedure
Figure 11. Typical setup for injecting ET-1. ET-1 is administered using a Hamilton's syringe connected to a stereotaxic device that precisely guides the needle onto the artery of interest (A). An enlarged view shows the injection of ET-1 into the artery (B). All experiments used the same needle, Doppler probe, and RVA positions.
After ET-1 therapy, the use of Doppler blood flow meter to track the effects of vasoconstrictions at RT is recommended. Using the stereotaxis device's measurement scale, make sure that the probe is positioned 5 ± 1 mm from the vasoconstriction site. During the persistent ischemic period (a prolonged and continuous restriction or absence of blood supply), the Doppler flow meter should show a significant reduction in flux, reaching up to 70% of the baseline value (Figure 12).
Figure 12. Typical Doppler blood flow flux readings during the ischemia phase. In minute counts, image A1 illustrates the flux transition from normoxia to the ischemia phase. A2 shows the changes in flux over the course of an hour. For up to 8 h, A3 displays flux readings. The black arrow represents flux under normoxia, while the red arrow represents flux measurement under ischemia. We conducted all Doppler blood flow flux experiments at RT following the windowing process. The y-axis represents blood flow flux measured in perfusion units, while the x-axis represents time.
For all embryos, monitor the Doppler blood flow reading until 8 h (see General note 24).
The duration of ET-1 treatment should be maintained from the onset of ischemia until 8 h (a persistent ischemic period, a non-reperfusion phase) (the period of no restoration of blood flow to RVA). This suggests that the treatment induces a persistent ischemic effect as evidenced by a 70% reduction in flux intensity from its baseline level (Videos 1 and 2 show the embryo's arterial system prior to and 8 h after ET-1 injection, respectively). After the ischemic procedure, carefully apply a few drops (two to three) of 1× PBS to the embryo using a 10 mL plastic Pasteur tube (Figure 13) (see General notes 25–27).
Video 1. Arterial system of 3-day-old chicken embryo before injecting ET-1
Video 2. Arterial system of 3-day-old chicken embryo after 8 h of injecting ET-1
Figure 13. Adding 1× PBS to prevent dehydration. This figure illustrates the periodic (at 2 h intervals) addition of a few drops of 1× PBS to prevent embryo desiccation during the ischemic period.
Following an 8 h ischemia period, proceed with RVA excision according to the protocol specified for drug treatment and downstream studies (see General note 28).
After 8 h of ischemia with ET-1, remove the eggshells and proceed with the designated treatment protocol.
Place the egg gently on a 90 mm Petri dish (see General note 29).
After placing the embryo on the Petri dish, locate and collect the RVA under the supervision of a stereo zoom surgical microscope (Figure 14B). Make sure the RVA's excision is up to 15 mm ± 1 mm (distal from the trunk), 5 mm ± 1 mm on each of the artery's left and right sides, and 2 mm ± 1 mm toward the trunk (Figure 14C). Measure the area that needs removal using a ruler (optional). Measure all distances from the point of occlusion.
Figure 14. Steps of RVA Isolation. To isolate the RVA, remove the eggshell (A). (B) shows the transfer of an egg embryo into a 90 mm Petri dish, followed by the isolation of the RVA using an ocular iris under a stereo zoom microscope. (C) shows a schematic image of the right side of a chicken embryo, illustrating the region of tissue excision around the RVA (for downstream studies). The annotation "1" represents the eye, and "2" represents the heart. The half-black circular line denotes the RVA's emergence from the embryo's thorax. The vertical line's asterisk indicates the location of the laser Doppler blood flow device installation. The intersection (in black lines) denotes the occlusion site. 1× PBS was used to wash RVA (D). RVA is collected and placed in a 1.5 mL centrifuge tube with 500 µL of Ringer's solution (E). The accompanying treatment process involved placing the RVA in a 37 °C laboratory incubator and incubating it for 4 h (F).
Rinse the excised RVA once in a sterile Petri dish containing 1× PBS (Figure 14D) before proceeding to the post-excision incubation step.
A control should be provided with a similar treatment to the ET-1+NAC-treated RVAs but no NAC.
Place the excised artery in a 1.5 mL centrifuge tube containing 500 µL of Ringer's solution, both with and without the selected drug, using ocular forceps (Figure 14E).
Subsequently, place the individual tubes containing the excised RVA, with or without the drug, in a laboratory incubator at 37 ± 1 °C for 4 h without agitation (Figure 14F) (see General note 30).
Following the 4 h incubation treatment, carefully remove the RVA from the tube with ocular forceps and rinse it twice in two consecutive tubes containing 500 µL of 1× PBS, each for 30 s (see General note 31).
Remove the tube from the incubator post-incubation and move on to process the artery for subsequent research tasks, such as Western blotting, ELISA, etc. To delicately remove the artery, use ocular forceps on aluminum foil. Subsequently, weigh the artery for protein estimation in preparation for downstream studies.
DNA isolation from RVAs
Crush 10 mg of tissue in 1 mL of absolute alcohol (see General note 32).
Transfer the homogenate into a fresh 1.5 mL centrifuge tube (see General note 33).
Centrifuge at 12,000× g for 10 min at 4 °C. Discard the supernatant.
Next, add 500 µL of lysis buffer (Table S1) containing proteinase K to the pellet. Resuspend and incubate at 55 °C for 2 h (see General note 34).
Next, centrifuge at 12,000× g for 10 min at 4 °C.
Transfer the supernatant to a fresh 1.5 mL centrifuge tube. Discard the pellet (see General note 35).
Next, add 200 µL of 3 M chilled sodium acetate to the sample. Incubate on ice for 15 min. Centrifuge at 12,000× g for 10 min at 4 °C.
Transfer the supernatant to a fresh 1.5 mL centrifuge tube.
Add an equal volume of phenol:chloroform:isoamyl alcohol, 25:24:1 (PCI) (see General note 36).
Mix properly by inverting the tube 15–20 times. Centrifuge at 12,000× g for 10 min at 4 °C.
Collect the aqueous phase.
Add an equal volume of isopropanol. Incubate for 30 min at 4 °C.
Centrifuge at 12,000× g for 10 min at 4 °C. Discard the supernatant.
Wash the pellet with 75% ethanol.
Dry the pellet properly (see General note 37).
Dissolve the pellet in 50 µL of TE buffer.
Finally, use a NanoDrop spectrophotometer to quantify the isolated DNA.
Agarose gel electrophoresis
Put a middle mold in the gel holder.
To prepare a 0.2% agarose gel, dissolve 0.14 g of agarose in 70 mL of 1× TAE buffer. To make the 1× TAE buffer, combine 20 mL of 50× TAE with enough distilled water to make a total volume of 1 L.
Heat it in the microwave for 3–4 min at a power of 300 watts; stir it occasionally.
Let it cool down to 50 °C and then add 14 µL of ethidium bromide.
Pour the agarose mixture into the mold, fill it with a comb, and let it solidify for 20 min.
After removing the gel from the comb and holder, place it in the electrophoresis chamber, and add enough 1× TAE buffer to cover the gel with a 2 mm layer of buffer.
Place the gel sample and the 3 µL marker into the agarose gel combs.
Run the gel for 1–2 h at 50 V (see General note 38).
Finally, analyze the gel in the Gel Doc system.
Western blotting
Preparation of the sample: In a lysis buffer containing protease and phosphatase inhibitors, homogenize RVA samples (see General note 39).
Next, maintain constant agitation for 30 min at 4 °C and isolate the protein using a high-speed centrifuge at 16,000× g for 30 min.
Place the tubes on ice after gently removing them from the centrifuge, collect the supernatant, transfer it to a new tube, and discard the pellet.
Next, determine the protein concentration using the Pierce BCA protein assay kit (see General note 40).
To prepare the resolving gel (Table S2), mix the constituents (acrylamide, resolving buffer, Milli Q, ammonium persulphate, and tetramethylethylenediamine) according to the desired gel percentage resulting in a total volume of 8 mL (see General note 41).
To prepare the stacking gel (Table S3), mix the constituents (acrylamide, stacking buffer, Milli Q, ammonium persulphate, and tetramethylethylenediamine) resulting in a total volume of 3 mL.
Load 20–30 μg of total protein and Laemmli buffer (for 5×). Then, incubate each sample for 5 min at 95–100 °C.
Subsequently, load an equivalent protein amount onto a commercially available SDS-PAGE gel and conduct the gel electrophoresis (Table S4) (run the gel at about 70 V) until the protein band nearly reaches the bottom of the gel (see General notes 42, 43).
Next, using wet transfer equipment, transfer proteins from the gel to a PVDF membrane.
For 2 min, place the gel in 1× transfer buffer (Table S5) to equilibrate; remove excess SDS, exchange buffers, and ensure better contact with the transfer membrane for efficient protein transfer.
Activate the PDVF membrane in methanol for 3–5 min prior to transfer.
Next, assemble the transfer sandwich, ensuring there are no trapped air bubbles.
Place the cassette in the transfer tank, along with an ice pack, and transfer at 90 V for 100 min (see General note 44).
To avoid non-specific antibody binding, soak the PVDF membrane in a 5% blocking solution for 1 h (see General note 45).
Subsequently, transfer the membrane to a refrigerator set at 4 °C and allow it to undergo overnight incubation in the presence of primary antibodies (see General notes 46, 47).
The next day, commence the washing procedure by thoroughly rinsing the membrane three times with 1× TBS-T (Table S6), each rinse lasting 3 min.
Next, incubate the membrane for 1 h at RT with secondary antibodies.
Next, rinse the membrane three times, for 3 min each (see General note 48).
Next, incubate the membrane with ECL for 10–15 s (see General note 49).
Using a Chemidoc (see General notes 50, 51), capture the image.
Using image analysis software, analyze the captured image to quantify band intensities and compare expression levels between samples. Evaluate the results in comparison with controls included in the experiment (e.g., GAPDH) for validation and interpretation (see General note 52).
ELISA
Place the artery on ice. Proceed to wash the tissue using the pre-cooled 1× PBS buffer (0.01 M, pH = 7.4).
Use lysate to grind artery homogenates on ice.
Centrifuge the homogenates at 5,000× g for 5 min. Next, collect the supernatant.
For data analysis, determine total protein concentration by BCA kit (taking reagent B:A in a 1:50 ratio) and add 190 µL to each well. Usually, the total protein concentration for the ELISA assay should be within 1–3 mg/mL.
Next, place the sample (10 µL) and control (blank) wells on the pre-coated plate and measure the samples. We recommend measuring each standard and sample in duplicate to reduce experimental errors.
In each standard well, aliquot 100 µL of stock solution on a dilution of ½ into five consecutive tubes. Additionally, add 100 µL of sample dilution buffer to the control (blank) well. Then, add 100 µL of test samples to each sample well. Seal the plate and incubate it for 90 min at 37 °C. Add the solution to the bottom of each well. Gently combine the mixture without touching the sidewall.
Allow the liquid to absorb in the plate or tap the plate on clean absorbent paper two or three times. Without immersion, add 350 µL of wash buffer to each well. Discard the liquid in the well and tap on the absorbent paper again. Repeat the washing steps twice.
In each well, add 100 µL of biotin-labeled antibody working solution. Seal the plate and incubate it for 60 min at 37 °C.
Repeat step E7.
In each well, add 100 µL of HRP-Streptavidin conjugate working solution. Seal the plate and incubate for 30 min at 37 °C.
Next, proceed to wash the plate five times using the wash buffer.
Add 90 µL of 3,3',5,5'-Tetramethylbenzidine substrate into each well, seal the plate, and incubate at 37 °C in the dark for 10–20 min.
After staining, keep the liquid in the well. Add 50 µL of stop solution to each well. The color will turn yellow immediately.
Read the absorbance at 450 nm in a microplate reader immediately and calculate.
Data analysis
The data distribution was checked by the Shapiro-Wilk test of normality. A one-way analysis of variance (ANOVA) with the Mann-Whitney test was used to compare the differences between the two groups and Tukey’s test was used to analyze the data for multiple comparison using SPSS Statistics 29.0 software (IBM). Statistics were judged significant at 0.001 (*/**/***P < 0.05/0.01/0.001 compared to the control/ET-1-treated group; #/##/###P < 0.05/0.01/0.001 compared to the ET-1/ET-1+NAC-treated group). The error bars represent the mean ± SD, at least n = 4.
Validation of protocol
This protocol aims to ascertain the success and reproducibility of ET-1-induced persistent ischemia in 3-day-old chicken embryos. Standard laboratory procedures employed in ischemic research globally were used, including Doppler blood flow imaging measurements [24,25], gel electrophoresis [26,27], ELISA [28,29], and western blotting [30] for model validation. In brief, first, we assess the model's efficacy via Doppler blood flow imaging, a well-established technology for detecting blood flow [31,32]. The comparative study began by comparing data from the control group to the ET-1 groups (Figure 15) (see General note 53).
Figure 15. Overlay graph comparing control vs. ET-1-treated ischemic RVAs. The ET-1 treatment demonstrates a significant reduction in flux overlay when compared to values observed in control-treated RVAs. The statistical analysis reveals a highly significant difference (Control vs. ET-1 ***P < 0.001) across all time points. The Shapiro-Wilk test of normality was used to check the data distribution. We used SPSS Statistics 29.0 software (IBM) to perform a one-way analysis of variance (ANOVA) with the Mann-Whitney test to compare the differences between the two groups. Statistics were judged significant at 0.001 (***P < 0.001 compared to the control/ET-1-treated group). The error bars represent the mean ± SD, n = 10.
This study confirmed ET-1's ability to induce persistent ischemia in 3-day-old chicken embryos. Subsequent molecular analyses included Southern blotting [33] (Figure 16A), ELISA (for TNF-α [34–36] and IL-1β [37,38]), and western blotting (for γ-H2AX [39,40], HIF-1α [41,42], NOX2 [43,44], IFN-γ [45,46], RIPK1 [47,48], cleaved caspase-3 [49,50], and LC3 [51,52] expression) (Figure 16B). Along the preceding lines, the model's efficiency in exploring hypoxia (Figure 17), oxidative stress [53,54] (Figure 18), inflammatory stress [55,56] (Figure 19A–C), necrosis [57], apoptosis [58] (Figure 20A), and autophagy [59] (Figure 20B) pathways was in line with the previous findings. Notably, pharmacological treatment with NAC in the ET-1 group outperformed ET-1 treatment alone [60], indicating the model's applicability for drug screening studies. In conclusion, our model has proven useful for analyzing persistent ischemia processes and has the potential to be used in research into tissue injury, inflammation, cell death, and drug screening applications.
Figure 16. Effect of prolonged ischemia, mediated by ET-1, on DNA damage. We used agarose gel electrophoresis to assess the DNA damage in 3-day-old chicken embryos treated with ET-1. The results confirmed the degradation of genomic DNA in ET-1 samples. NAC, an inhibitor of antioxidant therapy, significantly reduced DNA damage. A 1 kb DNA ladder was utilized as a size marker (A). Next, we employed western blot analysis to quantitatively analyze the expression of γ-H2AX in the presence and absence of NAC to further examine the ischemic stress reaction on DNA damage (B). The treatment with ET-1 significantly increased the expression of the DNA damage marker γ-H2AX. Notably, the use of NAC greatly reduced the reported effects. As an internal control, GAPDH was used (**P < 0.01 compared to the control/ET-1-treated group; #P < 0.05 compared to the ET-1/ET-1+NAC-treated group). Error bars represent the mean ± SD, n = 4.
Figure 17. Influence of ET-1-induced ischemia on HIF-1α expression. We used western blotting to look at the expression levels of the hypoxia marker HIF-1α to see whether ET-1-induced ischemia affected hypoxia or not. The results showed an increase in HIF-1α expression after ET-1 therapy; the use of NAC greatly reduced the reported effects, thus confirming ischemic development. GAPDH was used as an internal control (*P < 0.05 compared to the control/ET-1-treated group; #P < 0.05 compared to the ET-1/ET-1+NAC-treated group). Error bars represent the mean ± SD, n = 4.
Figure 18. Effect of persistent ischemia on oxidative stress. To assess the oxidative stress response during persistent ischemia, we used western blot analysis to look at the levels of NOX2 expression in the presence and absence of NAC. ET-1 treatment dramatically raised the expression of oxidative stress markers, indicating a considerable increase in oxidative stress in chicken embryos. Notably, the administration of NAC, an antioxidant, significantly reduced the observed effects. GAPDH was used as an internal control (*P < 0.05 compared to the control/ET-1-treated group; #P < 0.05 compared to the ET-1/ET-1+NAC-treated group). Error bars represent the mean ± SD, n = 4.
Figure 19. Effect of persistent ischemia on inflammation. To assess the inflammatory response after chronic ischemia, we used western blot analysis to examine IFN-γ expression levels in the presence and absence of NAC (A). We also used ELISA to evaluate TNF-α and IL-1β levels in the presence and absence of NAC (B and C, respectively). The expression of these inflammatory markers was dramatically increased by ET-1 treatment, indicating a significant rise in inflammation in embryos. GAPDH was used as an internal control (*/**P < 0.05/0.01 compared to the control/ET-1-treated group; #/##P < 0.05/0.01 compared to the ET-1/ET-1+NAC-treated group). Error bars represent the mean ± SD, n = 4.
Figure 20. Impact of persistent ischemia on cell death pathways. (A) depicts necrosis and apoptosis. By measuring the levels of RIPK1 expression, we first looked into the activation of prolonged cell death due to insufficient blood supply. We examined it both with and without NAC (Aii). The results showed that RIPK1 expression went up after ET-1 treatment, which suggests that ET-1 treatment induces necrosis. In addition, administering NAC reversed the observed effects, providing evidence for the involvement of oxidative stress. Through examining cleaved caspase-3 expression, our investigation focused on the induction of apoptosis following the administration of ET-1 (Aiii). ET-1 treatment increased the expression of cleaved caspase-3. (B) Subsequently, we analyzed the levels of LC3II/I expression to evaluate autophagy activation in response to prolonged ischemia. The administration of ET-1 markedly increased LC3II expression, while NAC therapy reduced it. The internal control utilized in the experiment was GAPDH (**/***P < 0.01/0.001 compared to the control/ET-1-treated group; #/##/###P < 0.05/0.01/0.001 compared to the ET-1/ET-1+NAC-treated group). Error bars represent the mean ± SD, n = 4.
General notes and troubleshooting
Eggs that are either newly laid (0-day) or stored for around 15 days under refrigeration can be employed. Older eggs might not successfully develop into embryos. Eggs that are freshly laid can be refrigerated at temperatures ranging from 15 to 16 °C for about two weeks. However, opting for a recently laid egg can enhance embryo viability and development potential.
Sterilization of eggs is recommended, as it will assist in removing any germs or microbes on the eggshell (Figures 1 and 2).
It is important to write the dates on the egg because it facilitates identification.
Once each egg has received a date and an ID, arrange the eggs on the egg rack in such a way that the written portions are at the bottom. This technique greatly aids in determining the embryo's orientation during windowing.
It is critical to set the egg incubator's temperature and humidity at 37 ± 1 °C and 60%–65%, respectively, as deviations from these optimal limits may result in inadequate or no growth.
A 5 mL syringe and an 18 G needle are recommended to facilitate the removal of albumin.
Albumin removal is necessary for all eggs, including both control and experimental eggs.
Albumin removal prevents its overspill while creating a window besides providing a development bed for the embryo. Further, the removal of albumin reduces the possibility of embryo injury during windowing because it prevents the embryo from sticking to the membrane adjacent to the inner surface of the shell.
To prevent damage to the yolk, insert the 18 G needle at a 60–70° angle. Puncturing the yolk significantly reduces egg survival. Hence, when removing albumin, it is essential to move slowly and concentrate on the thin outer layer. Extracting the viscous albumin can cause the embryo to rotate or change position. Obtaining dense albumin is a critical step that has the potential to significantly alter the embryo's position, emphasizing the importance of meticulous precision in procedure execution. By accounting for complex factors that influence embryo behavior, they enhance scientific accuracy and increase the likelihood of desired results. Recognizing and incorporating these insights is integral to a comprehensive and effective protocol implementation.
All eggs, including both control and experimental eggs, will undergo incubation.
A transparent packaging tape will help in reducing unwanted breakage of the shell during the cutting of the eggshell in an oval shape, besides helping in reducing the free fall of pieces of eggshell onto the embryo bed.
An oval-shaped window cutting provides a more stable platform for subsequent procedures.
Make sure the oval cut is large enough to provide straightforward access to the embryo from any direction. If necessary, adjust the egg's location to accommodate the embryo's position.
Use considerable caution during the surgical operation to prevent injuring the RVA or any neighboring arteries. If the RVA sustains damage by accident, remove the egg from the study.
A well-developed embryo should be visible at 72 h (Figure 8D).
All eggs, encompassing both control and experimental eggs, will go through the process of egg windowing.
For the experiment, precise standardization of ET-1 dosage and timing is critical. Following the settings outlined in this protocol, a concentration of 1 pM at the 8 h mark should yield similar results as indicated in the protocol.
Do not inject more than 1 µL of ET-1; doing so can cause the excess ET-1 to spill over in an unwanted area or across the embryo's extremes or ends, which would cause the embryo to die.
To maintain the integrity of the data, any eggs that were inadvertently exposed to more than 1 µL of ET-1 were promptly excluded and eliminated from the study. The Hamilton syringe, equipped with gradations, allows for precise dispensing of ET-1 dissolved in MilliQ.
Investigations should only consider embryos that exhibit a flux reading drop of at least 70% post-ET-1 treatment.
Although minor fluctuations may occur during normoxia, they are significantly less noticeable than the sharp decline observed after ET-1 treatment. Importantly, these minor fluctuations will not impact the statistical calculations when comparing the normoxia and ischemic phases.
The precision required for ET-1 injection and embryo handling may hinder reproducibility.
During incubation, it is critical to maintain constant humidity and temperature; any changes may impact the embryo's growth and the induction of ischemia.
Precise positioning of the Doppler probe (Figure S1) is essential for reliable blood flow measurements; even minor deviations can lead to conflicting results.
ET-1 after an 8 h period could result in mortality escalation of roughly 35%–40% until 12 h, with death rates increasing to 60%–70% within 24 h of ET-1 administration. As a result, it is best to conduct investigations within 8 h (Figure S2). The ET-1 used in these trials is 1 µL of 1 pM.
Use a 10 mL plastic Pasteur tube to apply 2–3 drops of 1× PBS every 2 h during ET-1 treatment until the experiment is over to maintain the embryo's moisture and prevent dehydration.
The standardized composition, buffering capacity, well-defined ionic composition, and osmolality of 1× PBS as an isotonic solution contribute to maintaining a stable pH environment and preventing osmotic stress, ensuring consistent conditions for proper embryonic development.
The process for control eggs will be the same as for ischemic eggs but without ET-1 treatment.
To remove the eggshell without disturbing the yolk or embryo, a precise incision is made in the middle of the pre-existing oval-shaped window cut on the shell using sterilized scissors (Figure 14A). Then, the upper region of the eggshell is carefully cut, allowing the contents of the egg to be gently poured into a dish. Throughout this process, the utmost care is taken to avoid any unnecessary shaking or disturbance, ensuring a smooth transfer of the yolk and embryo with minimal disruption.
Regardless of whether the excised RVA has undergone drug incubation, immerse all RVAs in Ringer's solution, the standard control for drug interventions, within a laboratory incubator.
At this point, the experiment can proceed as specified or, alternatively, the artery can be directly frozen at -80 °C for future use.
To achieve effective homogenization, ensure that the tissue is thoroughly and consistently crushed with absolute alcohol.
Avoid disturbances during supernatant removal. Repeat the centrifugation and washing processes, modifying the parameters as necessary, if the supernatant removal is not sufficient.
Ensure thorough resuspension and consistent incubation.
Check for efficient supernatant transfer to a new tube, while ensuring minimal pellet contamination. If the supernatant is not sufficiently washed, repeat the centrifugation and washing processes, modifying parameters as necessary.
To ensure successful DNA isolation, confirm the addition of PCI solution in equal volume and thorough mixing by inversion (15–20 times). Avoid insufficient mixing.
It is critical to dry the pellet properly. Methods for overcoming the problem include extended centrifugation, air drying, vacuum concentration, and optimizing the washing conditions by changing the ethanol content.
Consistently monitor the run time and voltage to prevent overheating and ensure proper separation of DNA fragments.
Ensure thorough homogenization in the lysis buffer with protease and phosphatase inhibitors to extract a representative protein sample.
Verify protein concentration using a protein assay and ensure samples are properly prepared and denatured.
Use freshly prepared or properly stored gels. Ensure there are no air bubbles in the gel during polymerization.
To guarantee appropriate separation during electrophoresis, it is imperative to load a precise and comparable quantity of protein onto the gel.
Verify the composition of the running buffer and ensure that the gel runs at the proper time and voltage.
To achieve optimal transfer without excessive heating, precise timing and voltage adjustments are required. During the transfer procedure, use an ice pack to preserve the transfer unit's low temperature.
Use an appropriate blocking agent (e.g., 5% BSA or non-fat dry milk) for sufficient time to reduce non-specific binding.
Optimizing incubation temperature and duration for specific antibodies to ensure effective binding is equally important in the current protocols.
Antibody concentration and specificity are crucial for accurate detection.
Thorough washing is essential to eliminate surplus secondary antibodies and minimize background. If the current washing is insufficient, consider increasing the number of washes or extending the washing process's duration.
Use fresh and high-quality enhanced chemiluminescence (ECL) reagents. Optimize exposure time to avoid underexposure.
Capture the image promptly using a Chemidoc to avoid signal decay and ensure accurate documentation.
If bands are still not visible, repeat critical steps (e.g., transfer, blocking, and antibody incubation) to identify potential errors.
Use a loading control (e.g., β-actin or GAPDH) to confirm equal protein loading across lanes.
To ensure the reproducibility and accuracy of model validation, the following key markers can be used: γ-H2AX for double-stranded DNA breaks; HIF-1α for hypoxia; ORP150, NOX2, and SOD1 for oxidative stress; TNF-α, NF-kβ, and IFN-γ for inflammatory stress; RIPK1 for necrosis; cleaved caspase-1, cleaved caspase-3, Apaf-1, Bax, and BCl2 for apoptosis; LC3, p62, Beclin1, Ambra1, LAMP1, and Cathepsin B for autophagy.
Acknowledgments
The authors thank the Department of Science and Technology-Science Engineering Research Board (Grant No. YSS/2015/01731), New Delhi, for providing the funds to purchase a Doppler blood flow meter. Arshi Waseem is thankful to the Uttar Pradesh Council of Science and Technology for providing a junior research assistant fellowship (Project ID 2837, reference no. CST/D-1544). We thank Prof. Joanki Sen of the Indian Institute of Technology's Bioscience and Bioengineering department for critically examining the article and providing input.
Competing interests
The authors declare no competing interest.
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Supplementary information
The following supporting information can be downloaded here:
Figure S1. An illustrative schematic of the Doppler blood flow imaging system, accompanied by annotations
Figure S2. Picture showing the arterial system of a 3-day-old chicken embryo before injecting ET-1 and at 1 h intervals until 8 h of ischemia
Table S1. Lysis buffer
Table S2. Resolving gel preparation
Table S3. Stacking gel preparation
Table S4. 1× running buffer
Table S6. 10× TBS-T buffer
Table S5. 1× transfer buffer
Article Information
Publication history
Received: May 21, 2024
Accepted: Jul 24, 2024
Available online: Aug 9, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Medicine > Cardiovascular system
Cell Biology > Tissue analysis > Tissue imaging
Cell Biology > Model organism culture > Embryo selection
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5,062 | https://bio-protocol.org/en/bpdetail?id=5062&type=0 | # Bio-Protocol Content
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Multiplexed Microfluidic Platform for Parallel Bacterial Chemotaxis Assays
MS Michael R. Stehnach *
RH Richard J. Henshaw *
SF Sheri A. Floge
JG Jeffrey S. Guasto
(*contributed equally to this work)
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5062 Views: 458
Reviewed by: Olga KopachHsih-Yin Tan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jul 2023
Abstract
The sensing of and response to ambient chemical gradients by microorganisms via chemotaxis regulates many microbial processes fundamental to ecosystem function, human health, and disease. Microfluidics has emerged as an indispensable tool for the study of microbial chemotaxis, enabling precise, robust, and reproducible control of spatiotemporal chemical conditions. Previous techniques include combining laminar flow patterning and stop-flow diffusion to produce quasi-steady chemical gradients to directly probe single-cell responses or loading micro-wells to entice and ensnare chemotactic bacteria in quasi-steady chemical conditions. Such microfluidic approaches exemplify a trade-off between high spatiotemporal resolution of cell behavior and high-throughput screening of concentration-specific chemotactic responses. However, both aspects are necessary to disentangle how a diverse range of chemical compounds and concentrations mediate microbial processes such as nutrient uptake, reproduction, and chemorepulsion from toxins. Here, we present a protocol for the multiplexed chemotaxis device (MCD), a parallelized microfluidic platform for efficient, high-throughput, and high-resolution chemotaxis screening of swimming microbes across a range of chemical concentrations. The first layer of the two-layer polydimethylsiloxane (PDMS) device comprises a serial dilution network designed to produce five logarithmically diluted chemostimulus concentrations plus a control from a single chemical solution input. Laminar flow in the second device layer brings a cell suspension and buffer solution into contact with the chemostimuli solutions in each of six separate chemotaxis assays, in which microbial responses are imaged simultaneously over time. The MCD is produced via standard photography and soft lithography techniques and provides robust, repeatable chemostimulus concentrations across each assay in the device. This microfluidic platform provides a chemotaxis assay that blends high-throughput screening approaches with single-cell resolution to achieve a more comprehensive understanding of chemotaxis-mediated microbial processes.
Key features
• Microchannel master molds are fabricated using photolithography techniques in a clean room with a mask aligner to fabricate multilevel feature heights.
• The microfluidic device is fabricated from PDMS using standard soft lithography replica molding from the master molds.
• The resulting microchannel requires a one-time calibration of the driving inlet pressures, after which devices from the same master molds have robust performance.
• The microfluidic platform is optimized and tested for measuring chemotaxis of swimming prokaryotes.
Keywords: Microfluidics Microfabrication Chemotaxis High-throughput screening Bacteria Serial dilution Gradient generation Photolithography Soft lithography Polydimethylsiloxane (PDMS)
Graphical overview
Background
Many motile microorganisms use chemotaxis to navigate complex environments by detecting and responding to gradients in chemical stimuli [1–3]. Chemotaxis mediates many key biological processes including reproduction [4], microbial foraging [5], and infection [6]. Over the past two decades, microfluidics have been established as a crucial tool for studying chemotaxis [7,8] by enabling precise control of the chemical conditions experienced by swimming cells [9]. Applications have included modeling nutrient patches [5], precise and systematic variation of concentration gradients [10], drug dose–response quantification [11], and infectious disease diagnostics [12]. Microfluidic chemotaxis assays inherently rely on diffusion and device geometry to establish well-defined and reproducible chemical gradients. Passive devices [8,13,14] use relatively large reservoirs of chemoattractant loaded into micro-wells to produce long-lasting spatial gradients. Many gradient generators [15–17] stratify chemostimulant and buffer solutions via laminar flow patterning to establish steady gradients under continuous flow or slowly evolving gradients via diffusion in a stop-flow device. Porous membranes and hydrogel microfluidics produce steady-state gradients without subjecting microbes to an external flow [9,18]. Each of these methods produces tunable gradients to quantify aspects of microbial chemotaxis. However, there is a trade-off between high-throughput measurement of population-level cell responses to a diverse range of chemostimulants across a range of concentrations (e.g., micro-well techniques) versus high-spatiotemporal resolution of cell behavior within a defined single chemical gradient (e.g., gradient generators).
Here, we present the multiplexed chemotaxis device [19] (MCD). This high-throughput microfluidic platform capitalizes on the parallelization afforded by microfluidics [11,12,20] to conduct simultaneous microbial chemotaxis screening across gradient conditions spanning the microbe’s entire sensitivity range. The MCD comprises a two-layer architecture: (i) a serial dilution layer [11] produces five logarithmically diluted chemostimulant solutions (plus a control) and (ii) a cell injection layer introduces microbes and buffer. The MCD performs six stop-flow diffusion assays simultaneously on a single chip, where the performance has been fully characterized, validated, and benchmarked against conventional single gradient generator assays [19]. The serial dilution ratios are robust and customizable during the design stage. Compared to existing microfluidic techniques, the MCD has significantly higher throughput by enabling the simultaneous measurements of chemotactic responses across a range of concentration gradient conditions. This methodology has recently been applied to rapidly screen bacterial chemotaxis toward a wide range of identified metabolites produced by phage-infected cyanobacteria [21].
The MCD limitations are derived from its relative structural complexity. Whilst typical microfluidic devices consist of a single layer that can be fabricated in almost any microfabrication facility, the dilution layer of the MCD requires a mask aligner. Care must be taken with the alignment of the two PDMS layers, as a misalignment could render the device inaccurate or inoperable. Different choices of dilution ratios require a new dilution layer geometry fabricated to new design specifications. Finally, in the current mode of operation, each observation chamber is imaged once every few seconds due to limitations in automated microscopy. Whilst typically insufficient to capture bacteria motility dynamics, this can be mitigated by periodically capturing a short video of each individual assay at the cost of decreasing the sampling frequency across the MCD. Other potential applications of the MCD include but are not limited to accelerating microfluidic approaches to human health studies [22,23], chemical synthesis and drug discovery [24], and studying protists and antibiotic-resistant bacteria [25].
Materials and reagents
Reagents
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906)
Fluorescein sodium salts (fluorescein) (Sigma-Aldrich, catalog number: F6377)
Polydimethylsiloxane mixture (PDMS) (SYLGARDTM 184) (Sigma-Aldrich, catalog number: 761036)
SU-8 3050 photoresist (Kayaku Advanced Materials)
SU-8 2025 photoresist (Kayaku Advanced Materials)
Tridecafluoro-1,1,2,2-tetrahydrooctyl trichlorosilane (silanization agent) (Gelest Inc., catalog number: SIT8174.0)
Propylene glycol methyl ether acetate (photoresist developer) (Sigma-Aldrich, catalog number: 484431)
Ethanol (VWR, CAS number: 64-17-5)
Acetone (Sigma-Aldrich, CAS number: 67-64-1)
Isopropyl alcohol (Sigma-Aldrich, CAS number: 67-63-0)
Deionized (DI) water
Buffer solution cell resuspension/dilution. For example, a minimal motility medium or artificial seawater (ASW) as appropriate. The choice of media is organism-dependent; for example, the commonly used M9 minimal media for bacteria [26]. In the case of marine microbes, two common examples include artificial seawater amended with Provasoli’s ES solution (ESAW) [27,28] or the commercially available Instant Ocean® sea salt
Solutions
Polydimethylsiloxane (PDMS) mixture (see Recipes)
BSA solution (see Recipes)
Fluorescein stock solution, 1 mM (see Recipe)
Fluorescein stock solution, 0.1 mM (see Recipes)
Recipes
Polydimethylsiloxane (PDMS) mixture
Note: A 10:1 mass ratio of elastomer base to curing agent was used. The total mass of each component depends on the open volume on the mold.
Reagent Final concentration Quantity or Volume
SYLGARDTM 184 silicone elastomer base 90.9% wt 30 g
SYLGARDTM 184 silicone elastomer curing agent 9.1% wt 3.0 g
Total (optional) n/a 33 g
BSA solution
Note: Filter mixture through a 0.2 μm syringe filter after mixing and store at 4 °C. Preferably use within two weeks.
Reagent Final concentration Quantity or Volume
BSA n/a 0.1 g
DI water n/a 20 mL
Total (optional) 0.5% w.v. 20 ml
Fluorescein stock solution (1 mM)
Reagent Final concentration Quantity or Volume
Fluorescein sodium salts n/a 0.0941 g
DI water n/a 250 mL
Total (optional) 1 mM 250 mL
Fluorescein stock solution (0.1 mM)
Reagent Final concentration Quantity or Volume
1 mM fluorescein solution (Recipe 3) n/a 25 mL
DI water n/a 225 mL
Total (optional) 0.1 mM 250 mL
Laboratory supplies
Precision wipes (Kimtech Science, VWR, catalog number: 115-2029)
Silicon wafer (University Wafer, catalog number: 452)
Glass Petri dish (BRAND Petri dish) (Sigma-Aldrich, catalog number: BR455751)
1.5 mm biopsy punch (Integra Miltex, VWR, catalog number: 95039-092)
Double-wide (75 mm × 50 mm) microscope slides (Corning, catalog number: 2947-75X50)
Tape (Scotch Magic Tape or Highland invisible tape, 6200)
Lab labeling tape (VWR, catalog number: VWRU89097-928)
15 mL conical centrifuge tubes (Falcon, Fisher Scientific, catalog number: 352096)
50 mL conical centrifuge tubes (Falcon, Fisher Scientific, catalog number: 352070)
20 mL Luer lock plastic syringe (Codan, catalog number: 62.7602)
Tygon tubing (0.020” ID × 0.060” OD) (Masterflex, Fisher Scientific, catalog number: NC0578437)
0.2 μm syringe filters (Filtropur, VWR catalog number: 103573-246)
Ratchet tubing clamp (Cole-Parmer Celcon, Fisher Scientific, catalog number: 11752703)
23G Luer lock blunt syringe needles (CellLink, catalog number: NZ5231005001)
Retort stand (R&L Enterprises, Fisher Scientific, catalog number: 11779593)
Retort stand clamps (R&L Enterprises, Fisher Scientific, catalog number: 11517722)
Plastic Petri dishes (150 mm diameter) (VWR, catalog number: 25384-326)
Cutting knife (e.g., X-ACTO, catalog number: X3602)
Cutting pad (e.g., Fiskars, catalog number: 183720-1001)
Compressed nitrogen (e.g., Middlesex Gases & Technologies Inc., NI 3133)
Equipment
Spin coater (Laurell Technologies Corporation, WS-400B06NPP-Lite Manual Spinner)
Wafer alignment tool (Laurell Technologies Corporation)
Mask aligner (OAI, model: 204IR Mask Aligner)
Stylus profilometer (Bruker Dektak)
Hot plate (Cimarec+TM, model: Stirring Hot Plate MA-1827)
Plasma oven (Plasma Etch Inc, model: PE-25)
Inverted microscope with motorized stage (Nikon, model: Ti-E)
Scientific camera (Andor, model: Zyla 5.5)
Pressure controller (Elveflow, model: OB1)
Vacuum desiccator (e.g., Nalgene, Thermo Fisher Scientific, catalog number: 5310-0250)
Mylar photomasks (Artnet Pro, formally CAD/Art Services)
Software and datasets
AutoCAD from AutoDesk (https://www.autodesk.com/products/autocad). Free to students and educators, requires a license. Alternative commercial CAD software: Solidworks
Image processing and analysis can be carried out with the users’ preferred analysis software. Here, all image analysis and processing were carried out using MATLAB 2021b (MathWorks), typically available on institution licenses. Open-source alternatives: Python, Fiji (ImageJ)
Stage, microscope, and camera were controlled by National Instrument Systems (NIS) Elements. Open-source alternatives: Micromanager versions 1.4 and 2.0
Single pressure controller, controlled by default software (Elveflow). Alternative: Fluigent’s microfluidic flow controller
All data and code examples used for the calibration and validation of this protocol are publicly available [19,29].
Procedure
Below, we describe the step-by-step procedure for the fabrication, experimental setup, and calibration of the MCD. This device has been used to screen the chemotactic response of motile swimming bacteria to a logarithmically spaced concentration of a range of potential chemoattractants [19,21]. For the two-layer device, standard photolithography procedures [30,31] are used to fabricate two master molds. Subsequently, soft lithography with polydimethylsiloxane (PDMS) is used to assemble the MCD on a standard double-wide glass slide. The photomask designs are included in the Supplementary Material, and further data and example scripts are publicly available (see Software and Datasets). Photomasks were printed on mylar at a resolution of 25,400 dpi (Arnet Pro, formally CAD/Art Services Inc.).
Outline of the protocol:
Fabricating microchannel molds via photolithography
Fabrication of the master mold for the multi-height serial dilution layer
Fabrication of the master mold for the single-height cell injection layer
Silanization of the two master molds by vapor deposition
PDMS soft lithography of the two master molds
Assembly of the MCD
Experimental procedure
Calibration of flow rates
Fabricating microchannel molds via photolithography
Silicon wafer cleaning
Submerge and soak wafer in a separate glass Petri dish containing 50 mL of acetone for 3 min.
Submerge and soak wafer in a separate glass Petri dish containing 50 mL of isopropyl alcohol for 2 min.
Submerge and soak wafer in a separate glass Petri dish containing 100 mL of deionized water (DI H2O) for 2 min.
Remove wafer from DI H2O bath and rinse with a spray bottle containing DI H2O.
Dry wafer with compressed nitrogen.
Place clean wafer on a hotplate set to 150 °C for at least 15 min.
Note: Before continuing to the photolithography step below (step A2), make sure the wafer has cooled to room temperature. Failing to do so may negatively impact photoresist film thickness during the spin coating step.
General photolithography protocol
Spin-coat the silicon wafer with SU-8 photoresist:
i. Place the wafer on the spin coater using the wafer alignment tool to ensure the wafer is placed on the center of the chuck.
ii. Pour SU-8 photoresist onto the center of the wafer, covering approximately a 50 mm diameter area according to manufacturer's recommendations.
Note: Sufficient photoresist should be poured onto the wafer to ensure the photoresist completely coats the wafer after spin coating steps are completed below.
iii. Program and run spin coater with the following steps to form the desired photoresist film thickness:
1) Spin wafer with an acceleration of 100 rpm/s up to 500 rpm and hold for 10 s to evenly spread the photoresist onto the wafer.
2) Spin wafer with an acceleration of 300 rpm/s up to the necessary rotation speed (Si,i) to achieve the desired film thickness and hold for 30 s.
3) Spin with a deceleration of 100 rpm/s to ramp down to 500 rpm, hold for 5 s, and complete the spin coating protocol.
Soft bake (also known as prebake):
i. Place the wafer on a level preheated hotplate for 1 min at 65 °C.
ii. Increase the hotplate temperature to 95 °C to ramp up the temperature of the wafer and hold for the duration specified in the manufacturer’s data sheets, which is based on the thickness of the photoresist.
iii. Reduce the hotplate temperature back to 65 °C and remove the wafer after the temperature is achieved.
Note: Steps A2b.ii and A2b.iii are optional but are performed to limit thermal stresses generated by changes in temperature.
Load the wafer into the mask aligner, carefully aligning the photomask with the center of the wafer.
Transfer photomask pattern to SU-8 through UV exposure using mask aligner.
Note: The time duration (t) of the UV exposure is set by the power output (P) of the mask aligner, and the required exposure energy (E) is set by the thickness of the photoresist based on the manufacturer’s protocol: t = E/P.
Post-exposure bake (PEB)
i. Place the wafer on a level preheated hotplate for 1 min at 65 °C.
ii. Increase the hotplate temperature to 95 °C to ramp up the temperature of the wafer and hold for the duration specified by the manufacturer’s data sheets, which is based on the thickness of the photoresist.
iii. Slowly reduce the hotplate temperature back to room temperature.
Note: Steps A2e.i and A2e.iii are highly recommended to avoid thermal stresses generated by changes in temperature.
Photoresist development
i. Place the wafer inside a glass Petri dish and rinse with the photoresist developer (propylene glycol methyl ether acetate). Submerge the wafer in the developer and agitate the wafer frequently with wafer tweezers and/or laboratory rocker for at least 5–10 min, until features are fully developed.
ii. Rinse the wafer first with isopropyl alcohol and then DI water until all traces of undeveloped photoresist are removed and the rinsing agents run clear. Dry gently with compressed air.
Optional: Hard bake. The hard bake step is recommended to increase the lifespan of the molds and/or to anneal cracks if any are observed when viewing the molds under a microscope.
i. Place the wafer on a level hotplate set to 65 °C for 1 min.
ii. Increase the hotplate to 100 °C and hold for 1 min.
iii. Increase the hotplate to 150 °C and hold for 3 min.
Note: Duration of step A2g.iii can vary if cracks are still visible.
Fabricating the master mold for the multi-height dilution layer
Note: See “General notes: Microfabrication tolerances” for previous ranges of microfluidic tolerances.
Clean a silicon wafer based on step A1.
Mount the wafer to the spin coater, apply the photoresist SU-8 3050, and program the spin coater described in step A2a where S1,1 = 2,000 rpm for a target height of 90 µm.
Soft bake the wafer with photoresist described in step A2b, baking the wafer at 95 °C for 45 min.
Allow the wafer to cool gradually, then load wafer into the mask aligner with the first mask for layer 1 (main channel pattern).
Transfer the pattern by exposing it for the appropriate duration (see note on step A2d).
Move the wafer back to the hot plate and carry out the post-exposure bake (step A2e) at 95 °C for 5 min.
Return the wafer to the spin coater, apply the photoresist SU-8 2025 on top of the baked features, and program the spin coater described in step A2a where S1,2 = 4,200 rpm for a target height of 37 µm.
Soft bake the wafer with photoresist described in step A2b, baking the wafer at 95 °C for 6 min.
Allow the wafer to cool gradually, then load it into the mask aligner with the second mask containing the herringbone ridges for the dilution layer.
Align the herringbone photomask with wafer by adjusting the wafer (x-y translation and rotation) until the alignment markers are placed correctly (Figure 1).
Transfer the pattern by exposing for the appropriate duration (see note on step A2d).
Move the wafer back to the hot plate and carry out the post-exposure bake (step A2e) at 95 °C for 12 min.
Develop the wafer to remove the unexposed photoresist as described in step A2f and then hard bake as described in step A2g.
Figure 1. Multiplexed chemotaxis device (MCD) channel layout and multilayer photolithography using alignment markers. (A, B) Dilution layer (A) and cell injection layer (B) microchannel geometry scaled drawing. Scale bars 2 mm. (C) Multilayer photolithography was necessary to fabricate the herringbone ridges of the micromixer channels of the dilution layer. Alignment markers designed into the photomasks ensured proper orientation of the design. An inset of the microchannel design (magenta box from A) shows the alignment marker on the silicon wafer (i) used to align the herringbone ridge pattern within the photomask (ii) to ensure the ridges are placed in the correct location on the main channel (iii).
Fabricating the master mold of the single-height cell injection layer
Clean a silicon wafer based on step A1.
Mount the wafer to the spin coater, apply the photoresist SU-8 3050, and program the spin coater described in step A2a where S2 = 2,550 rpm for a target height of 75 µm.
Soft bake the wafer with photoresist described in step A2b, baking the wafer at 95 °C for 45 min.
Allow the wafer to cool gradually, then load the wafer into the mask aligner with the photomask containing the cell injection layer pattern (Figure 1).
Transfer the pattern by exposing for the appropriate duration (see note on step A2d).
Move the wafer back to the hot plate and carry out the post-exposure bake (step A2e) at 95 °C for 8 min.
Develop the wafer to remove the unexposed photoresist as described in step A2f, then hard bake as described in step A2g.
Silanization of the molds by vapor deposition
Critical: Silanization of the wafers, especially the dilution layer master mold, will ensure easy release of the cast PDMS at the end of the soft lithography process. This step will need to be repeated periodically after several castings to maintain easy release, which is signaled by the casting becoming increasingly difficult to remove from the wafer. Use of this silanization agent does not result in any negative effects on microbes and has been used successfully with other biological microfluidic assays [32,33].
Carefully remove the wafers from any containers, rinse first with isopropyl alcohol and then with DI H2O, and blow dry with nitrogen/compressed air.
Load 1–2 drops of silanization agent (tridecafluoro-1,1,2,2-tetrahydrooctyl trichlorosilane) onto a glass slide, a small glass container, or a disposable weighing boat.
Place the two wafers and silanization agent inside a vacuum desiccator for a minimum of 1 h.
Note: The wafers can be silanized separately if required due to space constraints.
Caution: Filters are recommended to prevent the silanization agent from entering the vacuum system/pump. Where possible, use a dedicated vacuum chamber for chemicals to prevent cross-contamination for other applications.
Remove the wafers from the vacuum desiccator and place in fresh plastic Petri dishes for storage prior to soft lithography. Secure the edges of the wafers to the base of the Petri dishes with lab tape.
PDMS soft lithography of the two wafers
Prepare polydimethylsiloxane (PDMS) in a disposable container, e.g., plastic cup, in a 10:1 ratio (base:curing agent, see Recipe 1).
If the wafers have been previously cast, then approximately 20–30 g of PDMS base per wafer is required. Otherwise, if this is the first casting post-silanization, 80–100 g of PDMS base is recommended per wafer to ensure sufficient coverage of the entire Petri dish and wafer.
Note: Excess PDMS will be pourable for approximately 12 h (depending on the exact curing agent ratio), so multiple casts can be completed with a sufficient quantity of prepared PDMS.
Thoroughly mix the PDMS with a disposable implement (e.g., plastic spreader, plastic knife) for at least 1 min until the solution is opaque with bubbles.
Note: A figure-of-eight motion is recommended for improved incorporation of the two components, with regular scraping of the sides and bottom of the container.
Place the mixed PDMS inside a vacuum desiccator and degas the mixture for at least 10 min until all bubbles are removed.
Remove the PDMS from the vacuum chamber and gently pour over the two molds, tilting to ensure complete PDMS coverage of each wafer.
Replace the wafers back inside the vacuum chamber and degas for approximately 10 min until most bubbles have been eliminated.
Remove the wafers from the vacuum chamber.
Optional: Gently burst any remaining bubbles on the surface with gentle bursts of compressed air.
Place the poured mold inside in a level oven at 60–70 °C for at least 1 h until the PDMS is fully cured.
Note: The PDMS should be firm to the touch and not sticky when fully cured.
Carefully remove the PDMS slabs from the wafers by cutting around the edge of each layer with a sharp, precision knife until the PDMS perimeter has been fully cut. Carefully lift the edges of the slab away from the mold, working around the slab until it is fully free of the mold.
Caution: Care must be taken not to tear any of the PDMS features, particularly around the serpentine herringbone ridges of layer 1. The PDMS should release with relative ease; if not, repeat the silanization process (see section D).
Cover both sides of each PDMS slab with Scotch tape to protect from dust.
Carefully punch the tubing ports with a 1.5 mm biopsy punch (or equivalent tool) for both the dilution layer (chemical inlet, buffer inlet, Ai, and Bi) and cell injection later (cell inlet and buffer inlet) as shown in Figure 1A, B.
Caution: Punch the ports as perpendicular to the plane of the device as possible; sloped ports will make alignment of the two layers (see step F10) more difficult.
Store the completed PDMS layers in a Petri dish prior to assembly.
Pause point: The procedure can be paused here indefinitely, until the devices are required for assembly.
Assembly of the MCD
Rinse the dilution layer PDMS with ethanol and then deionized water, blow drying with nitrogen/compressed air.
Clean the PDMS with Scotch tape by repeatedly applying then removing tape from all surfaces of the PDMS, taking care to clean within individual features. Leave a layer of tape over the clean features to protect microchannels from dust before continuing to the next steps. Scotch tape is recommended as it does not leave a residue on the surfaces and remains easy to remove after a long time.
Clean the glass slide (standard double-wide glass slide, 75 mm × 50 mm × 1 mm) with isopropyl alcohol scrubbing firmly with tissue (e.g., Kimtech wipes). Rinse with DI water, then blow-dry with nitrogen/compressed air. If necessary, the glass slide can be soaked in harsher treatments, e.g., hydrogen peroxide or standard hydrochloric acid cleaning processes to remove any residue and/or coatings from the surface.
Place a double-wide glass slide and dilution layer PDMS feature-side up (removing any protective tape) inside a plasma oven and expose the surfaces to the plasma for 60 s.
Remove glass and PDMS from the plasma oven using tweezers, taking care not to touch the exposed faces of either material. Invert the PDMS, so microchannels are facing down, and gently bring into contact with the glass slide (Figure 1A).
Note: If air gaps get trapped between the glass slide and PDMS, lightly press the top of the PDMS with tweezers to remove.
Immediately after contact, place the partially assembled device on a hot plate at 110 °C for at least 30 min to promote covalent bonding.
Inspection: Under a microscope, inspect the microchannels to ensure no debris is blocking the microchannels. If debris is observed, attempt to remove it by flowing DI water through the channels; however, if the debris remains, it is recommended to cast a new PDMS layer instead.
Pause point: The protocol can be paused here indefinitely. It is advised and recommended that the first bonded layer is kept in a dust-free location (e.g., in a Petri dish) with Scotch tape covering the top of the PDMS to prevent dust accumulation in the inlets and on the bonding surface.
Clean both PDMS surfaces: feature-side of cell injection PDMS and exposed top of the bonded dilution PDMS. Use Scotch tape to clean by repeatedly applying then removing tape from all surfaces of the PDMS, taking care to clean within individual features. Leave a layer of tape over the clean features for dust protection.
Place glass-bonded dilution layer PDMS and cell injection layer PDMS feature-side up (removing any protective tape) inside a plasma oven and run for 60 s.
Remove the partially assembled device and PDMS from the plasma oven using tweezers, taking care not to touch the exposed faces of either material.
Invert the cell injection layer so the microchannels are facing downward. Carefully, by eye, align the features of cell injection layer with the punched ports of the dilution layer (Figure 1B) and then bring the two layers into contact.
Critical: At this stage, the PDMS-PDMS bond is partially reversible, so there is a margin (~3–5 s) for removing the cell injection layer if the two PDMS pieces are incorrectly aligned (Figure 1C, D). However, disturbing the bond could compromise the device’s integrity. It is recommended that steps F9–11 are repeated if the two layers are separated during alignment.
Note: A microscope (e.g., stereomicroscope) can also be used to aid in alignment if necessary.
Place the glass-side down on a hot plate at 110 °C for at least 1 h to improve bond quality. Then, place the device on a 70 °C hotplate overnight. Store in dust-free condition.
Experimental procedure
Critical: Careful loading and preparation of fluid connections is critical for successful operation of the MCD. It is essential, as with standard microfluidic techniques, that any air bubbles are removed from the device prior to experiments.
Prepare 100 mL of 0.5% (w/v) BSA in DI H2O (see Recipe 2), vortex thoroughly, and then filter through a 0.2 μm syringe filter. Store in cool dark conditions until required.
Optional: Place the fully assembled MCD inside a chamber under vacuum for 15 min to remove air from inside the device and PDMS.
Note: The total time to degas the MCD in the vacuum desiccator depends on the overall thickness of the device. If unsure, an extended degassing period of 30 min is recommended.
Prepare four 20 mL syringes for gravity-fed filling of the MCD. For each syringe:
Remove the plunger and attach a 23G Luer lock needle.
Thread the end of 60 cm of Tygon tubing (henceforth referred to as tubing) onto the needle.
Attach each syringe (facing downward) into a lab flask grip stand at least 50 cm high.
Immediately after removing the MCD from the vacuum chamber, quickly fill each syringe with the prepared BSA solution; once fluid begins to exit the tubing, connect the tubing to each inlet of the MCD (Figure 1A, Figure 2A).
Figure 2. Multiplexed chemotaxis device assembly. (A) Shown with food coloring, successful assembly results in passive serial dilution and injection of the chemical solution with buffer from the dilution layer as well as the injection of buffer and cells from the cell injection layer. Dashed box corresponds to the observation region where experimental data is recorded. Scale bar, 5 mm. After assembly, inspection of the connection ports between the cell injection layer and dilution layer is necessary to ensure the MCD operates correctly. (B) Good alignment between the dilution layer (green dashed circle) and cell injection layer (blue dashed circle) is achieved when the punched holes in the dilution layer do not overlap with the labeled microchannels (blue arrows). (C) An example of misalignment in the intersection of the microchannel (blue arrow) of the cell injection layer (dashed blue circle) and the punched port of the dilution layer (red dashed circle), which will change the hydraulic resistance of the microchannel thus negatively altering the symmetry of stratified chemical, cell, and buffer solutions in the observation region (see Figure 3). Scale bar (B and C), 1 mm.
Connect outlet waste tubing to the MCD and then clamp the tubing closed near the open end of the tubing once no more air bubbles can be seen exiting the device.
Note: The hydrostatic pressure from the BSA syringes will slowly remove any air pockets/bubbles from the device. The BSA will pre-treat the surfaces to reduce adhesion of bacteria to the device.
Leave the device under gravity-pressure until no bubbles are visible within the device; allow at least 30–60 min or until the bubbles have disappeared. The required duration depends on the pressure applied and initial wetting of the microchannels. If a desiccator is not used to remove air from the device and PDMS (see optional step G2), the device can be placed under gravity-pressure overnight to slowly remove any bubbles. Proceed with step G7 whilst step G6 completes.
Critical: Bubbles stuck to the channel walls will compromise the device operation, and thus enough time should be allowed for the air bubbles to be absorbed in the PDMS prior to experiments.
Notes:
More pressure can be applied by either (i) increasing the height of the syringes above the device, or (ii) reinserting the plungers and using an elastic band to exert pressure on each of the syringes.
Alternative methods for degassing the MCD are described in the General Notes section.
Prepare experimental solutions for the four inlets (Figure 1) connected to the pressure controller.
Note: See previous applications for examples for preparation of specific experimental solutions [19,21].
Dilution layer—chemical inlet: solution of chemostimulus dissolved into buffer solution (e.g., artificial seawater, minimal motility media), concentration C0.
Dilution layer—buffer inlet: buffer solution.
Cell injection layer—cell inlet: cells washed and resuspended into buffer solution.
Cell injection layer—buffer inlet: buffer solution.
Disconnect the MCD from all BSA syringes, making sure to leave a droplet on top of each inlet/outlet to facilitate attachment of the inlet tubing.
Attach each inlet tubing with the device in turn:
Briefly turn on the pressure controller to form a small droplet on the end of the tubing, then stop the flow.
Critical: Bring the tubing into contact with the droplet on top of the corresponding inlet of the MCD, maintaining a fluid-fluid connection at all times to prevent air being pushed into the device. If the end of the tubing exits the droplet before connection is established, reset the droplet using the pressure controller.
Push the tubing into the inlet, using tweezers if necessary to ensure a firm hold. Test that the tubing is inserted correctly with a light tug on the tubing; the tubing should remain firmly attached to the MCD.
Insert outlet tubing (40 cm) into the outlet and connect it to a waste container.
Secure the MCD to the microscope stage.
Ensure the liquid–air interfaces of each fluid inlet container and outlet container are level (i.e., at the same relative heights). Raise/lower individual Falcon tubes as necessary to achieve this.
Note: The liquid–air interface of each container should be level across all containers, so they have the same absolute pressure resulting in a more accurate applied pressure from the pressure controller. This process eliminates the potential for residual hydrostatic-driven flows.
Setup microscope acquisition. Locate a mid-plane observation point in each experimental channel (Figure 1A). An experimental time of approximately 10 min is recommended to start. Slower diffusing chemoattractants might require longer acquisition times due to a longer persisting gradient. The magnification should be selected to allow for imaging of the full channel width in a single field of view, which is dependent on the available scientific camera (typically expect a magnification between 4× and 10×).
Start the pressure controller using pressure ratios from calibration (see section H). The designed applied pressure ratio between the dilution layer (pin,1-2) and cell injection layer (pin,3-4) are 2:1, though minor pressure adjustments will be necessary to have symmetric stratified chemical, cell, and buffer solutions in the observation region (see Figure 3) due to minor variations in the replica mold manufacturing processes [19].
Figure 3. Multiplexed chemotaxis device (MCD) validation and calibration. (A) Shown with fluorescein (green) and DI H2O, the MCD is functioning properly when the stratified chemical, cell, and buffer solutions are symmetric in the observation region. Scale bar, 0.1 mm. (B) Normalized measured widths, , of the chemical, cell, and buffer streams illustrate their pronounced symmetry in each observation channels (C0-5, Figure 1A); where wi is the width of fluid stream (A; solid white lines) and W the total width of the intensity profile (A; dashed white-red lines). The bar plots and error bars indicate the mean and standard deviation of for three separate driving pressures of the dilution layer and cell injection layer: (i) 100 mbar and 70 mbar, (ii) 150 mbar and 105 mbar, and (iii) 200 mbar and 140 mbar for pin,1-2 and pin,3-4, respectively. To achieve symmetric laminar flow, the applied pressure, pin,3-4, was tuned. The slight deviations from the designed inlet pressures are due to a variety of experimental conditions such as (i) variations in the fabricated channel height of the dilution layer and cell injection layer master molds and/or (ii) variations in the inlet tubing lengths that supply the solutions to the MCD. These deviations are fixed per set of master molds; hence, flow rate calibration only needs to be carried out once per set of master molds.
After at least 2 min of continuous flow, simultaneously stop the pressure controller and clamp close to the open end of the tubing connected waste outlet. Then, begin acquisition.
After acquisition is completed, unclamp the waste outlet and cut the crimped tubing. Then, restart the flow and repeat steps G14–15 for as many technical replicates as required. Biological replicates can be easily conducted by replacing the cell suspension and restarting the flow.
Calibration of flow rates
Critical: Calibration of the MCD only needs to be done with one complete device per set of wafers. Subsequent devices from the same wafers will perform identically, provided they are assembled correctly. The purpose of this calibration step is to establish the correct pressure ratio between the two layers to account for minor deviations in geometry due to the microfabrication processes. The designed pressures are pin,1-2 = 100 mbar, pin,3-4 = 50 mbar.
Prepare 100 mL of 0.1 mM fluorescein solutions in DI H2O and 100 mL of DI water (see Recipes 3 and 4).
Setup the MCD as per section G, flowing the 0.1 mM fluorescein solution through both the chemical and buffer inlet of the dilution layer, DI H2O through the cell inlet of the cell injection layer, and 0.1 mM fluorescein through the buffer inlet of the cell injection layer.
Prepare the microscope for fluorescence microscopy with a GFP filter cube (for excitation of the fluorescein). Select a magnification that allows for imaging of the full width of the observation channel (1 mm) in a single field of view, typically in the range 4–10×.
Turn on the pressure controller at the designed pressure ratios (pin,1-2 = 100 mbar, pin,3-4 = 50 mbar; Figure 4).
Note: The observation regions will now have three distinct laminar fluid layers visible: two bright fluorescent bands from the chemical inlet of the dilution layer and the buffer inlet of the cell injection layer width (w1,3). The center width (w2) contains DI H2O from the cell inlet.
Acquire an image in each observation region (Figure 3).
Adjust the pressure of the cell injection layer (pin,3-4, Figure 4) until w1 ≈ w3 and then acquire a second image at each observation region (Figure 3).
Precise widths for each fluid stream can be easily measured using the spatial intensity across the width of the observation region channel, either in the microscope software or in post-processing.
Critical: These pressure ratios will be robust across all devices made from a particular set of molds as long as the viscosity of the solutions is the same, and the absolute pressures can be increased as long as the relative pressure ratio (pin,1-2/pin,3-4, Figure 4) remains constant. See previous applications [19] for examples of the device operating at different absolute pressures with a constant pressure ratio.
Figure 4. Multiplexed chemotaxis device (MCD) design. Circuit representation of the dilution layer (A) and cell injection layer (B) of the MCD. The pressure at each node (pi), microchannel volumetric flow rate (Qi), and microchannel hydraulic resistance (Ri) must be known before designing and fabricating the MCD.
Data analysis
A full description of data analysis methods has been previously described [19,21] and data analysis from previous applications and example calibration data are publicly available [29]. Alternatively, image analysis can be easily conducted with the software package of choice (e.g., Python, MATLAB, ImageJ) with commonly available image processing toolboxes. In summary, images were processed by removing the average (mean) background and applying a spatial bandpass filter; then, features were identified to sub-pixel accuracy. With the cell locations identified, the chemotactic behavior can be visualized through a variety of methods. Previous examples include heatmaps of the conditional probability [19,21] P(y|t), accumulation toward the boundaries [19,21], and migration coefficients calculated from average cell distance from the center of the channel [15,34,35].
Validation of protocol
This protocol has been used and validated in the following research articles:
Stehnach et al. [19]. Multiplexed microfluidic screening of bacterial chemotaxis. eLife.
Fabrication and calibration: Figure 1, Figure 1 Figure Supplements 1–4
Validation: Figure 2, Figure 2 Figure Supplement 1
Applications: Figure 3 and Figure 4
Dataset [29]
Henshaw et al. [21]. Early stage viral infection of cyanobacteria drives marine bacterial chemotaxis. bioRxiv.
Applications: Figure 4
General notes and troubleshooting
General notes
Alternative technique to assist in the PDMS-PDMS bonding.
After treating the two PDMS microchannels in the plasma oven (step F8), a small drop of DI H2O can be placed on the top of the dilution layer PDMS before aligning the cell injection PDMS [36]. This allows for the two PDMS pieces to slide into place to ensure they are properly aligned.
Microfabrication tolerances.
The microfabrication process can vary due to a range of factors including age and temperature of the SU-8 and accuracy of the spin coater. Therefore, there will be deviations in the final channel heights away from the designed target heights. It is crucial that the height of the microchannel remains constant across the wafer; a systematic deviation away from the target height is more desirable than spatially varying mold heights across the master mold. In previous applications of this protocol [19,21], the target and achieved channel heights were:
Dilution layer main channel: Target, 90 μm; Achieved, 90–94.5 μm.
Herringbone ridges of the dilution layer: Target, 37 μm; Achieved, 37–38.5 μm.
Cell injection layer: Target, 75 μm; Achieved, 71–73 μm.
The MCD will perform robustly despite these variations if each master mold is calibrated once prior to use (as outlined in the protocol).
Hydraulic circuit and microchannel design.
Designing the MCD using hydraulic circuits (Figure 4): The Hagen-Poiseuille law describes the flow characteristics for an incompressible fluid in a channel with laminar flow as:
Δp = QRH
where Δp is the pressure drop along the channel, Q is the volumetric flow rate, and RH is the hydraulic resistance defined by a combination of the geometrical properties of the channel and mechanical properties of the fluid [37]. The channel lengths within the provided hydraulic circuits for this protocol (Supplementary Materials) are determined by setting the following: channel height, applied inlet pressures pin,1-2, outlet pressure pout, outlet flow rate Qout, resistance of the bridge channel RB, resistance of the outlet R4,4, and resistance of the herringbone micromixer channel RM. A set of simultaneous equations can then be established and solved for the unknown flow rates, pressures, and hydraulic resistances (and corresponding channel dimensions). This process will need to be repeated if the user wishes to alter features of the MCD such as dilution ratios, linear versus logarithmic dilution, or to increase/decrease the number of observation channels for example. For a complete description of this process, which is out of the scope of this protocol, please see the original work [19].
Standard microfluidic procedures.
This protocol builds upon standard microfluidic techniques, which will be familiar to experienced users. However, for less experienced users, we recommend reviewing the cited literature throughout this protocol and additionally the following materials [38–40]:
O’Laughlin et al. [38]. Fabrication of Microfluidic Devices for Continuously Monitoring Yeast Aging. Bio-protocol.
Taly et al. [39]. Microfluidics Diagnostics: Methods and Protocols. Springer
Bruggeman et al. [40]. Microfluidics and fluorescence microscopy protocol to study the response of C. elegans to chemosensory stimuli. STAR Protocols
Alternative methods for degassing/filling of microfluidic devices.
Users can use any preferred method for removing air bubbles from the microchannels, such as the described “gravity feed” method in the main text. Below, we outline two commonly used alternative strategies to achieve the desired degassing and subsequent filling of the microfluidic device:
Flowing DI H2O through the device at a high flow rate with some perturbations to dislodge any bubbles stuck to the channel geometry, using either a syringe or peristaltic pump.
Fully submerge the device in DI H2O, then place in a vacuum desiccator. The vacuum will draw the air out from the device and pull the liquid into the microchannels.
Acknowledgments
We thank J. Vlahakis of the Tufts Micro and Nano Fabrication Facility for assistance in the device fabrication. This work was funded by NSF Awards OCE-1829827, CAREER-1554095, and CBET-1701392 (to J.S.G.), and OCE-1829905 (to S.A.F.). This protocol has previously been described and validated.
This protocol was derived from the original work [19] of Stehnach et al. 2023 and the generation of its derivative dataset [29].
Competing interests
The authors declare that no competing interests exist.
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Supplementary information
The following supporting information can be downloaded here:
SI_Material_MCD_CAD_Designs_AllLayers.dwg: CAD Art files containing the photomasks designs outlined in this protocol.
Article Information
Publication history
Received: Apr 29, 2024
Accepted: Jul 28, 2024
Available online: Aug 12, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cell Biology > Cell imaging > Microfluidics
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5,063 | https://bio-protocol.org/en/bpdetail?id=5063&type=0 | # Bio-Protocol Content
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FixNCut: A Practical Guide to Sample Preservation by Reversible Fixation for Single Cell Assays
SW Shuoshuo Wang
LJ Laura Jiménez-Gracia
AA Antonella Arruda de Amaral
IV Ioannis S. Vlachos
JP Jasmine Plummer
HH Holger Heyn
Luciano G. Martelotto §
(§ Technical contact)
Published: Vol 14, Iss 17, Sep 5, 2024
DOI: 10.21769/BioProtoc.5063 Views: 826
Reviewed by: Pilar Villacampa AlcubierreLeonor GouveiaSaba Asam
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Original Research Article:
The authors used this protocol in Genome Biology Mar 2024
Abstract
The quality of standard single-cell experiments often depends on the immediate processing of cells or tissues post-harvest to preserve fragile and vulnerable cell populations, unless the samples are adequately fixed and stored. Despite the recent rise in popularity of probe-based and aldehyde-fixed RNA assays, these methods face limitations in species and target availability and are not suitable for immunoprofiling or assessing chromatin accessibility. Recently, a reversible fixation strategy known as FixNCut has been successfully deployed to separate sampling from downstream applications in a reproducible and robust manner, avoiding stress or necrosis-related artifacts. In this article, we present an optimized and robust practical guide to the FixNCut protocol to aid the end-to-end adaptation of this versatile method. This protocol not only decouples tissue or cell harvesting from single-cell assays but also enables a flexible and decentralized workflow that unlocks the potential for single-cell analysis as well as unconventional study designs that were previously considered unfeasible.
Key features
• Reversible fixation: Preserves cellular and molecular structures with the option to later reverse the fixation for downstream applications, maintaining cell integrity
• Compatibility with single-cell assays: Supports single-cell genomic assays such as scRNA-seq and ATAC-seq, essential for high-resolution analysis of cell function and gene expression
• Flexibility in sample handling: Allows immediate fixation post-collection, decoupling sample processing from analysis, beneficial in settings where immediate processing is impractical
• Preservation of RNA and DNA integrity: Effectively preserves RNA and DNA, reducing degradation to ensure accurate transcriptomic and genomic profiling
• Suitability for various biological samples: Applicable to a wide range of biological samples, including tissues and cell suspensions, whether freshly isolated or post-dissociated
• Enables multi-center studies: Facilitates collaborative research across multiple centers by allowing sample fixation at the point of collection, enhancing research scale and diversity
• Avoidance of artifacts: Minimizes stress or necrosis-related artifacts, preserving the natural cellular physiology for accurate genomic and transcriptomic analysis
Keywords: Single cell Tissue Fixation Lomant's reagent DSP Sample preservation Single-cell sequencing
Graphical overview
Background
Single-cell techniques have revolutionized biological and clinical research by quantitatively capturing the genomic and transcriptomic state of individual cells at unprecedented resolution and scope [1]. This approach moves beyond averaging the bulk population, which consists of various cell types. However, the technical and logistical challenges of acquiring or generating high-viability single-cell suspensions have limited the application of single-cell genomics and transcriptomics.
Multi-center clinical studies often involve sample collection from remote areas lacking processing infrastructure [2,3]; additionally, longitudinal and cohort recruitment of patients occurs over the years, subject to patient availability and logistics constraints. Moreover, even in advanced clinical centers, the urgency of tissue harvesting during surgical procedures, biopsies, or autopsies can depend on factors such as the type of surgery, the purpose, and the specific protocol followed, as well as the transfer procedures between surgery and histopathology [4,5]. In basic research using organoids, embryos, or rare specimens, the limited cell number poses a significant bottleneck for conducting scientifically meaningful and economically feasible single-cell analysis.
In many experimental setups typical for clinical research, the freeze-thaw process remains the predominant option for sample collection en masse. However, freezing will physically damage the cell membrane, impact the post-thaw viability, and disrupt intracellular compartments where endogenous RNases are sequestered, leading to RNA degradation. In addition, cryopreservation may lead to the loss of certain cell types and induce cellular stress response [6]. Moreover, variably staggered freezing and thawing through aliquoting cells for parallel orthogonal assays, for example, single-cell mass cytometry (CyTOF), single-cell RNA-Seq, single-cell ATAC-Seq, and additional multiomic assays, can lead to inconsistency, introducing technical artifacts and batch-wise noise to the data. Recently, the fixed RNA (flex) assay has been gaining popularity due to its robustness; however, so far probes are only available for humans and mice and do not target immunologically and clinically paramount transcripts such as joining and variable regions in B and T-cell receptor (BCR and TCR), Killer cell immunoglobulin-like receptors (KIRs), and human leukocyte antigen (HLA), nor polyadenylated non-coding RNA.
Notably, many important immune cells—such as neutrophils, dendritic cells, monocytes, macrophages, and lymphocytes (B cell and T cells), as well as cells with higher intrinsic mechanical integrity such as epithelial cells—are considered fragile for single-cell applications due to their susceptibility to mechanical damage during sample preparation, including cell sorting or tissue dissociation [7–9]. Loss of these cells often leads to underrepresentation in downstream analysis. Therefore, identifying a method allowing the stabilization of fragile cells without perturbing RNA integrity will enhance the recovery of these important populations.
The ideal fixative must possess several characteristics: first, it should be small and able to penetrate the cell membrane and tissues. Second, it should preserve the structure and integrity of cells. Third, it should not destroy RNA integrity and should, ideally, inhibit RNase activity at least temporarily. Currently, one of the most common options for single-cell assays is using either standard or modified methanol fixation [10–14]. Methanol works by dehydrating samples and denaturing proteins in a gentle manner, similar to histology fixation. However, methanol as a standalone fixative has raised concerns regarding biased results or ambient RNA leakage in single-cell RNA-Seq [6]. A possible mechanistic explanation is the irreversible intracellular compartment disruption resulting in the loss of normal lipid and protein structure during dehydration–rehydration cycles. In addition, incomplete reverse transcription of mRNAs with more complex secondary structures was suggested as a major caveat [15].
In contrast, dithio-bis-succinimidyl propionate (DSP), known as Lomant's reagent [16], has been traditionally used for histological tissue fixation to stabilize cellular integrity and structures through covalent crosslinking of free amine groups found at the N-terminus of polypeptide chains (diagrammatic molecular basis is depicted in Figure 1 of Akaki et al. [17]). It has been repurposed for single-cell RNA sequencing by maintaining RNA integrity and yield in bulk RNA extractions [18–20]. The cell and membrane permeability have been proven to be particularly useful in detecting rare cell populations such as tissue-resident immune cells. Certain T-cell populations can exist in very low numbers upon activation, making them difficult to isolate and analyze with conventional single-cell methods. A protocol called CLInt-Seq (crosslinker regulated intracellular phenotype sequencing) was developed to overcome the hurdle by crosslinking intracellular cytokines [21]. This technique combines and improves upon existing techniques to collect and genetically sequence rare T cells. The reversibility of DSP-induced crosslinking is a key feature allowing cells and tissue to be further processed or dissociated for downstream applications. However, a recent study concluded that de-crosslinking the DSP-fixed samples is optional, and additional handling steps of de-crosslinking may contribute counterproductively to cell loss [22].
Recently, we developed the further optimized FixNCut protocol and demonstrated its robustness and benefits systematically by comparing fresh and fixed lung, colon, and pancreas samples from different species even under cryopreservation for an extended period [23,24]. Here, we provide an end-to-end solution in the form of a practical guide on the implementation of this method, to remove practical barriers by streamlining the transport of samples and scheduling of shared instruments for downstream single-cell isolation and processing.
Materials and reagents
Biological materials
Tissue samples and cell suspensions can both be used. For cells, the protocol is set up for up to 2 × 106 cells. For more cells, scale up the fixation accordingly. Tissues larger than 3 mm in diameter or edge length need to be partitioned into smaller pieces to facilitate fixative penetration
Reagents
DSP (Dithio-bis-succinimidyl propionate), also known as Lomant's reagent (Thermo Fisher, catalog number: 22586)
DMSO, anhydrous (Thermo Fisher, Molecular Probes, catalog number: D12345)
LiberaseTM research grade, 10 mg (Roche, catalog number: 5401119001)
Protector RNase inhibitor (40 U/μL) (Roche, catalog number: RNAINH-RO)
UltraPureTM 1 M Tris-HCI buffer, pH 7.5 (Thermo Fisher, Invitrogen, catalog number: 15567027)
10× PBS buffer, pH 7.4 (Invitrogen, catalog number: AM9624 or AM9625)
Ambion nuclease-free water (Invitrogen, catalog number: AM9932)
Bovine serum albumin (BSA) solution, sterile filtered and cell-culture tested (Sigma Aldrich, catalog number: A1595)
Tris base, UltraPure Tris buffer (powder format) (Thermo Fisher, Invitrogen, catalog number: 15504020)
Solutions
Fixation concentrate stock solution (1 mL, for 100 standard assays) (see Recipes)
PBS with 1% BSA (see Recipes)
1 M Tris pH 7.5 (optional, if not using ready-made buffer) (see Recipes)
Recipes
Fixation concentrate stock solution (1 mL, for 100 standard assays)
Reagent Final concentration Amount
DSP (powder) n/a 50 mg
DMSO n/a 1 mL
Total 50 mg/mL (w/v) 1 mL
PBS with 1% BSA
Reagent Final concentration Amount
10× PBS n/a 50 mg
BSA 10% n/a 1 mL
Total n/a 1 mL
1 M Tris pH 7.5 (optional, if not using ready-made buffer)
Start with 30 mL of water and adjust pH to 7.5 by adding 5 M HCl to a final volume of 50 mL.
Reagent Final concentration Amount
Tris base n/a 6.05 g
Nuclease-free water n/a 50 mL
Total 1 M 50 mL
Laboratory supplies
DNA LoBind tubes 1.5 mL (Eppendorf, catalog number: 022431021)
DNA LoBind tubes 2.0 mL (Eppendorf, catalog number: 022431048)
Flowmi® cell strainers, porosity 40 μm, for 1000 μL pipette tips (Sigma, Scienceware, catalog number: BAH136800040)
pluriStrainer mini 70 μm (pluriSelect, catalog number: 43-10070-40)
Falcon conical centrifuge tubes (Corning, catalog number: 352070 for 50 mL, 352095 for 15 mL)
Sterile serological pipettes with pipettor
Rainin Pipet-Lite LTS pipette tips (Rainin, catalog number: 30389240, 30389213, 30389226)
Equipment
Centrifuge 5810/5810R (Eppendorf, catalog number: EP022628188) with rotor S-4-104
Rotor F-35-6-30 (Eppendorf, catalog number: EP5427716009)
S-4-104 rotor adapters for 50 × 1.5/2 mL tubes (Eppendorf, catalog number: 58-257-40009)
Standard heavy-duty vortex mixer (VWR or Fisherbrand, catalog number: 97043-562)
ThermoMixer C (Eppendorf, catalog number: 05-412-503) or thermoblock with minimal temperature fluctuation
Ice bucket with cool blocks or Cool Rack CFT30 (Corning, catalog number: CLS432052)
Automated cell counter, e.g., Luna-FX7 (Logos Biosystems) or hemocytometer
CoolCell freezing container for 12 × 1 mL or 2 mL cryogenic vials (Corning, catalog number: 432000) or Mr. Frosty freezing container (Thermo Scientific, catalog number: 5100-0001)
Tissue-Tek cold plate (VWR Scientific, catalog number: 25608-942)
Procedure
50× fixation concentrate stock preparation (50 mg/mL)
DSP equilibration
Thoroughly equilibrate the DSP package to room temperature for 30 min prior to first opening. Critical: Only open the package after equilibration.
Note: The NHS-ester of DSP is susceptible to hydrolysis upon contact with the humid atmosphere and is therefore moisture sensitive. Thorough equilibration is crucial in preventing condensation and premature inactivation of DSP.
50× working solution stock preparation
Dissolve 50 mg of DSP in 1 mL of high-quality anhydrous DMSO for best performance. Final concentration: 50 mg/mL.
Working stock storage
For short-term storage, dispense 10 μL from the stock into 1.5 mL or 2 mL Eppendorf safe-lock microtubes and store at -80 °C in a sealed container or bag with desiccants.
Alternatively, 10 μL aliquots can be stored in cryogenic vials with compression O-rings to prevent moisture absorption. Avoid freeze-thaw cycles by limiting the size of aliquots (up to 100 μL). Any opened and unused stock must be discarded.
Working solution preparation (1 mg/mL)
Note: Time-sensitive step.Prepare fresh working solution immediately before the fixation process.
Using a P200 pipette, slowly and drop-by-drop add 490 μL of PBS to an Eppendorf tube containing 10 μL of 50× stock while vortexing (Video 1).
Caution: Choose “touch mode” and maximal speed (3,500 rpm or no less than 2,500 rpm) on the vortex instead of the “on mode”. Using a P1000 pipette is acceptable too, but adding the PBS too fast will cause DSP to precipitate, which perturbs the effectiveness of fixation. It is not concerning to see tiny precipitates on the wall of the tube initially; however, in case of substantial precipitates, repeat by slowing down the PBS addition or start with a new DSP aliquot.
Video 1. Working solution preparation
Filter the working solution once using a 40 μm Flowmi strainer and transfer it to a new 1.5 or 2 mL tube to remove larger precipitates.
Critical: Working solution standing for more than 10 min should not be used and must be discarded. If sample preparation takes longer than 10 min, prepare the working solution during the process to avoid leaving it standing.
Fixation
Note: For this section, follow instructions based on your sample type.
Fixation of tissue samples
For larger tissue samples or organoids (3 mm or larger in diameter/edge length), finely cut or mince the tissue with a sterile razor blade on a cold plate to facilitate working solution penetration. For effective fixation, tissue samples should be no thicker than 3 mm, with 1–2 mm being preferable for uniformity. This is because fixatives penetrate through passive diffusion, and the higher molecular weight of DSP (14.5-fold paraformaldehyde and 4-fold glutaraldehyde) significantly reduces its penetration speed and depth in tissue.
Note: Fine mincing without separating the tissue minimizes sample loss during washing and transferring into a new tube.
Add the minced tissue to a new Eppendorf tube containing 500 μL of working solution and incubate at ambient temperature for 30–45 min.
Gently invert the tube every 15 min. For certain tissue types, if the tube becomes overly bloody or cloudy, consider replenishing once with fresh working solution.
Fixation of cells
Wash the cells in cold, sterile RNase-free PBS.
Centrifuge the cells using an appropriate speed depending on type and fragility: usually between 150× g for 3 min (large cells) and 600× g for 10 min (small cells). Keep the cell pellet and remove the supernatant. Resuspend in cold, sterile RNase-free PBS.
Repeat this step for a minimum of two washes.
Note: Remnant media containing FBS (see General notes, Tip 4), storage additives, or sheath fluid components may interfere with the crosslinking.
Pellet the cells, resuspend in 500 μL of working solution, and incubate for 30 min at ambient temperature.
Gently invert the tube once at the 15-min mark.
Quenching of reactive DSP
Note: For this section, follow instructions based on your sample type.
For cells
Add 10 μL of 1 M Tris-HCl pH 7.5. Then, mix thoroughly on a vortex for 2–3 s followed by a 15 min incubation at room temperature.
Note: As an amine-reactive crosslinker, excessive reactive DSP must be quenched with Tris-HCl buffer before proceeding. The quantity is stoichiometrically optimized and must be adjusted accordingly if more fixative was used for larger tissues or a higher number of cells.
Centrifuge at 500× g for 5 min at ambient temperature. Remove supernatant.
Resuspend the cell pellet in 1,000 μL of PBS. Mix by vortexing for 2–3 s.
Centrifuge at 500× g for 5 min at ambient temperature. Remove the supernatant.
Repeat steps D1c–D1d for a total of two washes.
Resuspend the cells in 0.5–1 mL of cold PBS with 1% BSA.
Note: For maximal RNA integrity, add 0.2–1 unit/μL RNase inhibitor to the final resuspension buffer.
Filter the final resuspension through an appropriately sized filter for the cell type. Commonly used mesh sizes for single-cell suspensions are between 40 and 70 μm.
Count cells, bring concentration to 1,000–1,500 cells per microliter, and proceed to encapsulation.
For tissues
Add 10 µL of 1 M Tris-HCl pH 7.5. Then, mix thoroughly on a vortex for 2–3 s followed by a 15 min incubation at room temperature.
Centrifuge at 500× g for 20 s or for 5–10 s in a mini spinner and remove supernatant.
Add 1 mL of 200 μg/mL Liberase in PBS.
Incubate the tissue with digestion buffer at 37 °C for 30 min with agitation at 800 rpm on a ThermoMixer C with heated lid.
Note: The length of digestion may need to be optimized according to tissue type.
Pipette to mix the sample 5–10 times every 15 min to facilitate digestion.
After digestion is completed, filter the sample through a 70 μm filter and into a 15 mL falcon tube to remove coarse debris.
Add 10 mL of ice-cold PBS and mix well.
Centrifuge the sample at 500× g for 5 min in a pre-cooled centrifuge with swinging bucket rotor.
Discard the supernatant and resuspend the pellet again in 10 mL of ice-cold PBS with 1% BSA. Mix well.
Repeat steps D2g–D2h for three washes.
Filter cells with an appropriate filter depending on cell size and resuspend the cells in 0.5–1 mL cold PBS with 1% BSA. Count cells, bring the concentration to 1,000–1,500 cells per microliter, and proceed to encapsulation.
Note: For maximal RNA integrity, add 0.2–1 unit/µL RNase inhibitor.
Cryopreservation of samples (Optional)
Resuspend the cell pellet in 500 μL of fresh PBS or media.
Count and record the cell number.
Add an appropriate volume of chilled cryopreservation medium to obtain a cell concentration of 1–2 × 106 cells per milliliter.
Dispense cell suspension aliquots of 1–2 mL into pre-cooled cryovials and place the cryovials inside a pre-cooled cell freezing container, e.g., CoolCell FTS30, to ensure gradual freezing.
Place the cell freezing container in a -80 °C freezer for ≥4 h. After 4 h, transfer the cryovials to liquid nitrogen for long-term storage.
Thaw in a 37 °C water bath and wash cells twice with PBS + 0.5%–1% BSA before proceeding to cell encapsulation.
Validation of protocol
The current protocol has been routinely performed as a standard protocol on human, mouse, and rat tissues and primary cells at the Spatial Technologies Unit, Beth Israel Deaconess Medical Center, Harvard Medical School, being considered robust and reproducible. The whole procedure was repeated once after the completion of this manuscript by a person without previous knowledge using the current version to ensure details are correct, comprehensible, and executable.
Additional verification has been conducted in various studies, as documented, and systematically tested and optimized in Jiménez-Gracia et al. [23] (Figures 1–7) and in Aney et al. [24] (Figure 1).
Authors have validated the following sample types with successful single-cell RNA sequencing: human PBMC, mouse pancreas, mouse and rat liver, mouse lung, mouse colon, human colon biopsies, and human prostate.
General notes and troubleshooting
General notes
Tip 1: Only use freshly prepared DSP
Make single-use aliquots (20–50 μL) for 2–5 fixations. Do not re-freeze leftovers. Keeping the stock fixative away from water is key because it neutralizes the NHS-esters quickly. Currently, this can be a limitation in clinical settings, since clinical staff requires bandwidth, proper instrument setup, and training for the fixation process.
Tip 2: Avoid using Tris, glycine, or any other amine-active buffer to prepare the fixative
Avoid buffer components with primary amines such as Tris and glycine buffers, as they compete with proteins in the sample. DSP reacts with non-protonated aliphatic amine groups, including the N (amine) terminus of polypeptides and the ϵ-amino group of lysine (K) side chain.
Tip 3: Never bypass the stock preparation step by directly dissolving DSP solid powder in PBS
DSP is hydrophobic and needs to be dissolved in DMSO before being added to the aqueous reaction mixture. Besides the 50 mg (22586) size, DSP is also available in 10 × 1 mg (A35393) or 1 g (22585) packaging. Although a smaller size unit is more expensive per sample, consider using a smaller size matching the experimental scale to avoid exposure to moisture absorption. Lot-to-lot differences in DSP may exist; testing one aliquot on the optimization sample from the batch can be a QC option.
Tip 4: DSP crystal formation is normal but needs to be controlled; not all crystals are from DSP
DSP does not possess a charged group; it is lipophilic and membrane-permeable, making it suitable for intracellular and intramembrane crosslinking. However, it is water-insoluble and can form crystal precipitates when preparing the working solution. These crystals are removed by filtering in step B3. However, usage of BSA and even a high percentage of FBS are common to promote cell survival and boost viability. Still, calcium oxalate crystals can often be found in commercial fetal bovine serum (FBS) [25], being mistakenly perceived as DSP crystals. Using BSA is not concerning but a high percentage of BSA will deplete reactive DSP and lead to cloudy samples.
Tip 5: Calculate viability before fixation and re-count before storage
Viability is no longer a good measure for single-cell sample quality because DSP permeabilizes the cell membrane. Therefore, it is advisable to measure viability immediately before the fixation. Some cells will inevitably be lost during wash steps, so recounting cells before freezing and storage is necessary. Because fixed cells are not alive, no additive beyond DMSO or glycerol is required, such as FBS, ascorbic acid, or cell culture media components.
Tip 6: Pay attention to other potentially interfering components
The list here is not exhaustive and only includes common examples. With the incorporation of new components into the FixNCut workflow, caution is advised. The central disulfide bridge in DSP provides a reducible link that can be cleaved by reducing agents such as DTT (dithiothreitol) or β-mercaptoethanol. In single-cell assays, DTT is commonly used to inhibit RNase activity, inactivate reverse transcription inhibitors, and dissolve gel beads by breaking their disulfide bonds, thus releasing oligonucleotides essential for mRNA capture. However, DTT also de-crosslinks DSP-induced fixation. DTT, often used with sodium dodecyl sulfate (SDS), can rapidly break down cells. Therefore, even fixed cells should not be left in the reaction mix for more than a few min before chip loading. In contrast, EDTA in cell sorting buffer does not interfere with the crosslinking process. Once cells are fixed, EDTA cannot alter intracellular protein structure through chelating divalent cations or prevent cell clumping by inhibiting calcium-dependent adhesion, making its use in FACS buffer non-essential.
Final remark: As a reactive compound, DSP esters can cause eye and skin irritation and may be harmful if swallowed or inhaled. Therefore, it should be handled with care in a controlled laboratory environment. Always follow safety guidelines when handling chemicals, including wearing appropriate personal protective equipment and working in a well-ventilated area. The waste generated in this protocol is inactive but contains trace contents of DMSO and should be disposed of according to local regulations.
Acknowledgments
This project has received funding from the Innovative Medicines Initiative 2 Joint Undertaking (IMI 2 JU) under grant agreement No 831434 (3TR; Taxonomy, Targets, Treatment, and Remission). The JU receives support from the European Union’s Horizon 2020 research and innovation program and EFPIA. Also, this project has received funding from the European Union’s H2020 research and innovation program under grant agreement No. 848028 (DoCTIS; Decision On Optimal Combinatorial Therapies In Imids Using Systems Approaches), and the Commonwealth Standard Grant Agreement 4-F26M8TZ. L.J.-G. has held an FPU PhD fellowship (FPU19/04886) from the Spanish Ministry of Universities. Australian Prostate Cancer BioResource (APCB) collection in Adelaide is supported by funding from the South Australian Immunogenomics Cancer Institute and the South Australian Health and Medical Research Institute. S.W., A.A.A. and I.S.V. acknowledge support from Spatial Technologies Unit of Precision RNA Medicine Core (RRID:SCR_024905), as well as the National Institutes of Health under award number P01AI179405 and U54HL165440. The authors further thank Dr. Tatsuyuki Sato and Dr. Joji Fujisaki for their valuable feedback and discussion. The graphics were created with BioRender.com.
Competing interests
H.H. is a co-founder and shareholder of Omniscope, a scientific advisory board member of MiRXES and Nanostring, and a consultant to Moderna and Singularity. L.G.M is an advisor and shareholder of Omniscope, and advisor for ArgenTAG and BioScryb. Omniscope has filed a patent related to the application of the FixNCut protocol. All other authors declare no competing interests.
References
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Article Information
Publication history
Received: Jun 1, 2024
Accepted: Jul 28, 2024
Available online: Aug 13, 2024
Published: Sep 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Cell Biology > Single cell analysis
Molecular Biology > DNA > DNA sequencing
Cell Biology > Cell isolation and culture > Cryopreservation
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Can you cryoperserve after DSP fixation and quenching of tissue?
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Received: Jul 19, 2024
Published: Aug 5, 2024
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After official publication in Bio-protocol (https://bio-protocol.org/e5018), we received a question regarding our publication titled "Flow Cytometry Analysis of Microglial Phenotypes in the Murine Brain During Aging and Disease" (DOI: 10.21769/BioProtoc.5018) that highlighted a significant typo in our writing. The published paper states, "Prepare Enzyme Mix 2: Buffer Y (20 μL/sample) + Enzyme A (100 μL/sample)." The volume of Enzyme A should read 10 μL/sample and not 100 μL/sample. This volume has been corrected in the updated version along with other minor edits that were made while reviewing to improve clarity of the protocol. Changes are tracked in the attached version linked here.
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The following supporting information can be downloaded here.
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Received: Jul 13, 2024
Published: Aug 5, 2024
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In Vitro GT-array (i-GT-ray), a Platform for Screening of Glycosyltransferase Activities and Protein–Protein Interactions
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DOI: 10.21769/BioProtoc.5066 Views: 383
Reviewed by: David PaulWeidong An Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Mar 2024
Abstract
Progress in bioinformatics has facilitated the identification of a large number of putative glycosyltransferases (GTs) associated with many physiological processes. However, many of these GTs remain with unknown biochemical function due to numerous technical limitations. One of these limitations is the lack of innovative tools for large-scale screening of enzyme activity in vitro and testing protein–protein interactions (PPIs) between GT partners. Currently, testing the enzyme activity of a protein requires its production in a heterologous expression system and purification before enzyme assays, a process that is time-consuming and not amenable to high-throughput screening. To overcome this, we developed a platform called in vitro GT-array (i-GT-ray). In this platform, 96-well microplates are coated with plasmid DNA encoding for tagged GTs and a capture antibody. Tagged GTs are produced from plasmid DNA via a cell-free in vitro transcription/translation (IVTT) system and captured through the anti-tag capture antibody directly on microplates. After washing to remove IVTT components, the captured enzymes can be considered purified, and their activity can be tested directly on microplates. The whole process can be performed in less than two days, compared to several weeks for currently available screening methods. The i-GT-ray platform has also been adapted to investigate PPIs between GTs. Here, we provide a practical user guide for the preparation of GT-arrays coated with plasmid DNA and a capture antibody that can be used for monitoring enzyme activity and PPIs of GTs in a high-throughput manner.
Key features
• Synthesis of tagged proteins directly from plasmid DNA, which are captured by anti-tag antibody attached to microplates.
• Captured tagged proteins can be considered as purified proteins ready for enzyme assays.
• Our platform can be used for high-throughput screening of enzyme activity and protein–protein interactions in vitro in a short time.
• Our platform can be used for biochemical characterization of difficult proteins such as membrane-integrated glycosyltransferases.
• Our platform can be adapted to downstream analytical methods such as mass spectrometry (i.e., DPS-MS).
Keywords: Protein synthesis NAPPA In vitro GT-array (i-GT-ray) Protein–protein interaction Glycosyltransferases
Graphical overview
Background
Advancements in high-throughput genomics technologies, including DNA sequencing, DNA microarrays, RNA-Seq, and proteomics, have greatly facilitated the identification of numerous genes across various species. However, methods for high-throughput biochemical analyses of proteins encoded by these genes are lacking [1]. Determining the enzyme activity of a protein can be achieved through direct testing in vitro or indirectly via genetic complementation of mutants, which are poorly adapted for high throughput. Currently, characterizing the biochemical function of an enzyme involves expression in heterologous organisms that typically lack that protein, followed by measurement of enzyme activity of solubilized and partially purified protein [2–4]. This approach has several drawbacks, including high cost, low yield, time consumption, and limited applicability to high-throughput analyses.
The development of protein microarrays requires the production of substantial quantities of purified proteins for binding onto a solid support and necessitates specialized instrumentation, such as microarray printers and scanners [5–7]. A recent advancement in protein microarray technology, called nucleic acid programmable protein array (NAPPA), has facilitated the efficient generation of protein microarrays through in vitro cell-free transcription/translation (IVTT) systems directly from plasmid DNA [8]. Currently, NAPPA is primarily employed in screening for protein–protein interactions (PPIs) [5,9–12]. We have recently developed and validated a NAPPA-based platform for enzyme assays of glycosyltransferases (GTs) named in vitro GT-array (i-GT-ray) [13,14]. Additionally, it enables the study of PPIs of GTs [15] in a high-throughput manner. Most eukaryotic GTs are difficult to characterize biochemically through conventional methods because they are integral membrane proteins [16] and are present (along with their corresponding mRNAs) in very low quantities within the cell. These characteristics pose challenges in production and purification [16,17]. Interestingly, in this platform, all GTs tested during validation work could be produced in a soluble and active form even though the cell-free coupled IVTT systems do not contain any lipids or detergent (membrane mimic). The i-GT-ray platform utilizes 96-well microplates pre-coated with plasmid DNA-encoding tagged proteins and anti-tag capture antibody (CAb). This enables simultaneous protein production using IVTT systems and protein capturing via the CAb on microplates (Figure 1A and B). Following washing, the captured GTs can be screened for enzyme activity directly on microplates (Figure 1C). The products resulting from transferase reactions can be detected using mass spectrometry or the GLO system directly on microplates (Figure 1D). This unique platform facilitates the rapid and effective creation of GT arrays, significantly reducing the time required for enzyme activity screening. In this manuscript, we provide a step-by-step protocol for the preparation of GT arrays, which can be utilized for in vitro enzyme activity screening and PPIs of GTs. The potential pitfalls of using full-length membrane integral proteins are discussed.
Figure 1. Schematic representation illustrates the fundamental principle of the i-GT-ray platform. (A) The tagged glycosyltransferases (GTs) are simultaneously synthesized and attached on microplate wells using plasmid DNA and anti-tag capture antibody. (B) After washing, the captured tagged GTs are considered purified. (C) Transferase reactions can be carried out by adding nucleotide diphosphate (NDP) sugars and acceptors. (D) The monitoring of transferase reactions is achieved through various methods, including the UDP/GDP GLO system, radioactive assays, or DPS-MS.
Materials and reagents
Biological materials
Chemically competent E. coli cloning strain (e.g., One ShotTM TOP10 Chemically Competent E. coli, Invitrogen, catalog number: C404010 or equivalent)
Expression vector pJFT7_nHALO_DC (Ampicillin resistant, DNASU Plasmid Repository, Clone ID: EvNO000424503) and pANT7_cGST (Ampicillin resistant, DNASU Plasmid Repository, Clone ID: EvNO00023103)
GatewayTM donor vector (non-ampicillin resistant, e.g., pCRTM8/GW/TOPOTM TA, Fisher Scientific, catalog number: K250020)
LR ClonaseTM II enzyme mix (Invitrogen, catalog number: 11791-020)
High-fidelity polymerase (e.g., KAPA HiFi DNA Polymerase, Roche Sequencing and Life Science, catalog number: KK2102)
Taq DNA polymerase (New England Biolabs, catalog number: M0273L)
Template for full-length protein-coding sequences (CDS) of interest or a CDS clone
1-Step Human Coupled IVT Kit, DNA (Thermo Scientific, catalog number: 88882)
TnT® Quick Coupled Transcription/Translation System (Promega, catalog number: L1170)
Anti-GST antibody (Cytiva, catalog number: 27457701)
Anti-HaloTag® pAb (Promega, catalog number: G9281)
Secondary mouse anti-goat IgG HRP conjugated (Santa Cruz Biotechnology, catalog number: sc-2354)
Secondary anti-rabbit IgG HRP conjugated (Promega, catalog number: W4011)
PfoI restriction enzyme (Thermo Scientific, catalog number: ER1751)
Streptavidin (Thermo Scientific, catalog number: 434302)
Reagents
Gel-purification kit (Zymo Research, catalog number: D4007)
DNA Clean and concentratorTM (Zymo Research, catalog number: D4004)
LB broth (Fisher Scientific, catalog number: BP1426-2)
Agar (Research Products International, catalog number: A20020-100.0)
Agarose ME (Research Products International, catalog number: A20085-500.0)
Ampicillin (Sigma-Aldrich, CAS number: 69-52-3)
Spectinomycin (Research Products International Corp., catalog number: 22189-32-8)
NucleoBond Xtra Midi kit (MACHEREY-NAGEL, catalog number: 740420.10)
2× Laemmli sample buffer (Bio-Rad, catalog number: 1610737EDU)
SuperSignalTM West Femto maximum sensitivity substrate (Thermo Scientific, catalog number: 34096)
Tween-20 (Sigma-Aldrich, CAS number: 9005-64-5)
EDC [1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride] (Thermo Scientific, catalog number: 22980)
EZ-LinkTM Hydrazide-Biotin (Thermo Scientific, catalog number: 21339)
Non-fat dried milk bovine (Sigma-Aldrich, catalog number: M7409-1BTL)
Acrylamide/bis 37.5:1 (Bio-Rad, catalog number: 1610158)
Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: 7727-54-0)
TEMED (Bio-Rad, catalog number: 1610801)
Imidazole (Sigma-Aldrich, CAS number: 288-32-4)
ZebaTM Micro Spin desalting columns, 7 K MWCO (Thermo Scientific, catalog number: 89877)
AccuBlue Next Gen dsDNA Quantitation kit (Biotium, catalog number: 31060-T)
UDP-GLO Kit (UDP-GloTM Glycosyltransferase Assay) (Promega, catalog number: V6961)
GDP-GLO Kit (GDP-GloTM Glycosyltransferase Assay) (Promega, catalog number: VA1090)
DOWEX 1X8-100 resin (Cl) (Fisher Scientific, catalog number: AAL142570B)
Ethanol (Fisher Chemical, catalog number: A962-4)
ScintiVerseTM BD Cocktail (ScintanalyzedTM) (Fisher Chemical, catalog number: SX18-4)
Methanol (Fisher Scientific, catalog number: A434-20)
Tris-HCL (Sigma-Aldrich, catalog number: 1185-53-1)
Tris-Base (Sigma-Aldrich, catalog number: 77-86-1)
EDTA (Sigma-Aldrich, catalog number: 60-00-4)
Glycine (Bio-Rad, catalog number: 1610718)
HCl (Fisher Scientific, catalog number: A144-50)
SDS (Vivantis, catalog number: 151-21-3)
NaCl (Sigma-Aldrich, catalog number: 7647-14-5)
KCl (Sigma-Aldrich, catalog number: 7447-40-7)
NaOH pellets (Fisher Scientific, catalog number: S318-100)
Na2HPO4 (Fisher Scientific, catalog number: P285-500)
NaH2PO4 (Fisher Scientific, catalog number: S381-500)
KH2PO4 (Fisher Scientific, catalog number: S375-212)
Na2CO3 (Sigma-Aldrich, catalog number: 497-19-8)
NaHCO3 (Sigma-Aldrich, catalog number: 144-55-8)
DMSO (Sigma-Aldrich, catalog number: 67-68-5)
MgCl2 (Sigma-Aldrich, catalog number: 7786-30-3)
MnCl2 (Sigma-Aldrich, catalog number:7773-01-5)
Solutions
LB medium (1 L) (see Recipes)
100 mg/mL antibiotic stock (Ampicillin or Spectinomycin) (10 mL) (see Recipes)
LB agar plate with antibiotics (1 L) (see Recipes)
SDS-PAGE gel working solutions (see Recipes)
SDS-PAGE running buffer (1 L) (see Recipes)
SDS-PAGE transfer buffer (1 L) (see Recipes)
100 mM phosphate buffer pH 7.2 (1 L) (see Recipes)
10 mM phosphate buffer pH 7.2 (1× PBS) (1 L) (see Recipes)
10 mM phosphate buffer pH 7.2 (1× PBS) containing 5% (w/v) fat-free milk (100 mL) (see Recipes)
10 mM phosphate buffer pH 7.2 (1× PBS) containing 0.05% (v/v) Tween 20 (1 L) (see Recipes)
10 mM phosphate buffer pH 7.2 (1× PBS) containing 5% (w/v) fat-free milk and 0.05% (v/v) Tween 20 (100 mL) (see Recipes)
10 mM phosphate buffered saline with EDTA (PBS with EDTA) (1 L) (see Recipes)
Sodium bicarbonate buffer (pH 9.6) (see Recipes)
50 mM biotin hydrazide (1 mL) stock (see Recipes)
2.5 mM biotin hydrazide (1 mL) working solution (see Recipes)
100 mM Tris-HCl buffer (pH 7.2) (see Recipes)
Recipes
LB medium (1 L)
Reagent Final concentration Quantity
LB broth 25 g/L 25 g
ddH2O n/a ~980 mL
Total n/a 1,000 mL
Adjust the pH to 7.4 with NaOH. Autoclave at 121 °C for 15 min. Store at RT.
100 mg/mL antibiotic stock (ampicillin or spectinomycin)
Reagent Final concentration Quantity
Ampicillin or Spectinomycin 100 mg/mL 1 g
ddH2O n/a 10 mL
Total n/a 10 mL
Sterilize the solution with a 0.2 μm syringe filter. Store at -20 °C.
LB agar plate with antibiotics (1 L)
Reagent Final concentration Quantity
LB broth 25 g/L 25 g
Agar 15 g/L 15 g
Antibiotic stock (100 mg/mL) 100 μg/mL 1 mL
ddH2O n/a ~960 mL
Total n/a 1,000 mL
Adjust the pH to 7.4 with NaOH. Add agar after adjusting pH. Autoclave at 121 °C for 15 min. Allow to cool to 35–45 °C. Add 1 mL of 100 mg/mL antibiotic solution and mix well. Transfer 25 mL of the solution to petri dishes. Allow the agar plates to cool and dry in a laminar flow hood. Store the plates at 4 °C.
SDS-PAGE gel working solutions (for three 1.0 mm gels)
Separation gel (10%)
Reagent Quantity
ddH2O 6.7 mL
30% acrylamide/bis 37.5:1 5.3 mL
Lower solution (1.5 M Tris-HCl pH 8.8 and 0.2% SDS) 4 mL
Persulfate (10% Sol) 150 μL
TEMED 10 μL
Stacking gel
Reagent Quantity
ddH2O 6 mL
30% acrylamide/bis 37.5:1 1.5 mL
Upper solution (1 M Tris-HCl pH 6.8 and 0.2% SDS) 2.5 mL
Persulfate (10% Sol) 40 μL
TEMED 10 μL
SDS-PAGE running buffer (1 L)
Reagent Final concentration Quantity
Tris 30 mM 3.03 g
Glycine 200 mM 14.5 g
SDS 0.1% (w/v) 1 g
ddH2O n/a ~980 mL
Total n/a 1,000 mL
Store at room temperature (RT).
SDS-PAGE transfer buffer (1 L)
Reagent Final concentration Quantity
Tris 30 mM 2.525 g
Glycine 200 mM 13.824 g
ddH2O n/a ~780 mL
Methanol 20% (w/v) 200 mL
Total n/a 1,000 mL
Adjust the pH to approximately 8.3 after adding Tris and glycine in ~780 mL of ddH2O. Then, add 200 mL of 100% methanol and add ddH2O to 1 L. Store buffer at RT.
100 mM phosphate buffer pH 7.2 (1 L)
Reagent Final concentration Quantity
NaCl 1.36 M 80 g
KCl 26.83 mM 2 g
Na2HPO4 141.96 mM 14.4 g
KH2PO4 17.64 mM 2.4 g
ddH2O n/a ~900 mL
Total n/a 1,000 mL
Adjust pH to approximately 7.2. Autoclave at 121 °C for 35 min. Store at 4 °C.
10 mM phosphate buffer pH 7.2 (1× PBS) (1 L)
Reagent Final concentration Quantity
100 mM phosphate buffer pH 7.2 10 mM 100 mL
ddH2O n/a ~900 mL
Total n/a 1,000 mL
Adjust pH to 7.2. Store buffer at 4 °C.
10 mM phosphate buffer pH 7.2 (1× PBS) containing 5% (w/v) fat-free milk (100 mL)
Reagent Final concentration Quantity
100 mM phosphate buffer pH 7.2 10 mM 10 mL
Fat-free milk 5% 5 g
ddH2O n/a ~90 mL
Total n/a 1,00 mL
Adjust pH to 7.2 for 1× PBS, add fat-free milk, and mix for 1 h. Store buffer at 4 °C.
10 mM phosphate buffer pH 7.2 (1× PBS) containing 0.05% (v/v) Tween 20 (1 L)
Reagent Final concentration Quantity
100 mM phosphate buffer pH 7.2 10 mM 100 mL
Tween 20 0.05% (v/v) 0.5 mL
ddH2O n/a ~900 mL
Total n/a 1,000 mL
Adjust pH to 7.2 for 1× PBS, add Tween 20, and mix for 1 h. Store buffer at 4 °C.
10 mM phosphate buffer pH 7.2 (1× PBS) containing 5% (w/v) fat-free milk and 0.05% (v/v) Tween 20 (100 mL)
Reagent Final concentration Quantity
100 mM phosphate buffer pH 7.2 10 mM 10 mL
Tween 20 0.05% (v/v) 0.05 mL
Fat-free milk 5% 5 g
ddH2O n/a ~99 mL
Total n/a 100 mL
Adjust pH to 7.2 for 1× PBS, add Tween 20 and fat-free milk, and mix for 1 h. Store buffer at 4 °C.
10 mM phosphate buffered saline with EDTA (PBS with EDTA) (1 L)
Reagent Final concentration Quantity
NaCl 150 mM 8.8 g
EDTA (0.5 M, pH 8.0) 10 mM 20 mL
Na2HPO4 6.7 mM 0.95 g
NaH2PO4 3.34 mM 0.4 g
ddH2O n/a ~950 mL
Total n/a 1,000 mL
Adjust the pH to approximately 7.2. Autoclave at 121 °C for 35 min. Store at 4 °C.
Sodium bicarbonate buffer (pH 9.6)
Reagent Final concentration Quantity
Na2CO3 15 mM 1.59 g
NaHCO3 34.87 mM 2.93 g
ddH2O n/a ~900 mL
Total n/a 1,000 mL
Adjust the pH to approximately 9.6. Autoclave at 121 °C for 35 min. Store at 4 °C.
50 mM biotin hydrazide (1 mL) stock
Reagent Final concentration Quantity
Hydrazide-Biotin 50 mM 12.917 mg
DMSO n/a 1 mL
Total n/a 1 mL
Store the solution at -20 °C.
2.5 mM biotin hydrazide (1 mL) working solution
Reagent Final concentration Quantity
Hydrazide-biotin (50 mM) 2.5 mM 5 mL
0.1 M imidazole, pH 6 n/a 995 mL
Total n/a 1 mL
100 mM Tris-HCl buffer (pH 7.2)
Reagent Final concentration Quantity
Tris-HCl 100 mM 12.14 g
MgCl2 2 mM 0.19 g
MnCl2 2 mM 0.252g
ddH2O n/a ~900 mL
Total n/a 1,000 mL
Adjust the pH to approximately 7.2. Sterilize the solution by filtration with a 0.2 μm syringe filter. Store at 4 °C.
Laboratory supplies
Pipette tips 0.5–10 µL, 20–200 µL, 100–1,000 µL (Fisher Scientific, FisherbrandTM, catalog numbers: 02-707-438, 02-707-417, 02-707-403)
1.5 mL Eppendorf tubes (Fisher Scientific, FisherbrandTM, catalog number: 02-681-321)
2 mL Eppendorf tubes (Fisher Scientific, FisherbrandTM, catalog number: 02-681-320)
Liquid scintillation vials (DWK Life Sciences, catalog number: 986730)
0.2 mL PCR tubes (Fisher Scientific, Thermo ScientificTM, catalog number: AB-0620)
100 mm × 15 mm Petri dishes (Fisher Scientific, FisherbrandTM, catalog number: FB0875712)
14 mL culture tubes (Globe Scientific, catalog number: 110178)
0.2 μm sterile syringe filter (e.g., Corning, catalog number: 431227)
Immobilon membrane (Millipore Sigma, catalog number: IPFL85R)
96-well microplate (clear, flat bottom, half area, high binding, polystyrene) (Corning, catalog number: 3690)
96-well microplate (flat bottom, half area, non-binding, white polystyrene) (Corning, catalog number: 3642)
Whatman® cellulose chromatography papers (Whatman, catalog number: 3017-820)
Equipment
250 mL Erlenmeyer flask
Micropipettes 0.1–2.5 µL, 2–20 µL, 20–200 µL (Eppendorf, models: 3123000217, 3123000233, 3123000250)
Multichannel pipettes (8 channels) 0.1–10 µL, 10–100 µL (Eppendorf, models: 2231300043, 2231300045)
Benchtop microcentrifuge (Beckman Coulter, model: Microfuge 22R centrifuge, catalog number: 8043-30-1145)
PCR thermocycler (Applied biosystems, Veriti 96-well Thermal Cycler 2990215201, catalog number: 4375305)
Incubation shaker (New Brunswick Scientific, Classic series, model: M1247-0004, catalog number: 19525)
Incubator (Fisher Scientific, Thermo ScientificTM, model: HerathermTM IGS60, catalog number: 51028063)
Mini-PROTEAN Tetra vertical electrophoresis system (Bio-Rad, catalog number: 1658002EDU)
Gel electrophoresis tank (Bio-Rad Laboratories, model: Wide Mini-Sub®, catalog number: 1704468)
Gel electrophoresis power supply (Bio-Rad Laboratories, model: PowerPacTM Basic, catalog number: 1645050)
Gel imager (Bio-Rad Laboratories, model: Gel DocTM XR+, catalog number: 1708195)
Plate reader (BioTek Synergy/HTX multi-mode reader, model: S1LFA, catalog number: BTS1LFA)
ChemiDocTM imaging system (Bio-Rad Laboratories, Gel Doc EZ, catalog number: 1708270)
Multi-purpose scintillation counter (Beckman, model: LS6500, SKU: 8043-30-1194)
Freeze dry system (Labconco, model: 4.5 FreeZone, catalog number: 7750020)
Vortex
Water bath
Software and datasets
SnapGene Viewer v.7.1.1
Gen5 v.2.09.2
Microsoft® Excel V 16.86
Procedure
Part 1: i-GT-ray platform for efficient screening of enzyme activity of glycosyltransferases (Figure 2)
Figure 2. Schematic overview of enzyme assays performed on i-GT-ray platform. (A) Microplates can be pre-coated with anti-tag capture antibody (CAb) only to capture tagged glycosyltransferases (GTs) synthesized from plasmid DNA in an Eppendorf tube using in vitro transcription/translation (IVT) system. (B) Microplates can be pre-coated with both CAb and streptavidin to which the biotinylated plasmid DNA is attached. The IVT system is then added to wells for synthesis of tagged GTs that are captured by CAb. After washing, the captured tagged GTs are considered purified and transferase reactions are carried out by adding NDP-sugars and acceptors.
I. Cloning of protein coding sequences (CDSs)
Cloning of CDSs into a gateway donor vector
Note: After the PCR, two forms of CDS are created: one with a stop codon, and one without a stop codon. Therefore, for the PCR process, the primers are designed accordingly. The reverse primers are designed with and without a stop codon to ensure the generation of both forms of the CDSs.
The CDSs are PCR-amplified using a high-fidelity polymerase (e.g., KAPA HiFi DNA Polymerase or Phusion polymerase).
Purify the fragment from an agarose gel using a standard kit (e.g., Zymoclean Gel DNA Recovery Kits).
Add extra As to the CDSs using a reaction made of reagents listed in Table 1.
Table 1. Conditions for addition of As to the CDSs.
Component Final concentration Quantity
10× ThermoPol reaction buffer 1× 1 µL
10 mM dNTPs 500 mM 0.5 µL
Purified CDSs PCR DNA 25 ng/mL Variable (250 ng)
Taq DNA polymerase 2.5 units 0.5 µL
ddH2O n/a ~10 µL
Total n/a 10 µL
Incubate at 72 °C for 25 min in a PCR machine.
Create an entry clone by transferring the CDS to the GatewayTM donor vector (e.g., pCRTM8/GW/TOPOTM TA) through TA cloning following the manufacturer’s protocol.
Introduce the constructs into the chemically competent E. coli cloning strain (e.g., DH5a) using the heat-shock method.
Spread the transformed competent bacteria cells onto LB plates containing 100 mg/L spectinomycin and incubate overnight at 37 °C to form colonies.
Screen the colonies and confirm the reading frame, correct orientation, and full sequence of the inserted CDS through Sanger sequencing.
Cloning of CDSs into an expression vector
Perform Gateway cloning from pCRTM8/GW/TOPOTM entry vector to pJFT7_nHALO_DC or pANT7_cGST destination vectors using LR ClonaseTM II enzyme mix following the manufacturer’s protocols.
Note: For the creation of the N-terminal Halo-Tagged GT proteins, an entry vector containing the full-length GT gene with a stop codon is required. However, to generate the C-terminal GST-tagged GT proteins, an entry vector containing the full-length GT gene without a stop codon is necessary.
Transfer the constructs into the chemically competent E. coli cloning strain by the heat-shock method.
Spread the transformed competent cells onto LB plates containing 100 mg/L ampicillin and incubate overnight at 37 °C to form colonies.
Screen the colonies by PCR and confirm the reading frame, correct orientation, and full sequence of the insert through Sanger sequencing.
The correct colony is cultured in 200 mL of LB liquid medium supplemented with 100 mg/L ampicillin. Grow at 37 °C with shaking at 180 rpm until the OD600 value reaches 0.6–1.0. Then, extract and purify large amounts of plasmid using the NucleoBond Xtra Midi kit following the manufacturer’s protocol.
II. Recombinant protein production and analysis
Produce the tagged GT fusion proteins either in 1.5 mL Eppendorf tubes or on 96-well microplates using 1 µg of GT plasmid DNA and an expression kit (e.g., 1-Step Human Coupled IVT Kit, Thermo Scientific) according to the manufacturer’s recommendations at 30 °C for 6 h.
Confirm the production of Halo-GT and GT-GST fusion proteins by running western blotting as follows:
Transfer 4 µL of expressed protein to Eppendorf tubes, add 4 µL of 2× Laemmli sample buffer, and denature the proteins at 100 °C for 10 min.
Prepare an SDS-PAGE gel using separating gel and stacking gel solutions.
Run the proteins on SDS-PAGE gel using running buffer. After running is completed, transfer the proteins onto Immobilon membranes using the transfer buffer in the Mini-Protein Tetra cell system (Bio-Rad) for 80–90 min, maintaining the cold environment using the ice pack (or place in a cold room).
After transfer, rinse the membrane with 1× PBS and proceed to blocking the membrane with 5% (w/v) fat-free dry milk in 1× PBS at 4 °C overnight (or at least 1 h at room temperature).
Wash the membrane with 1× PBS containing 0.05% (v/v) Tween 20 three times for 10 min each with gentle shaking (approximately 50 rpm).
Apply the primary antibody (anti-GST or anti-Halo) at 1:10,000 dilution in 1× PBS containing 5% (w/v) fat-free milk with 0.05% (v/v) Tween 20 for 2 h with gentle shaking (approximately 50 rpm) at room temperature.
Wash the membrane with 1× PBS containing 0.05% (v/v) Tween 20, three times for 10 min each with gentle shaking (approximately 50 rpm).
Apply the corresponding secondary antibody (anti-goat or anti-rabbit) at 1:15,000 dilution in 1× PBS containing 5% (w/v) fat-free milk with 0.05% (v/v) Tween 20 for 1 h with gentle shaking (approximately 50 rpm) at room temperature.
Wash the membrane with 1× PBS containing 0.05% (v/v) Tween 20, three times for 10 min each with gentle shaking (approximately 50 rpm).
Wash the membrane with 1× PBS (without Tween 20), two times for 10 min each with gentle shaking (approximately 50 rpm).
Detect the presence of the fusion protein using SuperSignal West Femto Maximum Sensitivity Substrate according to the manufacturer’s protocols and observing the membrane in the ChemiDocTM Imaging system (Bio-Rad).
III. Coating of microplates with capture antibody (CAb) with or without plasmid DNA
Coating of microplate with CAb alone
Apply 50 µL of CAb (anti-GST or anti-Halo antibody) at a 1/200 dilution in 50 mM sodium bicarbonate buffer, pH 9.6, to the 96-well plate and incubate at 4 °C overnight (or at least 1 h at room temperature).
Wash the wells with 1× PBS three times, each time for 10 min.
Block the wells with 5% fat-free dry milk in 1× PBS at 4 °C for 4–6 h.
Wash the wells three times with 1× PBS, each time for 10 min. These coated microplates can be used immediately or stored at 4 °C for up to 60 days (without liquid).
Coating of microplates with both CAb and plasmid DNA
DNA linearization and biotinylation
For linearization of plasmid DNA:
• Use approximately 60 µg of plasmid DNA (containing GT with tag) using PfoI restriction enzyme according to the manufacturer’s recommendations.
Note: During linearization of plasmid DNA, choose the restriction site far from the gene of interest (at least 1,000 bp).
• Freeze-dry the linearized plasmid DNA and resuspend in 10 μL of water to achieve a concentration of 6 mg/μL.
Note: For freeze-drying of plasmid DNA, the linearized plasmid DNA solution in water is frozen rapidly in a -80 °C freezer or by immersing in liquid nitrogen. Then, the frozen sample is placed in the freeze-dryer after it is pre-cooled and the vacuum system is functioning properly. The freeze-drying process is carried out according to the manufacturer's instructions. The sample is placed in the freeze-dryer until the samples are thoroughly dried. They should appear as a dry, powdery material.
For biotinylation of plasmid DNA:
• Mix 7.5 µL (40–50 µg) of the linearized plasmid DNA with ~1.25 mg (6.52 μmol) of the linker EDC (1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride) and then immediately add 5 μL of biotin hydrazide working solution (2.5 mM in 100 mM imidazole).
• Vortex the mixture until the contents are dissolved completely and add 20 μL of 100 mM imidazole pH 6 after a short spin (to collect the solution to the bottom of the tube); then, incubate overnight at 37 °C in a water bath.
• Remove the non-reacted EDC and its by-products using spin desalting column (ZebaTM Spin Desalting Column) with PBS with EDTA (Recipe 12), following the manufacturer’s recommendations.
• Store the biotinylated DNA plasmid at -20 °C until use.
Coating of microplate with plasmid DNA followed by CAb
First, coat microplates with streptavidin by applying 50 µL of 50 mM sodium bicarbonate buffer, pH 9.6 containing 2 µg of streptavidin and incubating overnight at 4 °C.
Wash the microplate wells with 1× PBS three times, each for 10 min.
Apply 50 µL of 1× PBS buffer containing CAb (anti-GST or anti-Halo antibody) at 1/200 dilution to the well and incubate at 4 °C for 4 h.
Wash the microplate wells with 1× PBS three times, each for 10 min.
Block the wells with 5% fat-free dry milk in 1× PBS for 2–4 h at 4 °C.
Wash the microplate wells with 1× PBS three times, each for 10 min.
Apply 1 µg of linearized and biotinylated DNA plasmid (prepared in step III.B.1.a beforehand) dissolved in 50 µL of 5% (w/v) fat-free dry milk in 1× PBS to the wells and incubate overnight at 4 °C.
Wash the wells with 1× PBS three times, each for 10 min. These microplates can be used immediately or stored at 4 °C for up to 30 days (without liquid).
Note: To estimate the amount of plasmid DNA attached to each well, use the AccuBlue Next Gen dsDNA quantitation kit (Biotium) according to the manufacturer’s protocols.
IV. Binding of tagged GT fusion proteins
For microplates pre-coated with CAb only (Figure 2A)
Note: Proteins are synthesized in 1.5 mL Eppendorf tubes using in vitro transcription/translation kit according to the manufacturer’s recommendations at 30 °C for 6 h and then transferred to the microplates pre-coated with CAb. For synthesis of fusion protein, 25 µL of in vitro transcription/translation mixture is carried out containing 1 µg of plasmid DNA according to the manufacturer’s recommendations.
Add 50 µL of protein mixture [12.5 µL of tagged proteins + 37.5 µL of 1× PBS containing 5% (w/v) fat-free milk] to microplates pre-coated with CAb.
Incubate at 14 °C for 4–6 h with shaking at 250 rpm.
Wash the microplates three times with 1× PBS, each time for 10 min.
Perform transferase assays according to the procedure outlined below (section V).
For microplates pre-coated with CAb and plasmid DNA (Figure 2B)
Note: Proteins are synthesized directly on microplates pre-coated with CAb and plasmid DNA using in vitro transcription/translation kit according to the manufacturer’s recommendations. The procedure involves simultaneous protein synthesis and binding.
Directly load 50 µL of in vitro transcription/translation mixture onto the wells of microplates pre-coated with CAb and plasmid DNA, following the manufacturer’s recommendations.
Incubate the microplates at 30 °C for 4 h.
Incubate the microplates at 14 °C for 4–6 h with shaking at 250 rpm.
Wash the microplates three times with 1× PBS, each for 10 min.
Perform the transferase assays according to the procedure below (section V).
V. Transferase assays on microplates
For testing transferase activity using non-radioactive NDP-sugars
Note: The standard transferase assays are conducted in 50 µL volume utilizing NDP-sugars and acceptors (oligosaccharides and polysaccharides) in microplates pre-coated with fusion proteins.
In a 1.5 mL Eppendorf tube, combine the assemble transferase reactions as indicated in Table 2 and mix well before transferring to wells.
Table 2. Transferase reaction mixture using non-radioactive NDP-sugars
Reagents Final concentration Volume
Tris-HCl buffer (100 mM, pH 7.2) 50 mM 25 µL
NDP-sugar (50 mM) 3 mM 3 µL
Acceptor (10 mg/mL) 2 mg/mL 10 µL
ddH2O n/a To achieve 50 µL
Total n/a 50 µL
Transfer 50 µL of transferase reaction mix to each well containing captured tagged GTs.
Incubate the reactions at 25 °C for 90 min.
Prepare the NDP-detection reagent (GDP or UDP following manufacturer’s protocol) 10 min before transferase reactions are completed.
Transfer 25 µL of reactions from each well to a new white polystyrene 96-well microplate.
Note: If the donor used is UDP-sugar, the luminescence activity is measured using UDP-GLO kit; if the donor is GDP-sugar, then GDP-GLO kit must be used.
Add 25 µL of the pre-prepared NDP-detection reagent and mix properly (with a pipette) without creating air bubbles in the well.
Incubate in the dark for 1 h at room temperature.
Observe the luminescence activity in the microplate reader using Gen5 software with a gain setting of 100. (Figure 5 represents an example of the results obtained.)
For testing the transferase activity using the radioactive NDP-sugars
Note: The standard assays are similar to non-radioactive assays, except that the appropriate NDP-[14C] sugar is used in addition to non-radioactive NDP-sugars.
In a 1.5 mL Eppendorf tube, combine the assemble transferase reactions as indicated in Table 3 and mix well before transfer to wells.
Table 3. Transferase reaction mixture using radioactive NDP sugars
Reagents Final concentration Volume
Tris-HCl buffer (100 mM, pH 7.2) 50 mM 25 µL
NDP-Sugar (50 mM) 1 mM 1 µL
NDP-[14C]sugars (90,000–100,000) cpm/50 µL 1–5 µL
Acceptor (10 mg/mL) 2 mg /mL 10 µL
ddH2O n/a To 50 µL
Total n/a 50 µL
Transfer 50 µL of the transferase reaction mix to each well containing the captured tagged GTs.
Incubate the microplates for 90 min at 25 °C.
The detection method of the incorporated radioactive differs according to the charge of the oligosaccharide products formed. For neutral oligosaccharide products (i.e., for testing of AtXXT1), excess unused NDP-[14C] sugars are removed by chelation with DOWEX 1X8-100 resin (Cl) 1:1 (v/v). For charged oligosaccharide products (i.e., for testing of AtGUX1), excess unused NDP-[14C] sugars are removed through thin layer chromatography (TLC) according to published works [18–21]. When the acceptor used is a polysaccharide (i.e., such as xyloglucan for testing AtFUT1 and AtMUR3), the reactions are collected from wells into 2 mL Eppendorf tubes and treated as follows:
Add 1 mL of cold 70% ethanol and incubate overnight at 4 °C to precipitate the radioactive polysaccharides.
Centrifuge at 15,000× g for 15 min to collect the radioactive polysaccharides.
Rinse the pellets five times with 1 mL of cold 70% ethanol.
The incorporated radioactivity (in both oligosaccharides and polysaccharides) is measured as cpm using a LS 6500 multi-purpose scintillation counter (Beckman) by resuspension of the radioactive products in 0.3 mL of water and mixing with 3–5 mL of liquid scintillation solution in scintillation vials.
Part 2: i-GT-ray platform for screening of protein–protein interactions of glycosyltransferases (Figure 3 and Figure 4)
Figure 3. Schematic representation of the strategy used to investigate protein–protein interactions (PPIs) on microplates using NAPPA-based technology. Tagged proteins (GT1 and GT2) to be tested for PPIs are synthesized in an Eppendorf tube from their respective plasmid DNAs using in vitro IVT kit. The synthesized glycosyltransferases (GTs) are applied to microplates that are pre-coated with one of the two anti-tag capture antibodies (CAb1 or CAb2). (A) If the two GTs interact, the formed complex is detected by adding the appropriate detection antibody (DAb, anti-Halo, or anti-GST) and the appropriate HRP-linked secondary antibody. One GT acts as the prey and the second is the interacting partner (acts as a bait). (B) If there is no interaction between the two GTs, the addition of DAbs and HRP-linked secondary antibody will not result in a signal. (C) Halo-GST fusion protein is applied to the microplate pre-coated with CAb (positive control) and the microplate with no pre-coating (negative control) to assess the status of binding of CAb on the microplates. The detection of the formed complex (positive interaction) or the binding of Halo-GST fusion protein is performed via TMB colorimetric method using a microplate reader and absorbance at 450 nm.
Figure 4. Schematic representation of the strategy used to investigate protein–protein interactions (PPIs) of three GTs on microplates using NAPPA-based technology. Tagged proteins (GT1 and GT2) and untagged protein (GT3) to be tested for PPIs are synthesized in an Eppendorf tube from their respective plasmid DNAs using in vitro IVT kit. The synthesized glycosyltransferases (GTs) are applied to microplates that are pre-coated with one of the two anti-tag capture antibodies (CAb1 or CAb2). (A) If the three GTs interact, the formed complex is detected by adding the appropriate detection antibody (DAb, anti-Halo, or anti-GST) and the appropriate HRP-linked secondary antibody. One GT acts as the prey, the second is the bait, and the third GT forms the bridge between GT1 and GT2. As a control, the testing of GT1 and GT2 does not result in a signal. (B) If there is no interaction between the three GTs, the addition of DAbs and HRP-linked secondary antibody will not result in a signal. (C) Halo-GST fusion protein is applied to microplate pre-coated with CAb (positive control) and microplate with no pre-coating (negative control) to assess the status of binding of CAb on the microplates. The detection of the formed complex (positive interaction) or the binding of Halo-GST fusion protein is performed via TMB colorimetric method using a microplate reader and absorbance at 450 nm.
I. Cloning of protein coding sequences (CDSs) into expression vector
CDSs in pCRTM8/GW/TOPOTM entry vector are generated as described in Part 1, Section I.A (see above).
Gateway cloning is performed using pCRTM8/GW/TOPOTM entry vector and pJFT7_nHALO_DC or pANT7_cGST destination vectors in presence of LR ClonaseTM II enzyme mix according to manufacturer’s protocols.
Note: For creation of the N-terminal Halo-Tagged GTs, the full-length GT genes are cloned into the entry vector with a stop codon. Similarly, to generate the C-terminal GST-tagged GT protein, the full-length GT genes without a stop codon are cloned into the entry vector. However, to generate the untagged GT protein, an entry vector containing the full-length GT gene with a stop codon is gateway cloned to pANT7_cGST destination vectors.
Transformation, screening, and plasmid extraction are performed as described in Part 1, section I.B (see above).
II. Screening of protein–protein interactions (PPIs)
Coating of microplates with CAb
Coating of microplates with CAb and blocking with 5% fat-free dry milk are performed as described above (Part 1, section III.A). For PPIs testing, the microplates are pre-coated with anti-GST or anti-Halo CAb depending on the tagged GTs.
PPI assays
Note: The two tagged GTs with different tags (GST or Halo) are co-produced in vitro (Figure 3). In case more than two GTs are co-produced, the third GT should be untagged (Figure 4).
The tagged GTs to be tested are co-produced in 1.5 mL Eppendorf tubes using equal amounts of plasmid DNA and the expression kit (e.g., 1-Step Human Coupled IVT Kit, Thermo Scientific) according to the manufacturer’s recommendations at 30 °C for 6 h.
Note: For co-production of two tagged GTs, ~500 ng of each GT is used, while for co-production of three GTs, 330 ng of each of GT plasmid DNA is used.
Apply 50 µL of produced protein mixture [12.5 µL of co-expressed proteins + 37.5 µL of 1× PBS containing 5%(w/v) fat-free milk] to each well of the microplate that was pre-coated with anti-Halo or anti-GST CAb.
Incubate the microplates for 4–6 h at 14 °C with shaking at 250 rpm.
Wash the microplates three times with 1× PBS, each time for 10 min.
Apply 50 µL of detecting antibody (DAb) [1/200 dilution in 1× PBS containing 5%(w/v) fat-free milk].
Note: If the microplate is coated with anti-GST, the detection antibody should be anti-Halo, and vice versa.
Incubate at 14 °C for 2–4 h with shaking at 250 rpm.
Wash the microplates three times with 1× PBS, each time for 10 minutes.
Apply 50 µL of the secondary antibody fused to Horseradish peroxidase (HRP) [anti-goat or anti-rabbit; 1/200 dilution in 1× PBS containing 5% (w/v) fat-free milk].
Incubate the microplates for 1 h at 14 °C with shaking at 250 rpm.
Wash the microplates three times with 1× PBS containing 0.05% (v/v) Tween 20, each time for 10 min.
Wash the microplates two times with 1× PBS only, each time for 10 min.
Apply 50 μL of TMB solution (substrate for HRP) to each well.
Incubate at room temperature until the appearance of color (approximately 5 min).
Stop the reaction by adding 50 μL of 2 M H2SO4.
Take the absorbance values at 450 nm using the microplate reader (Figure 6 represents an example of the results obtained).
Data analysis
For validation of the i-GT-ray platform, we have conducted enzyme activity screening of 22 non-processive putative fucosyltransferases (FUTs) from the GT37 family against five acceptor substrates and one donor substrate (GDP-fucose). The screening involved a total of 280 assays that were performed in high-throughput manner [14]. Furthermore, we validated the i-GT-ray platform using five synthases involved in cellulose, xyloglucan, (gluco)mannan, and b-(1,3)(1,4)-mixed-linkage glucan synthesis [13]. All data are expressed as the mean and standard deviation of the mean (Figure 5).
Figure 5. Example of data collected showing the fucosyltransferase activity of eight Arabidopsis AtFUTs from GT37 family on five acceptor substrates and one donor substrate (GDP-fucose). The layout of the assays in a 96-well microplate is indicated (center). Duplicate values were obtained using a second 96-well microplate. Data were obtained as luminescence (RLU, top panel) that measures the release GDP from GDP-fucose during the transferase reaction using GDP-GLO kit. Average data analysis showed that the activity of FUTs was more than three times higher than the control without the acceptor. Data are expressed as the mean ± SD using Excel software (bottom panel). Reprinted/adapted from Bhattarai et al. [14]. © 2024 The Authors. Published by Elsevier Inc on behalf of American Society for Biochemistry and Molecular Biology. Distributed under a Creative Commons Attribution NonDerivatives NonCommercial License 4.0 (CC BY-NC-ND license). https://creativecommons.org/licenses/by-nc-nd/4.0/
For protein–protein interaction (PPI), the i-GT-ray platform was validated by conducting PPI assays between eight rice GTs from OsGT43 and OsGT47 families [15]. Each OsGT43 protein was used as a prey to probe interactions with each of the OsGT47s (using both Halo- and GST-tagged proteins in an 8 × 8 protein interaction matrix in triplicates, for a total of 192 assays) (Figure 6).
Figure 6. Example of data collected from PPI assays between eight members of OsGT43s and OsGT47s families using a 96-well microplate. Data are expressed as the mean ± SE of triplicate assays using Excel software. Reprinted/adapted from Javaid et al. [15]. © 2024 The Authors. The Plant Journal published by Society for Experimental Biology and John Wiley & Sons Ltd. Distributed under a Creative Commons Attribution NonDerivatives NonCommercial License 4.0 (CC BY-NC-ND license). https://creativecommons.org/licenses/by-nc-nd/4.0/
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Bhattarai et al. [14]. Streamlining assays of glycosyltransferases activity using in vitro GT-array (i-GT-ray) platform: Application to family GT37 fucosyltransferases. Journal of Biological Chemistry. 300:105734. (Figures 2–6)
Bhattarai et al. [13]. New Insights on Beta-Glycan Synthases Using in Vitro GT-Array (i-GT-Ray) Platform. SSRN 4753263. (Figures 1–5)
Javaid et al. [15]. Specific protein interactions between rice members of the GT43 and GT47 families form various central cores of putative xylan synthase complexes. The Plant Journal. 118: 856–878. (Figures 6 and 7)
General notes and troubleshooting
General notes
Despite this protocol being validated for GTs of plant cell wall, it could be applicable to other enzymes.
The in vitro biochemical characterization and PPI are technically challenging; therefore, all steps in this protocol must be carefully attended. Optimization of these steps contributes to the success of the assay.
Before starting the enzyme and PPI assays, it is very important to confirm the synthesis of fusion protein; thereafter, it is crucial for the validation of enzyme activity and PPIs.
Before starting the PPI assay, it is important to validate that the fusion proteins are synthesized at similar levels.
The detection of the final product of enzyme assays is also equally important, since they can synthesize products at very low amounts, and it depends on the sensitivity of the detection methods.
Limitations
Although the full-length GTs (which are type-II membrane proteins with transmembrane domains) were expressed in vitro and produced soluble tagged proteins with the expected enzyme activity, it is always possible that their folding is not optimal, which may impact their enzyme activity. Therefore, efforts should be taken to minimize issues with solubility and folding of membrane proteins. i-GT-ray platform will benefit from the use of soluble and active forms of GTs and below we propose some improvements to achieve this.
Proposed improvements
We propose the following improvements using practical approaches to enhance the solubility and functional assessment of the GTs (and other integral membrane proteins):
The easiest practical approach is to design GTs without the predicted TMDs (cloning of genes encoding for GTs without TMDs). There are several publicly available bioinformatics tools that can be used to efficiently predict the location of TMDs (e.g., TMHMM, TOPCONS). The in vitro biochemical function of many non-processive GTs (type-II membrane proteins) has been determined using this approach (standard method). However, this approach is less successful for processive GTs.
The second easy practical approach is the inclusion of detergents/membrane mimics (such as 0.1%–0.4% Triton-X, CHAPS, deoxycholic acid) during the in vitro synthesis of the GTs. This can help with the proper folding and solubilization of GTs. However, the detergent concentrations must be optimized to limit the impact on the IVTT systems.
The third quick practical approach is to determine the stability of the proteins after synthesis by implementing the ThermoFluor assay, which is a temperature-based assay. In this approach, a small quantity of protein (5 mM) is mixed with SYPRO Orange dye that binds to hydrophobic patches/denatured protein/molten globules and fluoresces. As the temperature increases and the protein unfolds, it is possible to determine a melting temperature, which provides information about the stability of the proteins. The temperature and fluorescence can be monitored using a real time PCR machine [22].
The fourth approach is more complex and consists of using nanodiscs (NDs) to mimic native-like membrane environment for GTs. Nanodiscs are soluble nanoscale phospholipid bilayers used to self-assemble integral membrane proteins for enzymatic or structural studies [23] that provide an excellent solution for full-length GTs in i-GT-ray platform. In this approach, GTs are produced in the presence of phospholipids along with an encircling amphipathic helical protein belt, termed a membrane scaffold protein (MSP), which is a derivative from the human apolipoprotein 1A [24]. The MSP would allow GTs to simultaneously assemble with phospholipids into a discoidal bilayer, bringing the assay closer to what might occur in a native cell membrane. Another advantage of ND technology is the possibility of forming small (~10 nm diameter) edge-stabilized lipid bilayer discs containing a single GT or a multi-protein complex with possible control of oligomerization of GTs, which might be necessary for proper activity of some GTs. This can be achieved by using MSP of various sizes. ND technology has been used to characterize the biochemical activity of a bacterial cellulose synthase complex [25].
The fifth approach is the use of a computational design to generate soluble and functional membrane protein analogues [26]. In this approach, a robust computational pipeline is used for de novo protein design, using inversion of the AF2 network [27] coupled with sequence design using ProteinMPNN [28]. The soluble analogues could be designed in soluble form while preserving native functional motifs. In this approach, inversion of the AF2 network (AF2seq) is used to generate sequences that adopt a desired target fold and then apply ProteinMPNN sequence optimization [28]. The structures of the resulting sequences are re-predicted with AF2 and filtered based on their structural similarity to the target topology.
Troubleshooting
Problem 1: Lack of expression of fusion proteins or expression of more truncated protein.
Possible cause: The expression of fusion proteins is influenced by several critical factors, including the quality and concentration of the plasmid DNA, the choice of in vitro expression kit, and the specific expression conditions employed. Furthermore, degradation of proteins during IVTT reactions can lead to the truncation of proteins, affecting its functionality and downstream applications.
Solution: Ensure high-quality plasmid DNA extraction by consistently utilizing an appropriate plasmid extraction kit to obtain purified DNA. Maintain the concentration of plasmid DNA within the range of 500–1,000 ng per reaction during protein expression. Employ only sterilized materials, such as pipette tips, tubes, and water, to minimize contamination risks during protein synthesis. Experiment with multiple in vitro expression kits initially to identify the most suitable kit for achieving minimal or no truncation of the protein of interest. It is advisable to assess the protein synthesis profile using western blotting when employing a new in vitro expression kit to verify the integrity and quality of the synthesized proteins.
Problem 2: Lack of binding of fusion protein to the well.
Possible cause: The binding of the fusion protein is influenced by several key factors, including the concentration of the capture antibody (CAb), the presence of truncated protein variants, and the duration of the binding process. Additionally, other factors such as the affinity of the CAb for the tag of fusion protein, the specificity of the interaction, and the purity of the CAb can also impact the binding efficiency.
Solution: It is essential to optimize the dilution of the CAb in the well to ensure the capture of the fusion proteins at the appropriate amounts. Excessive synthesis of truncated proteins can hinder the binding of functional proteins in the well. The ideal duration for effective binding of the fusion protein in the well typically falls within the range of 4–12 h. Additionally, using a new batch of CAb can enhance the binding capacity, resulting in increased capture of fusion proteins in the well.
Problem 3: Production of non-functional protein.
Possible cause: The expression of fusion proteins using an in vitro expression kit may yield a large quantity of protein, but it can often be in an inactive form. This can occur due to various factors, including improper protein folding, lack of post-translational modifications, or the absence of necessary co-factors. Additionally, issues such as incorrect buffer composition, inadequate incubation conditions, or suboptimal reaction pH can also contribute to protein inactivity.
Solution: Experiment with various in vitro expression kits to evaluate their ability to facilitate proper folding of the synthesized fusion protein. Select a kit that contains all the essential components necessary for correct protein folding, including chaperone proteins or other factors crucial for post-translational modifications. Furthermore, optimize enzyme activity by testing different co-factors that may enhance protein functionality. Consider factors such as temperature, pH, and incubation time during optimization to ensure maximal enzyme activity.
Problem 4: Lack of binding of plasmid DNA to the wells.
Possible cause: The binding of plasmid DNA in wells is influenced by several factors, including the biotinylation of the plasmid DNA, the quantity and quality of streptavidin present, and the amount of biotinylated plasmid DNA applied to the well.
Solution: The use of new batches of chemicals can enhance the efficiency of biotinylation of plasmid DNA. It is essential to optimize the amount of biotinylation of DNA to achieve maximum protein synthesis in the wells. Additionally, factors such as reaction time, temperature, and the ratio of biotin to DNA should be carefully considered and optimized to ensure optimal biotinylation efficiency.
Problem 5: Lack of enzyme activity detection using the GLO assay.
Possible cause: Enzyme activity is determined by various factors such as no or less capture antibody, high concentration of CAb, the loss of enzyme activity, or not suitable buffer system; a high amount of captured enzyme on the wells can lower accessibility of donor and acceptor to the enzyme. Therefore, it is critical to determine the optimal amount to attach to the wells. The lack of transferase activity can be due to the lack of metal ions in the buffer system. The production of truncated protein that binds to the capture antibody can prevent the capture of functional protein.
Solution: Include a positive control to make sure that CAb is attached to the microplates, use Halo-GST fusion protein (commercially available) to determine the capturing capacity of the CAb, and use AccuBlue Next Gen dsDNA quantitation kit (Biotium) to determine the amounts of plasmid DNA attached to microplates. For enzyme assays, potential issues can be identified through meticulous adjustments to antibody concentration, buffer composition, enzyme concentration, and metal ion presence to maximize enzyme activity detection.
Problem 6: Detection of enzyme activity using GLO assay, but inability to detect the oligosaccharide products.
Possible cause: Some GTs may have hydrolytic activity that effectively releases sugar from NDP-sugar substrates without sugar transfer to the acceptor. Low binding of GTs on the microplates (see Problem 1–5).
Solution: Consider including a control without the acceptor, stabilizing NDP-sugars with metal ions (Mn and/or Mg). If the amount of the products formed is low, increase the number of reactions that are subsequently combined, which can significantly enhance detection sensitivity and accuracy. If the amount of GT captured on the microplate surface is low, the potential reasons could include the lack of expression of fusion proteins, the expression of more truncated protein, the lack of binding of fusion protein to the well, or the lack of binding of plasmid DNA to the wells; see troubleshooting 1, 2, and 4 to solve the problem.
Problem 7: Detecting the PPI in negative controls.
Possible cause: Detection of PPI in negative controls may arise from inadequate washing of the secondary antibody and insufficient blocking of the wells to prevent non-specific binding.
Solution: Thorough washing to ensure that the secondary antibody is removed by incorporating 0.05% Tween 20 in the washing buffer and subsequent removal of Tween 20 residues through additional washing with buffer alone. Enhance blocking efficiency by incubating the wells with 5% fat-free milk for an extended period, such as overnight at 4 °C.
Acknowledgments
This work was supported by an USDA-NIFA award (#1019179). This protocol is adapted from Bhattarai et al. [14] and Javaid et al. [15] and has also been used in Bhattarai et al. [13].
Competing interests
The authors declare no competing interests.
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Javaid, T., Bhattarai, M., Venkataraghavan, A., Held, M. and Faik, A. (2024). Specific protein interactions between rice members of the GT43 and GT47 families form various central cores of putative xylan synthase complexes. Plant J. 118(3): 856–878.
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Article Information
Publication history
Received: May 4, 2024
Accepted: Aug 1, 2024
Available online: Aug 16, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Expansion Microscopy of Synaptic Contacts on the Mauthner Cells of Larval Zebrafish
SI Sundas Ijaz *
SC Sandra P. Cárdenas-García *
AP Alberto E. Pereda
(*contributed equally to this work)
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5067 Views: 431
Reviewed by: Munenori IshibashiSébastien GillotinDhruv Rajanikant Patel
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Original Research Article:
The authors used this protocol in eLIFE Jul 2024
Abstract
Because of its genetic tractability and amenability for live imaging, larval zebrafish (Danio rerio) have emerged as a model to study the cellular and synaptic properties underlying behavior. The accessibility of Mauthner cells, a pair of escape-organizing neurons located in the brainstem of teleost fish, along with their associated sensory inputs, enables exploration of the correlation between structural and functional synaptic features. This is the case of the endings of auditory afferents on the lateral dendrite of this cell, known as large myelinated club endings, which provide the excitatory drive for the initiation of auditory-evoked escape responses mediated by the Mauthner cell and its spinal network. Here, we describe the procedures that make it possible to expose the molecular composition of these synapses using protein-retention expansion microscopy (proExM). This method allowed us to generate a map of the distribution of synaptic proteins at these identifiable synapses, which could also be applied to examine the organization of other synaptic contacts in this cell.
Key features
• This protocol builds upon the method developed by Tillberg et al. [1]
• Optimized for the examination of the organization of molecular components at synaptic contacts on the Mauthner cells of larval zebrafish
• Requires at least three days to complete and should be preceded by immunostaining.
• Results in a linear expansion factor of ~3.9× and an area expansion factor of ~13×
Keywords: Auditory Escape Immunohistochemistry Nanoscale imaging Connexin Electrical synapse
Background
The Mauthner cells are a pair of large reticulospinal neurons involved in tail-flip escape responses in fish [2,3]. Because of their experimental accessibility, these cells are considered a valuable model for studying vertebrate synaptic transmission, as they more easily allow for the correlation between the structure and function of synaptic features [4–7]. Auditory afferents originate in the sacculus, an organ of the vestibular system with auditory function in fish, and terminate as single large myelinated club endings [8,9] or club endings (CEs) on the lateral dendrite of the Mauthner cells. These terminals support both electrical and chemical synaptic transmission [5,10]. Because of their anatomical and physiological identifiability, these terminals have historically been amenable to exploring synaptic structure and function with novel technical approaches. For example, the seminal work by J.D. Robertson [4], using an electron microscope at these contacts, provided early evidence for the anatomical bases for electrical transmission at the gap junction. We have recently applied protein-retention expansion microscopy (proExM) to study the overall organization of these terminals. Unlike the more labor-intensive electron microscopy, this technique allowed us to generate a map of the incidence and distribution of synaptic proteins associated with electrical and glutamatergic transmission [11]. ProExM was able to expose, with sufficient resolution, spatial features that have only been observed so far with electron microscopy. Moreover, unlike electron microscopy, expansion microscopy allowed us to more easily generate a map of the organization of the synaptic contact by labeling proteins that form its various synaptic components. The protocol that we describe here is based on a previously established methodology [1]; we propose that it would be useful for exploring the anatomical organization of other synapses on the distal portion of the Mauthner cell lateral dendrite. This includes inhibitory small vesicle boutons (SVBs) [12], known to surround CEs, as well as synapses located in other processes of this large cell and throughout the zebrafish brain in general.
Materials and reagents
Reagents
10× phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: 6506-OP)
Dimethyl sulfoxide (DMSO) (Honeywell, catalog number: 67-68-5)
Cell culture–grade water (Sigma-Aldrich, catalog number: W3500)
Trichloroacetic acid (TCA) (Sigma-Aldrich, catalog number: T6399)
Normal goat serum (NGS) (Vector Laboratories, catalog number: S-1000)
Acryloyl-X, SE (AcX) (Invitrogen, catalog number: A20770)
Primary antibodies used for immunostaining:
Mouse IgG1 anti-Cx35/36 (monoclonal) (Millipore Sigma, catalog number: MAB3045, 1:250 dilution)
Rabbit anti-Cx35.5 (monoclonal) (Fred Hutch Antibody Technology Facility: clone12H5, 1:200 dilution)
Mouse IgG2A anti-Cx34.1 (monoclonal) (Fred Hutch Antibody Technology Facility: clone5C10A, 1:200 dilution)
Mouse IgG1 anti-ZO1 (monoclonal) (Invitrogen, catalog number: 33-9100, 1:200 dilution)
Mouse IgG1 anti-N-cadherin (monoclonal) (BD Transduction Laboratories, catalog number: 610920, 1:50 dilution)
Mouse IgG1 anti-Beta-catenin (monoclonal) (Sigma, catalog number: C7207, 1:100 dilution)
Chicken IgY anti-GFP (polyclonal) (Abcam, catalog number: ab13970, 1:200 dilution)
Rabbit IgG anti-GluR2/3 (polyclonal) (EMD Millipore, catalog number: 07-598, 1:200 dilution)
Secondary antibodies used for immunostaining:
Mouse IgG Alexa Fluor 546 (polyclonal) (Invitrogen, catalog number: A11030, 1:200 dilution)
Mouse IgG Atto 647N (polyclonal) (Sigma-Aldrich, catalog number: 50185, 1:200 dilution)
Rabbit IgG Alexa fluor 546 (polyclonal) (Invitrogen, catalog number: A11010, 1:200 dilution)
Rabbit IgG Atto 647N (polyclonal) (Sigma-Aldrich, catalog number: 40839, 1:200 dilution)
Chicken IgY Alexa Fluor 488 (polyclonal) (Invitrogen, catalog number: A11039, 1:200 dilution)
Sodium acrylate (Sigma-Aldrich, catalog number: 408220)
Acrylamide (Sigma-Aldrich, catalog number: A9099)
N,N'-methylenebisacrylamide (Sigma-Aldrich, catalog number: M7279)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
4-hydroxy-TEMPO (4-HT) (Sigma-Aldrich, catalog number: 176141)
Tetramethylethylenediamine (TEMED) (Sigma-Aldrich, catalog number: T7024)
Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: A3678)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
1 M tris(hydroxymethyl)aminomethane (Tris), pH 8.0 (Invitrogen, catalog number: AM9855G)
0.5 M Ethylenediaminetetraacetic acid (EDTA) (Invitrogen, catalog number: AM9260G)
Proteinase K, >600 U/mL (Thermo Fisher, catalog number: EO0491)
Poly-D-Lysine, 1 mg/mL (Sigma-Aldrich, catalog number: A-003-E)
Reagent alcohol (ethanol) (Sigma-Aldrich, catalog number: 362808)
Solutions
10% blocking solution (see Recipes)
2% trichloroacetic acid (TCA) solution (see Recipes)
1% acryloyl-X, SE (AcX) stock solution (see Recipes)
38% sodium acrylate stock solution (see Recipes)
50% acrylamide stock solution (see Recipes)
2% N,N'-methylenebisacrylamide stock solution (see Recipes)
5 M sodium chloride (NaCl) stock solution (see Recipes)
0.5% 4-hydroxy-TEMPO (4-HT) stock solution (see Recipes)
10% tetramethylethylenediamine (TEMED) stock solution (see Recipes)
10% ammonium persulfate (APS) stock solution (see Recipes)
Anchoring solution (see Recipes)
Monomer solution (see Recipes)
Gelling solution (see Recipes)
Digestion buffer (see Recipes)
Proteinase K solution (8 U/mL) (see Recipes)
70% ethanol (see Recipes)
Recipes
10% blocking solution
Note: Prepare this fresh with each experiment.
Reagent Final concentration Quantity or Volume
NGS 10% (v/v) 200 μL
DMSO 1% (v/v) 20 μL
0.5% Triton X-100 in PBS (PBS-Trx) n/a Fill to a final volume of 2 mL
2% trichloroacetic acid (TCA) solution
Note: Prepare this fresh with each experiment.
Reagent Final concentration Quantity or Volume
TCA 2% (w/v) 1 g
1× PBS n/a Fill to a final volume of 50 mL
1% acryloyl-X, SE (AcX) stock solution
Note: Prepare 20 μL aliquots of this and store at -20 °C with a drying agent for up to two months.
Reagent Final concentration Quantity or Volume
AcX 1% (w/v) 5 mg
DMSO n/a 500 μL
38% sodium acrylate stock solution
Note: Store this at 4 °C for up to one month.
Reagent Final concentration Quantity or Volume
Sodium acrylate 38% (w/v) 1.9 g
Cell culture–grade water n/a Fill to a final volume of 5 mL
50% acrylamide stock solution
Note: Store this at 4 °C for up to six months.
Reagent Final concentration Quantity or Volume
Acrylamide 50% (w/v) 5 g
Cell culture–grade water n/a Fill to a final volume of 10 mL
2% N,N'-methylenebisacrylamide stock solution
Note: Store this at 4 °C for up to six months.
Reagent Final concentration Quantity or Volume
N,N'-methylenebisacrylamide 2% (w/v) 0.2 g
Cell culture–grade water n/a Fill to a final volume of 10 mL
5 M Sodium chloride (NaCl) stock solution
Reagent Final concentration Quantity or Volume
NaCl 5 M 14.6 g
Cell culture–grade water n/a Fill to a final volume of 50 mL
0.5% 4-hydroxy-TEMPO (4-HT) stock solution
Note: Prepare 100 μL aliquots of this and store at -20 °C with a drying agent for up to one month.
Reagent Final concentration Quantity or Volume
4-HT 0.5% (w/v) 50 mg
Cell culture–grade water n/a Fill to a final volume of 10 mL
10% Tetramethylethylenediamine (TEMED) stock solution
Note: Prepare 100 μL aliquots of this and store at -20 °C with a drying agent for up to one month.
Reagent Final concentration Quantity or Volume
TEMED 10% (v/v) 20 μL
Cell culture–grade water n/a 180 μL
Total n/a 200 μL
10% ammonium persulfate (APS) stock solution
Note: Prepare 100 μL aliquots of this and store at -20 °C with a drying agent for up to one month.
Reagent Final concentration Quantity or Volume
APS 10% (w/v) 1 g
Cell culture–grade water n/a Fill to a final volume of 10 mL
Anchoring solution
Reagent Final concentration Quantity or Volume
1% AcX stock solution 0.01% (v/v) 2 μL (dilute 1:100)
1× PBS n/a 198 μL
Total n/a 200 μL
Monomer solution
Note: The final concentration of these reagents will decrease slightly in the gelling solution with the addition of 4-HT, TEMED, and APS. These concentrations are listed in parenthesis. Additionally, this solution should be made fresh with every experiment.
Reagent Final concentration Quantity or Volume
38% sodium acrylate stock solution 9.1% (8.6%) 2.25 mL
50% acrylamide stock solution 2.7% (2.5%) 0.5 mL
2% N,N'-methylenebisacrylamide stock solution 0.16% (0.15%) 0.75 mL
5 M NaCl stock solution 2.13 M (2 M) 4 mL
10× PBS 1.06× (1×) 1 mL
Cell culture–grade water n/a 0.9 mL
Total n/a 9.4 mL
Gelling solution
Note: 4-HT, TEMED, and APS should be added sequentially, in the order they are listed, per individual sample. This solution should not be mixed beforehand.
Reagent Final concentration Quantity or Volume
Monomer solution n/a 376 μL
0.5% 4-HT stock solution 0.01% 8 μL
10% TEMED stock solution 0.2% 8 μL
10% APS stock solution 0.2% 8 μL
Total n/a 400 μL
Digestion buffer
Note: Prepare 5 mL aliquots of this and store at -20 °C for up to one year.
Reagent Final concentration Quantity or Volume
1 M Tris (pH 8.0) 50 mM 2.5 mL
0.5 M EDTA 1 mM 0.1 mL
Triton X-100 0.5% (v/v) 0.25 mL
5 M NaCl stock solution 0.5 M 5 mL
Cell culture–grade water n/a 42.15 mL
Total n/a 50 mL
Proteinase K solution (8 U/mL)
Note: Proteinase K should only be added prior to the digestion step. Do not freeze with digestion buffer for storage.
Reagent Final concentration Quantity or Volume
Proteinase K, >600 U/mL 8 U/mL 3.3 μL
Digestion buffer n/a 246.7 μL
Total n/a 250 μL
70% ethanol
Reagent Final concentration Quantity or Volume
Ethanol 70% 35 mL
Cell culture–grade water n/a 15 mL
Total n/a 50 mL
Laboratory supplies
Fine Science Tools Dumont #4 forceps 0.13 × 0.08 mm, 11 cm (Fisher Scientific, catalog number: NC9091939)
Fisherbrand Premium plain microscope slides, 25 × 75 × 1.0 mm (Fisher Scientific, catalog number: 125444)
Epredia cover slips, 22 × 22, No.1 (Fisher Scientific, catalog number: 102222)
Fisherbrand cover slips, 24 × 50, No.1 (Fisher Scientific, catalog number: 12-545-88)
Falcon 24-well tissue culture plate (Fisher Scientific, catalog number: 353047)
Greiner 6-well cell culture plate (Fisher Scientific, catalog number: 657160)
Heathrow Scientific slide mailer (Sigma-Aldrich, catalog number: HS120557)
Fisherbrand glass Pasteur pipette (Fisher Scientific, catalog number: 13-678-20A)
USA Scientific SealRite 0.5 mL microcentrifuge tubes (USA Scientific, catalog number: 1605-0000)
USA Scientific SealRite 1.5 mL microcentrifuge tubes (USA Scientific, catalog number: 1615-5510)
Ted Pella sable brush #0, 1.3 mm W × 8.0 mm L (Ted Pella, catalog number: 11810)
Ted Pella sable brush #1, 1.5 mm W × 9.5 mm L (Ted Pella, catalog number: 11812)
Equipment
Titer plate shaker model 4625 (Lab-line Instruments, catalog number: 0101-1383)
Heratherm incubator (Fisher Scientific, catalog number: 50125590)
Fisherbrand water bath (Fisher Scientific, catalog number: 15-462-2Q)
Stereo microscope (Leica, model: MZFLIII)
Confocal microscope (Zeiss, model: LSM 710)
Software and datasets
Zen v2.3 black edition (Zeiss), https://www.micro-shop.zeiss.com/en/us/softwarefinder/software-categories/zen-black/
Prism 10 for Windows 64-bit (GraphPad, version 10.2.3 9403, April 21, 2024)
Fiji ImageJ, open source, https://www.nature.com/articles/nmeth.2019)
Adobe Photoshop 2023 Adobe Photoshop Version: 24.7.0 20230719.r.643 efe3886 ×64, September 2023)
Procedure
Prior to expansion procedure: fixation, dissection, and immunostaining
Fixation
Prior to the expansion procedure, fix 5-days-post-fertilization (dpf) larvae in 2% TCA solution for 3 h at room temperature (RT) on a rocker.
Remove the 2% TCA solution and wash larvae with 1× PBS three times.
Dissection
Pin larvae down to a Sylgard-coated dish and remove the eyes and skin on top of the head of the larvae with forceps.
Similarly, remove the notochord with forceps.
Remove the head of the larvae by making a cut at the level of the yolk sac.
Place the dissected brain into a 0.5 mL microcentrifuge tube with 350 μL of blocking solution (for more details, see Figure 1).
Immunostaining
Incubate dissected heads in the blocking solution for 3 h at RT on a rocker.
Remove the blocking solution, add appropriately diluted primary antibody in 300 μL of blocking solution, and incubate the samples overnight at RT on a rocker.
Remove the primary antibody solution and rinse with 0.5% PBS-Trx three times.
Wash samples in 0.5% PBS-Trx for 10 min, 15 min, and 30 min with fresh solution changes each time, on a rocker.
Remove 0.5% PBS-Trx and add appropriately diluted secondary antibody in 300 μL of blocking solution. Incubate the samples for 4 h at RT in the dark on a rocker.
Remove the secondary antibody solution and rinse with 1× PBS three times.
Wash samples in 1× PBS four times for 15 min on a rocker.
Proceed with the following protocol (expansion steps) on the same day.
Note: All incubations and washes take place on a rocker.
Note: Some fluorescent dyes conjugated to antibodies are not compatible with the expansion protocol and will degrade, especially those in the cyanine family (Cy3, Cy5, and Alexa 647) [1,13]. Recommended secondary antibodies include CF 405M, Alexa 488, Alexa 546, Alexa 594, CF 633, and Atto 647N. Please refer to the Reagents section for a list of fluorescent antibodies that have been successfully used in this protocol [1].
Figure 1. Brain dissection of larval zebrafish
Day 1: anchoring
Note: Stained samples should be protected from light as much as possible throughout the remainder of the procedure to prevent bleaching of the fluorophores.
Following staining, carefully slide the tips of the forceps beneath the larva’s skin and remove the skin from the brain.
Individually place the brains into a 24-well plate using a glass pipette and add 200 μL of anchoring solution to each well.
Place the plate on a shaker overnight (for at least 16 h) at RT to enhance the diffusion of the anchoring solution through the sample.
Note: It is crucial to set the shaker at an appropriate speed (40–70 rpm) to prevent the brains from sticking to the walls of the well, as this may lead to them drying out and/or fail to allow the anchoring solution to penetrate the samples deep enough for the expansion process to work effectively.
Day 2: gel polymerization
After the overnight incubation, remove the anchoring solution from each well and wash samples with 200 μL of 1× PBS. This step should be performed on a shaker (at speed 40–70 rpm) for 10 min at RT and repeated twice.
While the samples are washing, construct custom-made coverslip wells for use during gel polymerization: Use a microscope slide as a base, place two coverslips (22 × 22), one on either end of the slide, and add ~2 μL of water to adhere the coverslips to the slide. Apply a second set of coverslips over the first set using super glue. Make enough coverslip wells for every sample. (For more details, see Figure 2A. For a video demonstration, see Tillberg et al. [1].)
Note: Two sets of coverslips are used to accommodate for the thickness of the sample (~300 μm). Use as many coverslips as needed to avoid crushing the sample.
After washing with 1× PBS, remove the 1× PBS from each well and replace it with 376 μL of monomer solution, and then sequentially add 8 μL of 0.5% 4-HT stock solution.
Rock the plate on a shaker (at speed 40–70 rpm) for 10 min at RT, allowing the 376 μL of monomer solution to mix with the 8 μL of 0.5% 4-HT stock solution and to allow the mixture to diffuse into the sample.
Remove the solution from step C4 and add 376 μL of fresh monomer solution. Then, add 8 μL of 0.5% 4-HT stock solution.
This time, rock the plate on a shaker (at speed 40–70 rpm) for 5 min at RT, again allowing the 376 μL of monomer solution to mix with the 8 μL of 0.5% 4-HT stock solution and to allow the mixture to diffuse into the sample.
Note: Steps C7–9 should be performed on one sample at a time and executed quickly to prevent premature polymerization of samples. Once step C9 is performed on one sample, go back to step C7 and repeat with each sample.
Add 8 μL of 10% TEMED stock solution and rock the plate on a shaker (at speed 40–70 rpm) for 1 min.
Add 8 μL of 10% APS stock solution and rock the plate on a shaker (at speed 40–70 rpm) for 30 s. Together, these reagents form the gelling solution and initiate polymerization.
Before the polymerization is completed, transfer the samples using a glass Pasteur pipette with a small amount of gelling solution (~100 μL) to the custom-made coverslip wells. Mount the sample either dorsal or ventral side up, ensuring that there are no air bubbles, and place a coverslip over the sample (24 × 50) (for more details, see Figure 2B).
Note: Make sure that enough solution is added along with the sample to the coverslip well. Too much could disturb the structure of the well, and too little could dry out the sample.
Figure 2. Sample mounting for gel polymerization
Place the samples into a slide mailer and incubate them at 4 °C for 50 min. This will help to reduce the polymerization rate of the gel and ensure proper diffusion of the solution into the brain [15,16].
Note: Handle the samples with care while placing them in a slide mailer, taking care not to move the coverslip on top of the sample, as this will shift the mounting orientation.
Move the samples into a 37 °C incubator for 2 h, resulting in gel formation in and around the brain [15,16].
While the samples undergo incubation, add 250 μL of proteinase K solution (8 U/mL) to 1.5 mL microcentrifuge tubes (one tube per sample).
Once the incubation period is complete, remove the coverslips (both 22 × 22 and 24 × 50) from the mounted sample under a dissecting microscope.
Moisten the top of the sample (now in the hydrogel) with the digestion buffer using a paintbrush to ensure that it remains hydrated, and carefully cut the gel surrounding the sample using a scalpel or razor blade (for more details, see Figure 3).
Note: Introduce an asymmetrical cut to the gel to establish the orientation of the sample (ventral or dorsal) for reference later. Due to TCA fixation, pigment from the skin over the head remains on the dorsal side of the brain, making it easy to identify the orientation.
Figure 3. Identify the orientation of the sample (dorsal side here) embedded in the hydrogel
Using a paint brush, transfer the samples into the microcentrifuge tubes containing the proteinase K solution (8 U/mL).
Incubate the samples overnight for 12 h in a water bath at 50 °C. This step will allow proteinase K to digest the brain tissue, resulting in a translucent sample.
Day 3: expansion
Following the overnight incubation, transfer the samples into a 6-well plate using a paintbrush, with one sample per well.
Remove any remaining digestion buffer.
Wash the samples with 5 mL of cell culture–grade water for 30 min on a shaker. To induce gel expansion, perform four additional washes, each one for 30 min to allow the tissue to gradually expand.
Note: If imaging samples on a different day, do not wash with water; instead, store samples in 5 mL of 1× PBS at 4 °C in the dark, until you are ready to image.
While the gel is expanding, prepare the microscope slides for imaging (one slide per sample): start by cleaning the slides with 50 μL of water in a laminar fume hood and dry with Kimwipes. Repeat this twice. Next, clean slides with 50 μL of 70% ethanol and dry with Kimwipes. After the ethanol has evaporated, apply 50 μL of Poly-D-Lysine (1 mg/mL) to the center of the slide and incubate at RT, in the fume hood, for 30 min. Subsequently, remove any excess Poly-D-Lysine and perform three washes with 50 μL of cell culture–grade water and let slides dry before mounting sample (for more details, see Figure 4).
Note: It is essential to mark the reference point of Poly-D-Lysine placement on the slide, as that will serve as the designated spot for mounting the sample.
After the washes are done and the expansion process is complete, carefully remove all the water from the well (for more details, see Figure 5A).
Note: Only work with one sample at a time. Leave water in the wells of the samples you are not actively working with to ensure that they remain hydrated.
Dry the bottom of the well with a Kimwipe carefully to avoid touching the gel.
Place a coverslip in the well and gently slide the sample over the coverslip using a paintbrush. Ensure that the orientation for mounting is ventral side up in an upright confocal microscope. This ensures the Mauthner cell is easily accessible, as it is located on the ventral side of the brainstem (for more details, see Figure 5B)
Note: This method ensures that the sample is not directly manipulated, thereby reducing the risk of breaking the gel.
Using forceps, carefully remove the coverslip with the gel and slide the gel onto the microscope slide with Poly-D-Lysine using a paint brush (for more details, see Figure 4 and 5C).
Figure 4. Mount the gel and image
To locate the brain, specifically the Mauthner cell within the gel, use a dissecting scope and place a small piece of Kimwipe (~3 mm × 3 mm) on top of the gel, around the hindbrain region, to serve as a reference point for the general vicinity of the Mauthner cell.
Note: Although the tissue has been digested, it is still possible to determine the original location and orientation of the brain due to the presence of residual pigment on the dorsal side of the brain, an effect of TCA fixation (for more details, see Figure 3B).
To find the sample using an upright confocal microscope, use a 10× objective with the transmitted light on and identify the position of the Kimwipe.
Carefully remove the Kimwipe with forceps and add ~1 mL of cell culture–grade water to maintain gel hydration and assist in visualizing the sample using a 40× water immersion objective.
Locate the lateral dendrite of the Mauthner cell and begin imaging (for more details, see Figure 4C).
Figure 5. Remove sample from the 6 well-plate
Data analysis
Images were acquired using the Zeiss LSM 710 confocal microscope with a 40× (1.0 NA) water immersion objective. The acquired images of expanded samples, when compared to non-expanded samples, retained normal anatomical features (Figure 6, 10). Data analysis methods, which have been described in Cárdenas-García et al., 2024 (Results and Methods, 10), were used to determine the achieved expansion factor and colocalization, distribution, and occupancy of synaptic components at CEs. To give a brief overview of these methods, labeling of the oval CE contact areas on the Mauthner cells was measured pre- and post-expansion in Fiji. These measurements resulted in a ~3.9× lateral expansion factor and a ~13× area expansion factor (10). Colocalization of components was determined by analyzing fluorescence overlap and was performed in Fiji, with the JACoP plugin, followed by quantification using the Manders’ coefficient. Differential distribution of molecular components was also determined in Fiji by defining regions of interest (ROIs) in different areas of the contact and quantifying the fluorescence intensity in each ROI. Additionally, pre- vs. post-synaptic distribution of electrical components was more easily discernible given the increase in resolution in post-expanded samples and was found using line scan in Fiji. Similarly, line scan analysis was used to determine the distribution of fluorescence of any component relative to another. Finally, all this data was combined to determine the organization and proportion of these synaptic proteins at CE contacts (please see Cárdenas-García et al. [11] for more details). Finally, while this protocol proved to be very efficient, its main limitation is that it results in some loss of fluorescence intensity due to the necessary use of protease, which allows for isotropic expansion.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Cárdenas-García et al. (2024). The components of an electrical synapse as revealed by expansion microscopy of a single synaptic contact. eLife (Figure 1, panel E; Figure 2, panel A-C; Figure 3, panel A-D; Figure 4, panel C-E; Figure 5, panel A-C; Figure 6, panel C, E, G, H).
Acknowledgments
This protocol was used in the companion paper Cárdenas-García et al. (2024) eLife (https://doi.org/10.7554/eLife.91931) [11]. We thank Alyssa Brunal and Albert Pan for sharing their expansion protocol at the early stages of our experiments.
Supported by National Institute on Deafness and Other Communication Disorders (R01DC011099), National Institute of Neurological Disorders and Stroke (R21NS085772) and National Institute of Mental Health (RF1MH120016) grants to Alberto Pereda.
Competing interests
We declare no competing interests.
Ethical considerations
Studies using this protocol were performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to approved institutional animal care and use committee (IACUC) protocol of the Albert Einstein College of Medicine.
References
Tillberg, P. W., Chen, F., Piatkevich, K. D., Zhao, Y., Yu, C. C., English, B. P., Gao, L., Martorell, A., Suk, H. J., Yoshida, F., et al. (2016). Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies. Nat Biotechnol. 34(9): 987–992. https://doi.org/10.1038/nbt.3625
Pereda, A. and Faber, D. (2011). BRAIN AND NERVOUS SYSTEM | Physiology of the Mauthner Cell: Discovery and Properties. Encycl Fish Physiol. 66–72. https://doi.org/10.1016/b978-0-12-374553-8.00007-1
Faber, D. and Pereda, A. (2011). BRAIN AND NERVOUS SYSTEM | Physiology of the Mauthner Cell: Function. Encycl Fish Physiol. 73–79. https://doi.org/10.1016/b978-0-12-374553-8.00280-x
Robertson, J. D., Bodenheimer, T. S. and Stage, D. E. (1963). The ultrastructure of Mauthner cell synapses and nodes in goldfish brains. J Cell Biol. 19(1): 159–199. https://doi.org/10.1083/jcb.19.1.159
Furshpan, E. J. (1964). "Electrical Transmission" at an Excitatory Synapse in a Vertebrate Brain. Science. (1979). 144(3620): 878–880. https://doi.org/10.1126/science.144.3620.878
Flores, C. E., Nannapaneni, S., Davidson, K. G. V., Yasumura, T., Bennett, M. V. L., Rash, J. E. and Pereda, A. E. (2012). Trafficking of gap junction channels at a vertebrate electrical synapse in vivo. Proc Natl Acad Sci USA. 109(9): e1121557109. https://doi.org/10.1073/pnas.1121557109
Pereda, A. E., Curti, S., Hoge, G., Cachope, R., Flores, C. E. and Rash, J. E. (2014). Corrigendum to “Gap junction-mediated electrical transmission: Regulatory mechanisms and plasticity” [Biochim. Biophys. Acta 1828 (2013) 134–146]. Biochimica et Biophysica Acta (BBA) - Biomembranes 1838(3): 1056. https://doi.org/10.1016/j.bbamem.2013.10.013
Bartelmez, G. W. (1915). Mauthner's cell and the nucleus motorius tegmenti. J Comp Neurol. 25(1): 87–128. https://doi.org/10.1002/cne.900250105
Bartelmez, G. W. and Hoerr, N. L. (1933). The vestibular club endings in ameiurus. Further evidence on the morphology of the synapse. J Comp Neurol. 57(3): 401–428. https://doi.org/10.1002/cne.900570303
Lin, J. and Faber, D. (1988). Synaptic transmission mediated by single club endings on the goldfish Mauthner cell. II. Plasticity of excitatory postsynaptic potentials. J Neurosci. 8(4): 1313–1325. https://doi.org/10.1523/jneurosci.08-04-01313.1988
Cárdenas-García, S. P., Ijaz, S. and Pereda, A. E. (2024). The components of an electrical synapse as revealed by expansion microscopy of a single synaptic contact. eLife. https://doi.org/10.7554/eLife.91931
Tuttle, R., Masuko, S. and Nakajima, Y. (1987). Small vesicle bouton synapses on the distal half of the lateral dendrite of the goldfish mauthner cell: Freeze‐fracture and thin section study. J Comp Neurol. 265(2): 254–274. https://doi.org/10.1002/cne.902650209
Gao, R., Asano, S. M. and Boyden, E. S. (2017). Q&A: Expansion microscopy. BMC Biol. 15(1): 50. https://doi.org/10.1186/s12915-017-0393-3
Freifeld, L., Odstrcil, I., Förster, D., Ramirez, A., Gagnon, J. A., Randlett, O., Costa, E. K., Asano, S., Celiker, O. T., Gao, R., et al. (2017). Expansion microscopy of zebrafish for neuroscience and developmental biology studies. Proc Natl Acad Sci USA. 114(50): e1706281114. https://doi.org/10.1073/pnas.1706281114
Wassie, A. T., Zhao, Y. and Boyden, E. S. (2018). Expansion microscopy: principles and uses in biological research. Nat Methods. 16(1): 33–41. https://doi.org/10.1038/s41592-018-0219-4
Chen, F., Tillberg, P. W., Boyden, E. S. (2015). Optical imaging. Expansion microscopy. Science. 347(6221): 543–548. https://doi.org/10.1126/science.1260088
Article Information
Publication history
Received: Mar 25, 2024
Accepted: Jul 28, 2024
Available online: Aug 12, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
Neuroscience > Cellular mechanisms > Synaptic physiology
Cell Biology > Cell structure > Plasma membrane
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5,068 | https://bio-protocol.org/en/bpdetail?id=5068&type=1 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
GATK Variant Discovery Pipeline
CH Cheng He
SL Sanzhen Liu
Published: Sep 20, 2024
DOI: 10.21769/BioProtoc.5068 Views: 29
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Abstract
Phenotypic variations of most biological traits are largely driven by genomic variants. The single nucleotide polymorphism (SNP) is the most common form of genomic variants. Multiple algorithms have been developed for discovering genomic variants, including SNPs, with next-generation sequencing (NGS) data. Here, we present a widely used variant discovery pipeline based on the software Genome Analysis ToolKits (GATK). The pipeline uses whole-genome sequencing (WGS) data as input and includes read mapping, variant calling, and the variant filtering process. This pipeline has been successfully applied to many genomic projects and represents a solution for variant calling using NGS data.
Keywords: Variant discovery Single nucleotide polymorphism Next-generation sequencing Whole-genome sequencing GATK
Graphical overview
Workflow of the single nucleotide polymorphism (SNP) calling pipeline using GATK software
Background
As the technology allows regularly producing sequencing data in a high-throughput manner, genome-wide variant discovery becomes routine for many applications, including genotyping of variants for their phenotypic connection. Single nucleotide polymorphisms (SNPs) and small insertion–deletions (indels) are two common forms of genomic variants. The SNP, as the simplest molecular marker, has been widely used to connect genetic variation with phenotypic variation of biological traits, such as human diseases and plant architectures [1,2]. The reference-based SNP calling is the process by which DNA polymorphisms between sequencing reads and the reference are identified based on their alignments [3]. Algorithms and software packages have been developed for SNP calling. Simple SNP calling methods determine read counts at each polymorphic site and set a cutoff of read counts for SNP identification [4]. However, the power of SNP discovery using such methods is subject to sequencing depths. More sophisticated approaches incorporate the uncertainty of SNP calling in a probabilistic framework, such as SOAP2, SAMtools, or Genome Analysis ToolKits (GATK) [5–7]. Among them, GATK is the industry standard for discovering SNPs and small indels using sequencing data from genomic DNA and RNA-seq [7]. Although GATK was originally developed for human sequencing data, it has evolved to handle genome data from other organisms with different levels of ploidy [8]. Here, we present a variant calling pipeline using whole-genome sequencing (WGS) data based on GATK. The pipeline includes reads mapping, variant calling, and variant filtering.
Software and datasets
Software installation and data availability: All software installation instructions and test data have been deposited to GitHub: https://github.com/Bio-protocol/GATK-SNP-Calling
Software
All software packages have been tested on a Linux (CentOS 7, x86_64) operating system.
Anaconda (version 2024.02-1) (https://www.anaconda.com/download)
Trimmomatic (version 0.39, Java 1.8.0+) (http://www.usadellab.org/cms/?page=trimmomatic)
BWA (version 0.7.17) (https://github.com/lh3/bwa)
SAMtools (version 1.9) (http://www.htslib.org/)
GATK4 (version 3.8, Java 1.8.0+) (https://gatk.broadinstitute.org/)
vcftools (version 0.1.16) (http://vcftools.sourceforge.net/)
Perl (version 5) (https://perl.org)
Python (version 3) (https://python.org)
Data sets
Sample data for GATK SNP Calling workflow include:
B73/A188.R1/2.fq.gz: Paired-end whole genome sequencing reads for two test samples.
B73Ref4.fa: The maize B73 version4 reference genome.
TruSeq3-PE.fa: The adaptor sequences for reads trimming.
Sample reads and adaptor sequences were included in the “input” folder. The B73 version4 reference genome can be downloaded from MaizeGDB (https://download.maizegdb.org/Zm-B73-REFERENCE-GRAMENE-4.0/Zm-B73-REFERENCE-GRAMENE-4.0.fa.gz)
Procedure
You can download the GitHub repository with this command line:
git clone https://github.com/Bio-protocol/GATK-SNP-Calling.git
cd GATK-SNP-Calling
Step 0: Installation of required software
Note: Conda should be installed first and bioconda channel needs to be added into the conda configure with this command “conda config --add channels bioconda”.
Install the Anaconda and add it into ~/.bashrc.
wget https://repo.anaconda.com/archive/Anaconda3-2024.02-1-Linux-x86_64.sh
bash Anaconda3-2024.02-1-Linux-x86_64.sh
echo 'export PATH="~/anaconda3/bin:$PATH"' >> ~/.bashrc
source ~/.bashrc
All dependencies can be installed through the GATK_SNP.yaml file in the GitHub repository.
conda env create -f GATK_SNP.yaml
conda activate GATK_SNP
Alternatively, the conda environment can be created by following scripts if GATK_SNP.yaml is not used.
conda create -n GATK_SNP python=3
conda activate GATK_SNP
conda install -c bioconda trimmomatic
conda install -c bioconda bwa
conda install -c bioconda samtools
conda install -c bioconda gatk4
conda install -c bioconda vcftools
Step 1: Raw reads trimming
The Trimmomatic software is a flexible read trimming tool for Illumina NGS data [9]. It is widely used to remove low-quality reads and sequence adapters. As test inputs are paired-end reads, we use the paired-end mode for sequence trimming.
The code for raw reads trimming is as follows:
#!/bin/bash
## B73 reads trimming ##
trimmomatic PE \
./input/B73.R1.fq.gz ./input/B73.R2.fq.gz \
./cache/B73_trim.R1.fq.gz ./cache/B73_unpaired.R1.fq.gz \
./cache/B73_trim.R2.fq.gz ./cache/B73_unpaired.R2.fq.gz \
ILLUMINACLIP:./input/TruSeq3-PE.fa:3:20:10:1:true LEADING:3 TRAILING:3 SLIDINGWINDOW:4:13 MINLEN:40 2> ./cache/B73_trim.log
## A188 reads trimming ##
trimmomatic PE \
./input/A188.R1.fq.gz ./input/A188.R2.fq.gz \
./cache/A188_trim.R1.fq.gz ./cache/A188_unpaired.R1.fq.gz \
./cache/A188_trim.R2.fq.gz ./cache/A188_unpaired.R2.fq.gz \
ILLUMINACLIP:./input/TruSeq3-PE.fa:3:20:10:1:true LEADING:3 TRAILING:3 SLIDINGWINDOW:4:13 MINLEN:40 2> ./cache/A188_trim.log
For one-step running, you can simply run this script:
sh ./workflow/1_reads_trim.sh
Note:
(1) The Trimmomatic software should be run on a 1.8.0+ Java environment.
(2) If you want to trim data with your own adapter sequences, you should replace the adapter sequences in TruSeq3-PE.fa file with your own adapter sequences.
Step 2: BWA read alignment
BWA is a software package for mapping low-divergent sequences against a reference genome [10]. Here, we use BWA to map the remaining trimmed reads to the B73v4 reference genome.
BWA index for the reference genome
cd input
wget https://download.maizegdb.org/Zm-B73-REFERENCE-GRAMENE-4.0/Zm-B73-REFERENCE-GRAMENE-4.0.fa.gz
gunzip Zm-B73-REFERENCE-GRAMENE-4.0.fa.gz
mv Zm-B73-REFERENCE-GRAMENE-4.0.fa B73Ref4.fa
bwa index B73Ref4.fa
cd ../
BWA alignment
## BWA alignment ##
bwadb=./input/B73Ref4.fa
B73_R1=./cache/B73_trim.R1.fq.gz
B73_R2=./cache/B73_trim.R2.fq.gz
A188_R1=./cache/A188_trim.R1.fq.gz
A188_R2=./cache/A188_trim.R2.fq.gz
bwa mem -t 8 -R '@RG\tID:B73\tSM:B73' $bwadb $B73_R1 $B73_R2 > ./cache/B73.sam
bwa mem -t 8 -R '@RG\tID:A188\tSM:A188' $bwadb $A188_R1 $A188_R2 > ./cache/A188.sam
Adjustable parameters for BWA:
-t INT: Number of threads
-R STR: Complete read group header line. “\t” can be used in STR and will be converted to a TAB in the output SAM. The read group ID will be attached to every read in the output. An example is “@RG\tID:foo\tSM:bar”.
Note: Most parameters of BWA were set as default here; you can find the information of all the parameters in the BWA manual reference (https://bio-bwa.sourceforge.net/bwa.shtml)
For one-step running of this step, you can simply run this script:
sh ./workflow/2_bwa_align.sh
Step 3: SAM filtering and compressing
In this step, raw alignments (SAM format) are filtered and sorted based on alignment coordinates on the reference genome.
SAM filtering
Here, we use a Perl script to filter BWA alignment results (SAM format).
## SAM filtering ##
perl ./lib/samparser.bwa.pl -i ./cache/B73.sam -e 100 -m 3 100 --tail 5 100 --gap 0 --insert 100 800 1>./cache/B73.parse.sam 2>./cache/B73.parse.log
perl ./lib/samparser.bwa.pl -i ./cache/A188.sam -e 100 -m 3 100 --tail 5 100 --gap 0 --insert 100 800 1>./cache/A188.parse.sam 2>./cache/A188.parse.log
Adjustable parameters for samparser.bwa.pl are:
--input|i: SAM file
--identical|e: minimum matched and identical base length, default = 30 bp
--mismatches|mm|m: two integers to specify the number of mismatches out of the number of basepairs of the matched region of reads; matched regions are not identical regions, and mismatch and indel could occur, e.g., --mm 2 36 represents ≤ 2 mismatches out of 36 bp
--tail: the maximum bp allowed at each side, two integers to specify the number of tails out of the number of basepairs of the reads, not including "N", e.g., --tail 3 75 represents ≤ 3 bp tails of 75 bp of reads without "N"
--gap: if a read is split, an internal gap (bp) is allowed, default = 5000 bp
--mappingscore: the minimum mapping score, default = 40
--insert: insert range, e.g., 100 600 (default)
--help: help information
Note: We expect the percentage of mapped reads to be ~70% for our example organism.
BAM sort & index
The bam files are required to be sorted and indexed before GATK SNP calling.
## BAM sort & index ##
cd cache
samtools view -u B73.parse.sam | samtools sort -o B73.parse.sort.bam
samtools view -u A188.parse.sam | samtools sort -o A188.parse.sort.bam
samtools index B73.parse.sort.bam
samtools index A188.parse.sort.bam
rm *.sam
cd ../
For one-step running of this step, you can simply run this script:
sh ./workflow/3_SAM_filter.sh
Step 4: Data pre-processing for GATK SNP calling
Before GATK SNP calling, aligned reads need to be pre-processed to meet the GATK requirements.
(Optional) Add labels for alignment reads
In BWA alignment step, we have already set sample names/groups. If this is not added during the BWA alignment, the following step must be added to BAM alignments. The GATK module “AddOrReplaceReadGroups” can be used to add the sample name/group information.
## Add labels for inputs ##
gatk AddOrReplaceReadGroups --java-options '-Xmx24g' \
-I ./cache/B73.parse.sort.bam \
-O ./cache/B73.RG.bam \
-LB A -PL illumina -PU A -SM B73 -ID B73
gatk AddOrReplaceReadGroups --java-options '-Xmx24g' \
-I ./cache/A188.parse.sort.bam \
-O ./cache/A188.RG.bam \
-LB A -PL illumina -PU A -SM A188 -ID A188
Adjustable parameters for AddOrReplaceReadGroups:
-LB: Read-Group library#
-PL: Read-Group platform#
-PU: Read-Group platform unit#
-SM: Read-Group sample name#
-ID: Read-Group ID#
(Optional) MarkDuplicates
The GATK package “MarkDuplicates” is used to mark duplicated reads occurring during sample preparation such as library construction using PCR. The MarkDuplicates tool works by comparing sequences in the five prime positions of both reads and read-pairs in a SAM/BAM file. A BARCODE_TAG option is available to facilitate duplicate marking using molecular barcodes. After duplicated reads are marked, the GATK tool can differentiate the primary and duplicated reads using an algorithm that ranks reads by using the sum of quality scores of each read, which can improve the accuracy of variant calling.
## MarkDuplicates ##
gatk MarkDuplicates --java-options '-Xmx24g' --REMOVE_DUPLICATES true \
-I ./cache/B73.RG.bam \
-M ./cache/B73.metrics.txt \
-O ./cache/B73.RG.RD.bam
gatk MarkDuplicates --java-options '-Xmx24g' --REMOVE_DUPLICATES true \
-I ./cache/A188.RG.bam
-M ./cache/A188.metrics.txt
-O ./cache/A188.RG.RD.bam
## BAM index ##
cd cache
samtools index B73.RG.RD.bam
samtools index A188.RG.RD.bam
cd ../
(Optional) Recalibrate base quality scores
BQSR stands for base quality score recalibration. Using this module, alignment scores can be recalibrated according to known genomic variation data to improve variation calling. In our test samples, this step was skipped. However, the script is provided here.
## Generates recalibration table for Base Quality Score Recalibration (BQSR) ##
gatk BaseRecalibrator \
-I input.RG.RD.bam \
-R reference.fa \
--known-sites sites_of_variation.vcf \
--known-sites another/optional/setOfSitesToMask.vcf \
-O recal_data.table
## Apply base quality score recalibration ##
gatk ApplyBQSR \
-R reference.fa \
-I input.RG.RD.bam \
--bqsr-recal-file recalibration.table \
For one-step running of this step, you can simply run this script:
sh ./workflow/4_data_preprocess.sh
Note: GATK and 1.8.0+ Java environment are required for this step.
Step 5: GATK short variant calling
GATK is a collection of command-line tools for analyzing high-throughput sequencing data with a primary focus on variant discovery. Here, we focus on short variants and use the “HaplotypeCaller” module to call SNPs and indels relative to a reference genome.
Create a GATK index of the reference genome
## gatk index for reference genome ##
samtool faidx input/B73Ref4.fa
gatk CreateSequenceDictionary -R input/B73Ref4.fa -O input/B73Ref4.dict
GATK short variant calling
## GATK SNP Calling (2 samples together) ##
gatk HaplotypeCaller --java-options '-Xmx24g' -R input/B73Ref4.fa -I cache/B73.RG.RD.bam -I cache/A188.RG.RD.bam -O output/B73_A188.raw.0.vcf
Notes:
(1) The GATK short variant calling step can identify both SNPs and short indels; here, we focus on SNPs and filter out indels (Step 6: SNP filtering).
(2) We use the default parameters of GATK HapotypeCaller for short variant calling.
(3) We suggest doing GATK short variant calling with all samples together. If there are too many samples to load at the same time, you can use a bam list file including all sample information instead. An example bam.list file can be found in /input/bam.list; the format of the bam list file consists of the bam file path for each row. The code is as follows:
## GATK Short Variant Calling (with bam list) ##
gatk HaplotypeCaller --java-options '-Xmx24g' \
-R ./input/B73Ref4.fa \
-I ./input/bam.list \
-O ./output/B73_A188.raw.0.vcf
For one-step running of this step, you can simply run this script:
sh ./workflow/5_GATK.sh
Step 6: SNP filtering
To remove indels and retain high-quality SNPs, GATK provides the script module for filtering variants [11].
Select bi-allelic SNPs
## select bi-allelic SNPs ##
vcf=./output/B73_A188.raw.0.vcf
ref=./input/B73Ref4.fa
gatk SelectVariants -R $ref -V $vcf \
--restrict-alleles-to BIALLELIC -select-type SNP \
-O ./output/B73_A188.bi.1.vcf.gz &>./output/B73_A188.bi.log
Filtering variants
## Hard-filtering germline short variants ##
gatk VariantFiltration --java-options '-Xmx24g' \
-R $ref -V ./output/B73_A188.bi.1.vcf.gz \
--filter-expression "QD < 2.0 || MQ < 40.0 || FS > 60.0 || MQRankSum < -12.5 || ReadPosRankSum < -8.0 || SOR > 3.0" \
--filter-name "hard_filter" \
-O ./output/B73_A188.HF.2.vcf.gz
## Extract PASS SNPs ##
gatk SelectVariants --java-options '-Xmx24g' \
-R $ref -V ./output/B73_A188.HF.2.vcf.gz \
--exclude-filtered \
-O ./output/B73_A188.PASS.3.vcf.gz
(Optional) Convert heterozygous SNPs to missing
## (Optional) Convert heterozygous SNPs to Missing ##
gatk VariantFiltration --java-options '-Xmx24g' \
-V ./output/B73_A188.PASS.3.vcf.gz \
-O ./output/B73_A188.mark.hetero.4.vcf.gz \
--genotype-filter-expression "isHet == 1" \
--genotype-filter-name "isHetFilter"
gatk SelectVariants --java-options '-Xmx24g' SelectVariants \
-V ./output/B73_A188.mark.hetero.4.vcf.gz \
--set-filtered-gt-to-nocall \
-O ./output/B73_A188.het2miss.4.vcf.gz
(Optional) Filter SNPs by minor allele frequency (MAF) & missing rate (MR)
## (Optional) Filter SNPs by Minor Allele Frequency (MAF) & Missing Rate (MR) [12]##
vcftools ./output/B73_A188.het2miss.4.vcf.gz \
--maf 0.02 --max-missing 0.2 --recode --recode-INFO-all \
--out ./output/B73_A188.MAF.MR.5.vcf.gz
For one-step running of this step, you can simply run this script:
sh ./workflow/6_SNP_filter.sh
Note: The recommended filtering parameters for SNPs by GATK can be found on the link https://gatk.broadinstitute.org/hc/en-us/articles/360035890471-Hard-filtering-germline-short-variants.
Result interpretation
Read trimming results
The read trimming information is saved in ./cache/*_trim.log files. The percentage of “Both Surviving” represents the ratio of high-quality pair-ended reads that passed read trimming. Here is an example:
Input Read Pairs: 400000
Both Surviving: 400000 (100.00%)
Forward Only Surviving: 0 (0.00%)
Reverse Only Surviving: 0 (0.00%) Dropped: 0 (0.00%)
Read alignment and filtering results
The read alignment and filtering information are saved in ./cache/*.parse.log files. Here is an example:
./cache/B73.sam Total reads in the SAM output 400000
./cache/B73.sam Reads could be mapped 399854
./cache/B73.sam Passing criteria reads 282380
./cache/B73.sam Unmapped reads 1943
GATK SNP calling results
The GATK SNP calling results are saved in ./output/*.vcf files as a VCF format. Here is an example of the VCF format:
CHROM POS ID REF ALT QUAL FILTER INFO FORMAT B73 A188
1 242449 . T C 23.19 PASS AC=2;AF=1.00;AN=2;DP=2;ExcessHet=3.0103;FS=0.000;MLEAC=2;MLEAF=1.00;MQ=46.50;QD=11.60;SOR=0.693;GT:AD:DP:GQ:PL GT:AD:DP:GQ:PL 1/1:0,2:2:6:49,6,0 ./.
The first eight columns of the VCF records (up to and including INFO) represent the properties observed at the level of the variant (or invariant) site. Keep in mind that when multiple samples are represented in a VCF file, some of the site-level annotations represent a summary or average of the values obtained for that site from all samples. Sample-specific information such as genotype and individual sample-level annotation values are contained in the FORMAT column (9th column) and sample-name columns (10th and beyond). More details of the GATK result information can be found in VCF format wiki and https://gatk.broadinstitute.org/hc/en-us/articles/360035531692-VCF-Variant-Call-Format.
Acknowledgments
Cheng, H. and Liu, S. were supported by the USDA NIFA (award no. 2018-67013-28511 and 2021-67013-35724) and by the NSF (awards no. 1741090, 2011500, and 2311738).
Competing interests
The authors declare no competing interests.
References
Shastry, B. S. (2002). SNP alleles in human disease and evolution. J Hum Genet 47(11): 561-566.
Tian, F., Bradbury, P. J., Brown, P. J., Hung, H., Sun, Q., Flint-Garcia, S., Rocheford, T. R., McMullen, M. D., Holland, J. B. and Buckler, E. S. (2011). Genome-wide association study of leaf architecture in the maize nested association mapping population. Nat Genet 43(2): 159-162.
Nielsen, R., Paul, J., Albrechtsen, A. and Song, Y. S. (2011). Genotype and SNP calling from next-generation sequencing data. Nat Rev Genet 12(6): 443-451.
Burland, T. G. (2000). DNASTAR’s Lasergene sequence analysis software. Methods Mol Biol 132: 71-91.
Li, R., Yu, C., Li, Y., Lam, T. W., Yiu, S. M., Kristiansen, K. and Wang, J. (2009). SOAP2: an improved ultrafast tool for short read alignment. Bioinformatics 25(15): 1966-1967.
Li, H., Handsaker, B., Wysoker, A., Fennell, T., Ruan, J., Homer, N., Marth, G., Abecasis, G. and Durbin, R. (2009a). The sequence alignment/map format and SAMtools. Bioinformatics 25(16): 2078-2079.
McKenna, A., Hanna, M., Banks, E., Sivachenko, A., Cibulskis, K., Kernytsky, A., Garimella, K., Altshuler, D., Gabriel, S., Daly, M. and DePristo, M. A. (2010). The Genome Analysis Toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res 20(9): 1297-1303.
Brouard, J. S., Schenkel, F., Marete, A. and Bissonnette, N. (2019). The GATK joint genotyping workflow is appropriate for calling variants in RNA-seq experiments. J Anim Sci Biotechnol 10(1): 1-6.
Bolger, A. M., Lohse, M. and Usadel, B. (2014). Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30(15): 2114-2120.
Li, H. and Durbin, R. (2009b). Fast and accurate short read alignment with Burrows–Wheeler transform. Bioinformatics 25(14): 1754-1760.
De Summa, S., Malerba, G., Pinto, R., Mori, A., Mijatovic, V. and Tommasi, S. (2017). GATK hard filtering: tunable parameters to improve variant calling for next generation sequencing targeted gene panel data. BMC bioinformatics 18(5): 57-65.
Danecek, P., Auton, A., Abecasis, G., Albers, C. A., Banks, E., DePristo, M. A., Handsaker, R. E., Lunter, G., Marth, G. T., Sherry, S. T., et al. (2011). The variant call format and VCFtools. Bioinformatics 27(15): 2156-2158.
Supplementary information
Data and code availability: All data and code have been deposited to GitHub: https://github.com/Bio-protocol/GATK-SNP-Calling
Article Information
Publication history
Accepted: Aug 4, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
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Systems Biology > Genomics > Screening
Plant Science > Plant molecular biology
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5,069 | https://bio-protocol.org/en/bpdetail?id=5069&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
A Step-by-step Protocol for Crossing and Marker-Assisted Breeding of Asian and African Rice Varieties
Yugander Arra
EL Eliza P.-I. Loo
BD B.N. Devanna
MS Melissa Stiebner
WF Wolf B. Frommer
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5069 Views: 429
Reviewed by: Samik BhattacharyaWenrong He Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Academic Radiology Oct 2013
Abstract
Improving desirable traits of popular rice varieties is of particular importance for small-scale food producers. Breeding is considered the most ecological and economic approach to improve yield, especially in the context of pest and pathogen-resistant varieties development. Being able to cross rice lines is also a critical step when using current transgene-based genome editing technologies, e.g., to remove transgenes. Moreover, rice breeders have developed accelerated breeding methods, including marker-assisted backcross breeding (MABB) to develop novel rice varieties with in-built resistance to biotic and abiotic stressors, grain, and nutritional quality. MABB is a highly efficient and cost-effective approach in accelerating the improvement of recipient variety by introgressing desirable traits, especially from landrace cultivars and wild rice accessions. Here, we provide a detailed protocol including video instructions for rice crossing and MABB to introgress target trait(s) of interest into the elite rice line. Further, we also highlight tips and tricks to be considered for a successful crossing and MABB.
Key features
• This protocol provides detailed information on techniques for crossing rice varieties and for breeding rice varieties with new traits
• The protocol includes instructions for making rice crosses as well as MABB
• The protocol provides beginners with detailed instructions including troubleshooting guides
Keywords: Rice Backcross breeding Donor parent Recipient parent Gene introgression MABB
Graphical overview
Illustration of crosses of rice lines and marker-assisted backcross breeding (created with BioRender.com)
Background
Rice is one of the most important food staples for over 3.5 billion people. Classical breeding enables the development of new varieties by trait introgression for important agronomic characters and increased tolerance/resistance to abiotic/biotic stressors. Backcross breeding uses donor and recipient parents (DP and RP, respectively); the DP carries the desired traits to be introgressed into an RP, typically an elite variety. Classical breeding inadvertently introduces undesirable traits from the donor and/or the loss of beneficial RP traits. It is, therefore, necessary to backcross the resulting progeny, i.e., the F1 generation, multiple times with the RP to eliminate as much of the chromosomal segments from the DP that do not carry the trait of interest as possible. Crossovers close to trait gene loci are desirable to eliminate the chance that linked loci adversely impact the resulting variety's performance.
Developing a new rice variety through breeding takes 7–8 years (two seasons per year, maximum of 16 seasons of cultivation) [1]. New varieties must be tested in multilocation field trials to validate trait improvements relative to parental lines before new varieties are registered, which takes another two years. Marker-assisted backcross breeding (MABB) reduces this timeline to three years and reduces linkage drag from the DP [1]. MABB introgresses loci encoding desired traits into the RP and reconstitutes the genome of RP by background selection, eliminating undesirable genomic fragments, i.e., linkage drag, from the DP. MABB uses gene-based/gene-linked molecular markers for foreground; to select the desired trait/loci from DP, it uses background selection (to determine the percentage recovery of RP genome) and recombinant selection (to determine the presence of linkage drag) [2,3]. The success of MABB depends on, among other things, differentiating the genome of DP from the genome of RP. Typically, simple sequence repeat (SSR) markers are used as polymorphic molecular markers for differentiating genomic backgrounds [4]. SSR markers are short DNA motifs (2–6 nucleotide repeats) that exhibit variable repeat numbers in the genome. SSRs are abundant, multi-allelic, co-dominant, hypervariable, and relatively uniformly distributed in the genome. SSR markers have become important and widely used in rice breeding [2,5,6].
Apart from its application in plant breeding, crosses are an essential step for eliminating transgenes introduced via genome editing. Genome editing offers the potential to target multiple loci for providing broad-spectrum resistance, nutritional fortification, and yield improvement in crops [7–10]. Notably, the term new breeding methods, frequently used in the context of genome editing, is not an optimal choice, since it is rather an alternative for generating genetic variation, in particular targeted modifications compared with the use of naturally occurring mutations or chemical- or radiation-induced mutagenesis; the breeding process, on the other hand, remains in the hands of breeders and requires their technologies and experience. In countries with appropriate regulations, genome-edited crops are treated as equivalent to crops generated by classical breeding [11].
Materials and reagents
Biological material
Two rice parents, DP and RP, to develop new desirable rice varieties.
Choice of DP
Reproductive isolation can limit the ability to cross diverse rice varieties or landraces [12,13]. Incompatibility between DP and RP crossing leads to sterility in subsequent crosses [14]. Therefore, choose a DP genetically compatible with the RP that produces fertile F1 seeds after the cross. In rice, crosses involving indica and japonica subspecies often produce sterile seeds. Similarly, crosses between Oryza sativa spp. indica (RP) and the DP from a wild rice species/Oryza glaberrima are reported to show embryo abortion/sterility; for example, Komboka (accession: IR05N221) (RP), and FARO 44 (accession: WAB0004879) (DP).
The DP should possess at least a few characters of distinctness, uniformity, and stability (DUS) to the RP. DUS characters play a key role during varietal nomination and evaluation [15]. MABB-derived lines with at least 1–2 DUS characters to the RP are desirable. The choice and variety of releases also depend on DUS characters.
Choose a DP that provides a novel resource for pest/disease resistance, stress tolerance, yield attributes, agronomic characters, or grain quality.
In most cases, DP is either a landrace or a wild accession. Note that it is challenging to make a successful cross between some rice species due to crossing barriers (e.g., Oryza sativa and Oryza glaberrima).
Choice of RP
Choose an elite RP that possesses desirable characteristics including plant phenotype, protection against biotic/abiotic factors, or utilization traits (grain and cooking quality attributes).
The improved RP with higher gain yield and better economic returns tends to have higher acceptance among large-scale farmers.
Laboratory supplies
Murashige and Skoog basal salt mixture (Duchefa, catalog number: M0254)
Sucrose (Sigma-Aldrich, catalog number: S0389)
Phytagel (Sigma-Aldrich, catalog number: P8169)
peqGOLD Plant DNA Mini Kit (VWR, catalog number: 13-3486-02)
GoTaq G2 Green Master Mix (Promega, catalog number: M7823)
Agarose (Sigma-Aldrich, catalog number: A6013)
GeneRuler 1 kb Plus DNA Ladder, ready to use (Thermo Scientific, catalog number: SM1334)
Equipment
Analytical and precision balances (Precisa Gravimetrics AG, Series 321LS, model: LS 2200C)
Laboratory reagent bottles with screw cap, 1,000 mL (Duran®, catalog number: 218015455)
Benchtop pH meter, WTWTM inolab TM 7110 (Fisher Scientific, catalog number: 11731381)
Magnetic stirring bars, cylindrical 25 × 8 mm (VWR, catalog number: 442-0483)
Autoclave (Systec, model: 3850 EL)
Sterile workbench (Thermo Scientific, model: HeraguardTM Eco)
Petri dishes (Sigma-Aldrich, catalog number: P5606-400EA)
Tissue culture vessel, MagentaTM GA-7 (VWR, catalog number: SAFSV8505-100EA)
Hand-held wooden cereal dehusker
Centrifuge tubes with screw cap, 15 mL (Thermo Scientific, catalog number: 339650)
Permanent marker pen
Precision tweezers (VWR, catalog number: 232-1220)
Precision forceps, extra sharp, curved (VWR, catalog number: BOCH1940)
Growth chamber with 27 °C, 16/8 h day/night photoperiod (Percival, model: CU-41L5)
Head-band magnifying glasses (may be advantageous for proper emasculation)
Seed paper bag (11.5 × 6 cm)
Retro floor lamp with E27 Bulb including 3 lights tree floor lamp
Plastic pots, 11 × 11 × 12 cm (Mayer shop, catalog number: 722009)
Plastic tray to keep plastic pots, 50 × 32 × 6 cm (Mayer shop, catalog number: 749112)
Whatman filter paper (VWR, catalog number: 516-0593)
Height adjustable stool, useful during emasculation
Spray bottle for water, 1000 mL (Roth, model: IC8T.1)
Nanodrop (Thermo Fisher, catalog number: 13-400-519)
Thermocycler (Bio-Rad, model: T100TM, catalog number:1861096)
EasyPhor Medi gel electrophoresis system 3GT 15 × 10 cm (Biozym, catalog number: 615162)
Gel documentation unit (Vilber, model: E-BOX CX5 TS)
Drying and heating cabinets with mechanical adjustment (Binder, catalog number: E28)
DanKlorix® original (2.8 g/100 mL sodium hypochlorite)
Procedure
Germination and growth conditions
Dehusk the rice seeds using a small hand-held wooden cereal dehusker (https://orcainstruments.com/product/palm-husker/).
Transfer the dehusked seeds into a 15 mL centrifuge tube and add 7–8 mL of 70% ethanol.
Incubate the seeds for 2 min at room temperature shaking at 80 rpm.
Discard the ethanol and add 7–8 mL of 4% sodium hypochlorite into the tube, followed by 5 min of incubation at room temperature with shaking at 80 rpm.
Discard the bleach solution into a waste collection reagent bottle on a clean sterile bench and rinse the seeds repeatedly (≥ 5 times) with sterile water until no traces of bleach are noticeable. Finally, air-dry the sterilized seeds on sterilized filter papers on a clean workbench for 15–20 min.
Transfer the dry sterilized rice seeds onto Petri dishes (20 seeds/Petri dish) containing autoclaved half-salt strength Murashige and Skoog salt (½ salt strength MS) media supplemented with 1% sucrose.
Germination in Petri dishes appears to improve germination efficiency. To start seed germination, incubate the seeds (on the Petri dishes) in a plant growth chamber at 27 under dark conditions for three days.
After three days, transfer germinated seeds into a tissue culture vessel, MagentaTM GA-7, filled with 50 mL of ½ salt strength MS media and incubate at 27 with 16/8 h light/dark for six days.
After six days, transfer the seedlings to 11 × 11 × 12 cm (L × B × H) pots (one seedling per pot) filled with soil and grow under greenhouse conditions (28 ± 1 ; relative humidity between 60% and 80%) until the booting stage [16]. Water regularly and apply fertilizers as required [16]. Remove old and/or dried leaves, if present, to avoid hindrance during the crossing process.
Emasculation
The ideal criteria for emasculation are: (i) plants reached the post-booting stage, (ii) show healthy panicles, (iii) have no obvious signs of anthesis yet, and (iv) floral organs (stigma and stamen) are still located inside the lower half of the spikelet (Figure 1). In our case, 100–110-day-old plants were found to be the perfect stage for emasculation; however, booting is dependent on the growth conditions and rice varieties differ substantially with respect to flowering time.
Figure 1. The rice panicle represents the floral organs inside of the spikelets. Stigma and stamen are located inside the lower half of the spikelet.
Day 1: Spikelet opening leading to anthesis is triggered by high light intensity and humidity. Therefore, the plant growth conditions, including light intensity, temperature, and humidity, should be taken into consideration when deciding the appropriate time for emasculation. It is recommended to perform emasculation either in the early morning hours (5–7 am) or early evening hours (5–7 pm).
Carefully inspect rice plants for panicles with more than 50% exsertion (panicle exsertion: the distance between the leaf cushion of the flag node and the neck-panicle node; Figure 2). A correct selection of panicles is key to avoiding self-pollination.
Figure 2. Different stages of rice panicles from the booting stage to emergence from flag leaf. A. No panicle exsertion. B. 40% panicle exsertion. C. >70% panicle exsertion. D. Emasculated spikelet. E. Cross-cutting of spikelet.
Remove the leaves surrounding the panicle, except the flag leaf, to prevent interference during the crossing. Remove young spikelets at the base of the panicle and flowered spikelets in the upper part of the panicle to achieve the optimum number (50–60) of spikelets per panicle (Video 1).
Video 1. Removing spikelets from rice panicles before crossing
For emasculation, it is recommended to cut individual spikelets at a 35–40-degree angle just above the anthers with a pair of sterile scissors. Be careful not to damage the stigma while cutting the spikelet (Video 2).
Video 2. Emasculation of individual spikelets
Carefully remove each stamen (six stamens in total) from a rice spikelet using sharp tip forceps. Do not damage or break the anthers in the spikelet to avoid pollen contamination, hence self-fertilization. It is important to emasculate all the flowers in the panicle: a single anther is sufficient to self-pollinate the entire panicle.
Dusting and bagging
The DP plants with 30%–50% of flowering panicles should be used as a source of anthers for dusting. While carrying out the crosses in greenhouse conditions, care should be taken to position the recipient and the donor plants in such a way that the flowers of these plants are at similar heights for ease of crossing.
The inflorescence or panicle of DP plants close to flowering was exposed to high light intensity (>400 μE m-2 s-1) by lowering the height of the overhead light source for 15–30 min to encourage flowering. Exposure to high light intensity also increases the microclimate temperature of the panicle to 30–40 . Inflorescence with flowers with anthers at the stage of dehiscence should be used for dusting on emasculated panicles of the RP (Figure 3, video 3).
Video 3. Dusting of emasculated panicles of the RP with pollen from the DP
Figure 3. Panicles at (a) pre- and (b) post-flowering stages of individual plants.
It is important to begin dusting from the top of the emasculated panicle and slowly slide the DP panicle down the RP panicle with a gentle stroke. Immediately after dusting, cover the panicle with pre-labeled white paper bags [important information on labels: name of the cross, date, and filial (F) generation].
Depending on the availability of anthers, dusting can be repeated with the same panicle on the same or the next day to increase the percentage seed set.
Tip: Use a flag leaf to support the paper bag as florets are not supposed to be folded after the crossing.
Day 2–6: Provide plants with optimal growth conditions.
Day 7: If desired, one can observe the seed set one week after dusting. Gently remove the paper bag from the panicle. Healthy, immature, greenish-colored seeds are typical. The percentage seed set can be calculated by a ratio of filled to unfilled spikelets. Keep in mind to re-bag the panicle with the paper bag.
Day 15: Since the spikelets were trimmed, mature seeds could develop without husk (Figure 4a). Dried lemma and palea contribute to the light or dark yellow coloration of the husk. Since developing seeds lack husk, care should be taken to keep insects, pests, and diseases from damaging the seeds.
Day 22: All mature seeds develop a light brown color (Figure 4b).
Day 27: Seeds (F1 generation) with a maximum of 30% moisture [17] are ready for harvest (Figure 4b). A moisture tester helps measure the rice seed moisture (e.g., https://best4grain.at/product/moisture-meter-farmpoint/?lang=en).
Figure 4. Different stages of seed setting on recipient parent after pollination. A. Fifteen days after pollination. B. Twenty-two days after pollination. Insets represent the different maturity stages of rice seed after pollination.
Seed storage
Incubate harvested rice seeds at 40 for 2 days under dark conditions to reduce their moisture content to below 12%. Higher moisture content leads to seed damage due to quick degradation of nutrients, which affects seed quality, storage life, and germination.
The dried seeds can be either used for germination or preserved at 4 for long-term storage. Harvested F1 seeds (#100–1,000) can be dried in an incubator set at 40 , allowing for faster drying.
Marker-assisted backcross breeding (MABB)
The MABB is an application of DNA-based molecular markers to monitor and choose the target loci (foreground selection) and to accelerate the RP genome recovery (background selection) during backcrosses [2]. MABB was mainly deployed to develop near-isogenic lines (NILs) with far greater precision than classical breeding. MABB accelerates the recovery of the RP genome and reduces the number of backcross generations. These plants carry the target loci in the genetic background of RP, and the percent recovery of the RP genome can be estimated in these lines [18].
Foreground selection
Incubate F1 seeds from the cross at 40 for 2 days to facilitate uniform germination.
Germinate the F1 seeds as described in section A.
Identify the positive heterozygous plants, i.e., foreground selection, and harvest approximately 100 mg of healthy leaf tissues from 25–30-day-old plants (F1 plants, and both DP and RP) with sterilized scissors into 2 mL microcentrifuge tubes containing 3–5 glass beads.
This protocol recommends the peqGOLD Plant DNA Mini Kit for genomic DNA isolation from harvested leaf samples. Other DNA extraction kits can be adopted.
Measure the genomic DNA concentration using Nanodrop (for details, see step E4). Use gene-specific, co-dominant DNA markers (preferably SSR/SNPs) to PCR amplify target gene(s) from the isolated genomic DNA (https://archive.gramene.org/markers/microsat/).
Access the hybridity of the F1 plants based on the PCR amplicon sizes compared to RP and DP. Homozygous dominant, homozygous recessive, or heterozygous for the target gene(s) of interest are typical in the F1 generation (Figure 5). Pick the heterozygous F1 plants for backcross breeding (Figure 6).
Figure 5. Validation of heterozygosity in the F1 generation by gene-specific markers (foreground selection). M: 1 kb Plus ladder; DP: Donor parent, 270 bp; RP: Recipient parent, 150 bp; Arrow indicates heterozygous for both parents 270bp/150 bp.
Figure 6. Crossing scheme adopted for marker-assisted backcross breeding. Three backcrosses bring maximum recipient parent genome recovery into improved lines by implementing molecular markers (SSR/SNPs). BC1-3: number of backcrosses; F2-4: number of generations for advancing the population by selfing.
Background selection
Recombinant selection helps to reduce the linkage drag and to identify the breeding lines with maximum RP genome recovery [19]. In the present protocol, we chose a minimum of two polymorphic SSR markers (differentiating the DP and RP) for each arm of the 12 rice chromosomes. Also, we made sure that the chosen markers were at a uniform distance along the chromosome for the better recovery of the RP genome. For genome recovery calculation, we chose breeding material that was confirmed for the presence of candidate target gene(s) through foreground selection and performed PCR analysis using polymorphic SSR markers. Further, to overcome the linkage drag, we designed a set of SSR primers close to the target gene(s) loci, identified the polymorphic SSRs differentiating the RP and the DP, and used them again to screen the breeding material in the advanced (BC2F3 generation) generation for better RP genome recovery and overcome linkage drag using the software Graphical Geno Types (GGT) Version 2.0 [20].
Selection of simple sequence repeats (SSRs)
More than 18,000 SSR markers have been reported for rice [21,22]. Information for the SSR makers is available on the GRAMENE database (https://archive.gramene.org/markers/).
Choose around 500 SSR markers distributed uniformly on all 12 rice chromosomes (with at least 1 Mb distance between each marker) for PCR analysis of DP and RP.
Isolate genomic DNA from DP and RP using a DNA isolation kit or CTAB method [23].
Genotype DP and RP with selected SSRs using PCR and identify markers showing polymorphism between DP and RP.
Choose at least four polymorphic SSRs between DP and RP for each rice chromosome.
Use identified polymorphic SSRs to genotype F1 up to BC2-3F2 generations to identify homozygous lines with maximum recovery of the RP genome (> 97%).
Advance lines to BC2F4 for registration or nomination and possible varietal release.
Genotyping for foreground or background selection
Collect leaf samples with sterilized scissors (5–8 cm long) from 25–30 days-old individual breeding plants in 2 mL microcentrifuge tubes.
Isolate genomic DNA from collected leaf samples using an available plant DNA isolation kit or CTAB method.
Measure the concentration of genomic DNA with Nanodrop and adjust the concentration to 10 ng/μL.
Prepare PCR master mix (10 μL volume) with 1 μL of template DNA (10 ng/μL), 0.5 μL of each primer (5 mM of each gene-specific primer solution), 5 μL of GoTaq® Green Master Mix, and 3 μL of nuclease-free water.
Run a PCR with the following thermal profile: Initial denaturation at 95 °C for 3 min, followed by 30 cycles consisting of denaturation at 95 °C for 30 s, annealing at 55 °C for 30 s, and primer extension at 72 °C for 1 min (1 kb/min). The final extension is at 72 °C for 5 min.
Confirm the presence or absence of targeted genes via electrophoresis.
Take a photograph of the gel with a gel documentation unit and record the data for the presence of the gene (using gene-specific markers) and also the allelic nature of these genes based on the fragment size.
For background selection, use the SSR marker data generated for each breeding line and calculate the RP genome recovery using the software Graphical Geno Types (GGT) Version 2.0 [20].
Tips and tricks
For beginners, start with a bold grain shape (length to breadth ratio value, <2.0 mm) rice genotype/cultivar and practice removing anthers (emasculation) before attempting a major crossing program. It is more challenging to remove anthers from fine grain and medium slender gains.
Use the headband magnifier to facilitate observation for better emasculation.
It is best to remove at least the top 3–5 spikelets from each secondary branch and remove anthers with forceps to avoid anthesis and self-pollination.
Try to cut the spikelet with an upright triangle shape from both sides of the spikelet as it facilitates the opening of the spikelet and emasculation.
Choose rice DP plants with 30%–40% flowering for anthers.
Spray the panicles with water before moving them to high light intensity to facilitate fast anthesis.
After dusting, it is best practice to clean and remove old, dried leaves from the base of the plant. Dried leaves serve as physical contact for ants and insects, which will damage the early-stage green embryos developed after a successful cross.
Monitor crossed panicles regularly for insects and ants both on the panicle or inside the bag covering the panicle.
Try to keep crossed plants in a clean area and fill the pots or tray with water to the maximum level (>95%) for good seed setting.
For fine grain–type rice plants, use 40–50 spikelets/panicles for emasculation and dust with two flowered panicles.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Yugander et al. (2018). Incorporation of the novel bacterial blight resistance gene Xa38 into the genetic background of elite rice variety Improved Samba Mahsuri. PLoS One.
Schepler-Luu et al. (2023). Genome editing of an African elite rice variety confers resistance against endemic and emerging Xanthomonas oryzae pv. oryzae strains. eLife.
Acknowledgments
This work was supported by grants from the Bill & Melinda Gates Foundation (1155704), the Alexander von Humboldt Professorship (WBF), Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) under Germany´s Excellence Strategy – EXC-2048/1 – project ID 390686111 (CEPLAS), and fellowships to YA by the Alexander von Humboldt Foundation, CEPLAS and DBT-Ramalingaswami Re-entry Fellowship (2024), Department of Biotechnology, New Delhi, India.
Author contributions
Y.A., E.L., and B.F.W. developed the concept. Y.A., B.N.D., and S.M. performed experiments. Y.A., E.L., B.N.D., and B.F.W. wrote the manuscript. All authors have approved to approve the final version of the manuscript.
Competing interests
The authors declare no conflict of interest.
References
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Article Information
Publication history
Received: Apr 9, 2024
Accepted: Jul 10, 2024
Available online: Aug 26, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,070 | https://bio-protocol.org/en/bpdetail?id=5070&type=0 | # Bio-Protocol Content
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Mouse Renal Artery Catheterization for Local Delivery of Drugs in Capsulated or Free Forms
OS Olga A. Sindeeva
OG Olga I. Gusliakova
EP Ekaterina S. Prikhozhdenko
NS Natalia A. Shushunova
GS Gleb B. Sukhorukov
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5070 Views: 301
Reviewed by: Olga KopachKomuraiah Myakala Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Pharmaceutics May 2022
Abstract
Arterial delivery to the kidney offers significant potential for targeted accumulation and retention of cells, genetic material, and drugs, both in free and encapsulated forms, because the entire dose passes through the vessels feeding this organ during the first circulation of blood. At the same time, a detailed study on the safety and effectiveness of developed therapies in a large number of experimental animals is required. Small laboratory animals, especially mice, are the most sought-after in experimental and preclinical testing due to their cost-effectiveness. Most of the described manipulations in mice involve puncturing the walls of the abdominal aorta or renal artery for direct administration of solutions and suspensions. Such manipulations are temporary and, in some cases, result in long-term occlusion of the aorta. Ultimately, this can lead to disruption of blood flow as well as functional and morphological changes to the kidneys. In addition, few of these protocols describe targeted delivery to the kidney. The presented protocol involves the injection of test substances or suspensions through the renal artery into one of the kidneys. The catheter is implanted into the femoral artery and then advanced into the abdominal aorta and renal artery within the vessels. In this case, the integrity violation of the renal artery or abdominal aorta is absent. Occlusion of the renal artery is necessary only immediately at the time of injection to minimize the entry of the injected substance into the aorta. This protocol is similar to the clinical procedure for delivering a catheter into the renal artery and is designed for real-world operating conditions.
Key features
• The protocol involves implantation of a catheter into the vascular system through a puncture of the femoral artery, similar to the clinical procedure.
• The catheter is moved inside the vessels without puncture or ligation to the aorta or renal artery.
• The protocol involves only a short-term block of the blood supply to the target kidney (the time required for direct administration of the drug).
• Suitable for chronic experiments on mice, since the catheter is removed from the vascular system immediately after drug administration.
Keywords: Femoral artery catheterization Renal artery catheterization Targeted drug delivery Local drug accumulation Mouse kidney Target kidney
Graphical overview
Mouse renal artery catheterization through the femoral artery
Background
Arterial delivery of drugs to the kidneys in free and encapsulated forms has enormous potential to increase local drug concentrations and reduce the side effects of therapy on other organs. The choice of therapy parameters (active substance, pharmaceutical form, dosage regimen, volume, etc.), as well as understanding drug accumulation and retention in the kidneys during arterial delivery, requires a wide range of complex studies on animals. In this regard, small laboratory animals, especially mice, are the most attractive and economical.
The success of the study directly depends on the correctness of the arterial delivery procedure, which is especially difficult for mice due to their small size. Incorrectly performed injections in the renal artery could disrupt the kidney blood supply and lead to tissue damage. The vast majority of drug delivery protocols through the renal artery are based on ligation and direct injection into the abdominal aorta [1]. Chen et al. reported that direct injection of the rAAV2-GFP vector into the rat abdominal aorta using 45 min ligation of renal vein and abdominal aorta led not only to successful transfection of tubular epithelial cells but also to focal areas of chronic interstitial infiltration with mononuclear cells, tubular atrophy, and scar formation [2]. Authors suggested that these changes are most likely the result of ischemic injury inherent in the procedure and not a consequence of recombinant adeno-associated viral vector administration or an adverse immune response to it or the transgene. A similar protocol for mice with short-term occlusion (several minutes) and puncture of the abdominal aorta was presented by Dahlqvist et al. [3]. Zaw Thin et al. developed a percutaneous minimally invasive ultrasound-guided renal artery injection to enhance cell delivery to mice kidneys [4]. This procedure appears to be less invasive (only puncture of muscles and skin with a syringe) than the others but requires a special ultrasound imaging system. In addition, in this protocol, the renal artery wall is punctured with a 29-gauge needle (0.33 mm diameter). Note that the diameter of the mouse renal artery is in the range of 0.3–0.4 mm [5], which determines a high risk of significant blood loss after injection without the ability to stop bleeding. At the same time, the authors reported 70% success.
In our protocol, we describe the catheterization of the mouse renal artery by inserting a thin catheter into a small puncture in the femoral artery and then approaching the left renal artery through the abdominal aorta [6–8]. This protocol is similar to the standard clinical approach [9–11]. It does not require blockade of the target kidney blood flow, disruption of the integrity, and ligation of the renal artery or abdominal aorta, unlike other described methods [1–4, 12, 13]. This protocol requires microsurgery skills but can be extremely useful for studies in which maintaining a normal blood supply to the target kidney as well as vascular integrity is critical.
Materials and reagents
Biological materials
Balb/c mice 8–10 weeks old (20–25 g weight) were obtained from the vivarium of the Saratov State Medical University. Other mouse strains could also be used in this protocol.
Reagents
Corneregel 5% (Dr. Gerhard Mann, catalog number: 4030571000075)
Zoletil 100 Virbac SA (Valdepharm / Delpharm Tours, catalog number: SAGARPA Q-0042-306)
Xyla (Xylazine 2%) 50 mL (Interchemie werken, catalog number: IX2)
Iodine alcohol solution for external use, alcohol 5% (Pharmaks, catalog number: 322645 or Rigas Farmfabrika, catalog number: 110004)
Sodium chloride solution (saline) for injection 0.9% 5 mL (Grotex LLC, catalog number: 250309 or Hameln, UK, catalog number: SC005)
Evans Blue (Sigma-Aldrich, catalog number: E2129)
70% alcohol antiseptic solution (BioPharmKombinat, catalog number: N1x1)
Solutions
5% Evans Blue solution in saline (see Recipes)
Recipes
5% Evans Blue solution in saline (1 mL)
Reagent Final concentration Quantity or Volume
Evans Blue 5% (w/v) 5 mg
Sodium chloride solution (0.9%) 0.9% (w/v) 1 mL
Total (optional) 5% (w/v) 1 mL
Laboratory supplies
Nylon surgical threads (USP: 5/0, EP: 1) (Mosnitki, catalog number: 5/0(1))
Cotton pads, cleanic soft & сomfort (Harper Hygienics, catalog number: 96190075)
Cotton buds (Harper Hygienics, catalog number: 96190075)
Fixing adhesive plaster 2.5 cm × 5 m (Nordeplast, catalog number: 1145541)
Sterile surgical cotton wool 100 g (Hygrovata, catalog number: GOST 5556-81)
Hair removal cream for sensitive skin Veet (Reckitt Benckiser, catalog number: BP_29060)
Suture material Prolene 5-0 (Ethicon, Johnson & Johnson, catalog number: W8310)
BD Micro-Fine Plus Demi insulin syringes with sterile interior 0.3 mm (30G) × 8 mm, 0.3 mL (Becton Dickinson, catalog number: 320829)
Polyurethane intravascular tubing 32ga/0.8Fr, 0.13 mm × 0.25 mm (Instech Laboratories, Inc., catalog number: BTPU-010)
Polyethylene tubing PE-10, 0.28 mm × 0.61 mm (BD Intramedic, catalog number: 22-204008)
Equipment
Dressing forceps 10 cm (Figure 1A)
Tweezers 11 cm (Figure 1B)
Angle tweezers 10.5 cm (Figure 1C)
Toothed tweezers 14 cm (Figure 1D)
Tweezers 14 cm (Figure 1E)
Micro scissors 14 cm (Figure 1F)
Microneedle holder 14 cm (Figure 1G)
Stereomicroscope (Nexcope, model: NSZ-608)
LED ring light (Lomo-microsystems, model: LED-144A)
Digital camera MS-8.3 USB 3.0 with matrix 1/1.2" SONY (Lomo-microsystems, model: MS-8.3)
Analytical balance (OHAUS, model: Pioneer PR224)
Trimmer Wahl Micro Lithium (Wahl GhbH, model: 5640)
Figure 1. Catheterization tools. Dressing forceps 10 cm (A), tweezers 11 cm (B), angle tweezers 10.5 cm (C), toothed tweezers 14 cm (D), tweezers 14 cm (E), micro scissors 14 cm (F), and microneedle holder 14 cm (G).
Software and datasets
For video recording: Pylon Viewer (6.3.0.23157, 02.11.2021)
For video processing: Shotcut (24.02.29, 29.02.2024)
Procedure
Preparation of catheter for the renal artery catheterization
Prepare the components of the catheter (Figure 2A):
Unwind the nylon surgical threads and take one of the three fibers as a ligature (10–12 cm).
Cut 5 cm of Polyurethane (PU) intravascular tubing 32ga/.8Fr, 0.13 mm × 0.25 mm (thin catheter).
Cut 7 cm of Polyethylene tubing PE-10, 0.28 mm × 0.61 mm (thick catheter).
Insert the thin catheter into the thick one to a depth of 0.5 cm (Figure 2B).
Tie a ligature tightly to secure the thin catheter. The patency of the catheter should not be impaired.
Cut the tip of the thin catheter at a 45° angle (Figure 2C).
Figure 2. Preparation of catheter for the renal artery catheterization. (A) Components of the catheter: ligature, thin catheter (5 cm)—Polyurethane intravascular tubing 32ga/.8Fr, 0.13 mm × 0.25 mm; thick catheter (7 cm)—Polyethylene tubing PE-10, 0.28 mm × 0.61mm. (B) A thin catheter is inserted into a thick catheter to a depth of 0.5 mm and fixed with a ligature. (С) The tip of the thin catheter is cut at a 45° angle. (D) Checking the tightness and patency of the catheter using saline solution. (E) Saline solution does not flow at the junction of the thin and thick catheters. (F) Saline solution flows from the tip of a thin catheter.
Check the tightness and patency of the catheter by injecting saline solution into it using an insulin syringe with sterile interior and needle 0.3 mm (30G) × 8 mm (Figure 2D):
Check that saline solution does not leak at the junction of the thin and thick parts of the catheter (Figure 2E).
Check that the saline solution leaks at the tip of the thin catheter (Figure 2F).
The internal volume of the catheter, manufactured according to the described method, is 5 μL. This must be considered when administering the drugs. If other tubes are used to create the catheter, the internal volume should be measured or calculated based on internal diameter and length.
Mouse preparation for catheterization
Prepare a cotton roll for the mouse:
Take three cotton pads (Figure 3A).
Take 3 cm of fixing adhesive plaster 2.5 cm × 5 m (Figure 3B).
Prepare a roll of cotton pads and adhesive plaster (Figure 3С).
Figure 3. Preparation of cotton roll for mouse. (A) Three cotton pads, (B) fixing adhesive plaster 2.5 cm × 5 m, (C) and cotton roll for mouse.
Inject general anesthesia into the mouse intraperitoneally [Zoletil mixture (40 mg per kg) and 2% Xylazine (10 mg per kg)]. Instead of Xylazine, 2% Rometar can be used.
Wait 5–7 min and check the hind limb reflex. If there is no hind limb reflex, begin the operation; otherwise, wait a few more minutes.
Apply Corneregel 5% or other protective gel to mouse eyes.
Place the mouse on the cotton roll and remove hair from the left side of the body using a trimmer and then hair removal cream for sensitive skin (Figure 4A). Remove the cream thoroughly using cotton pads and saline or water. Do not apply the cream for more than 30–60 s to avoid damaging the skin.
Place the animal on its back and secure its paws using adhesive plaster (Figure 4B).
Remove hair on the inner side of the left paw using a trimmer and then hair removal cream (Figure 4С).
Figure 4. Mouse preparation for catheterization. (A) Hair is removed from the left side of the mouse body above the left kidney. (B) The mouse is fixed lying on its back, and (C) the hair on the inner side of the left paw is removed.
Catheter implantation into the left femoral artery and abdominal aorta
Treat the mouse's left paw skin with 70% alcohol antiseptic solution.
Cut the skin over the femoral vein and artery of the mouse's left paw using micro scissors and toothed tweezers (Figure 5A). Do not allow the vessels to dry out during the entire operation. Moisten them periodically with saline.
Separate the vessels from the connective tissue on the right side using a 30 G needle (Figure 5B).
Place the angle tweezers under the vessels on the right side (Figure 5C) and completely separate them from the connective tissue and muscle (Figure 5D).
Carefully separate the nerve and femoral vein from the femoral artery using tweezers (Figure 5E).
Place two ligatures under the femoral artery (Figure 5F).
Tie the lower ligature, stretch it, and secure it using a plaster.
Prepare an upper ligature to quickly secure the catheter, but do not tie it at this stage.
Puncture the femoral artery using a 30G needle (Figure 5G).
Insert the tip of the thin catheter (cut side up) through the puncture into the femoral artery using two tweezers, as shown in Figure 5H. The catheter should first be connected to a syringe with a 30G needle and completely filled with saline solution without air bubbles (Figure 2D).
Inject a small amount (20–30 µL) of saline through the catheter into the artery to inflate it.
Slowly advance the catheter through the femoral artery using two tweezers (Figure 5I) until the junction of thin and thick catheters (Figure 5J). At this point, the tip of a thin catheter will be positioned in the abdominal aorta. Be careful not to squeeze the catheter too hard to avoid deformation.
Check that the catheter is inside the artery. Pull the plunger of the syringe toward you so that arterial blood enters the catheter (Figure 5K). Then, inject saline into the catheter (Figure 5I).
Secure the catheter with an upper ligature, but do not tie it completely (Figure 5L). The ability to move the catheter inside the vessel must be maintained.
Figure 5. Catheter implantation into the left femoral artery and abdominal aorta. (A) Skin incision over the femoral vein and artery. (B) Separation of vessels from connective tissue on the right with a 30 G needle. (C, D) Separation of nerve and vessels from connective tissue using angle tweezers. (E) Separation of the nerve from the femoral artery. (F) Ligation of the femoral artery. (G) Femoral artery puncture. (H) Inserting the tip of a thin catheter through a puncture of the femoral artery. (I) Pushing a thin catheter into the femoral artery, (J) until the junction of thin and thick catheters. (K) Checking the location of the catheter in the artery. Arterial blood flows into the catheter when the syringe plunger is pulled back. (L) Fixation of the catheter in the femoral artery with an upper ligature.
Catheter implantation into the left renal artery
Remove the fixing adhesive plaster from the mouse's paws and lower ligature.
Turn the mouse over, being careful not to pull on the catheter. Place the mouse on the cotton roll in the same position as shown in Figure 4A.
Treat mouse's skin with 70% alcohol antiseptic solution.
Pull back the skin to the right of the kidney using toothed tweezers (Figure 6A) and make a 1 cm long incision parallel to the spine (Figure 6В).
Pull back the muscle to the right of the kidney using toothed tweezers and make a 1 cm incision (Figure 6C).
Pull apart the connective and fatty tissue to visualize the kidney (Figure 6D).
Gently spread the muscle and tissue using dressing forceps to visualize the renal vein (Figure 6E). The renal artery is closely adjacent to the renal vein to its left, but the location of the vessels may differ slightly in different animals.
Figure 6. Catheter implantation into the left renal artery. (A, B) Skin incision along the spine over the kidney. (C) Muscle incision over the kidney. (D) Removal of connective and fatty tissue to reveal the kidney. (E) Renal vein. (F) Abdominal aorta and renal artery cleared of connective and fatty tissues. (G) Palpation of a thin catheter inside the abdominal aorta, searching for the tip of the catheter. (H) Transferring the tip of a thin catheter from the abdominal aorta to the renal artery. (I) The tip of a thin catheter inside the renal artery. (J) Target kidney before, (K) during, and (L) after saline injection.
Carefully clean the junction of the renal arteries and the abdominal aorta using tweezers and sterilized cotton buds (Figure 6F). Cotton buds should be moistened with saline solution. Try to minimize the duration of this procedure to avoid damage to the vessels due to repeated mechanical pressure on them.
Locate the tip of the thin catheter in the abdominal aorta by palpating its entire length with tweezers (Figure 6G). If the catheter is palpable throughout the visualized portion of the abdominal aorta, gently pull the 2–3 mm catheter out of the femoral artery. Then palpate the aorta again with forceps. Pull the catheter, preferably holding the thin part with tweezers.
Gradually pull out the catheter until its tip is in the aorta 2–3 mm above the renal artery.
The saline syringe should be replaced with a drug syringe immediately prior to implantation of the catheter tip into the left renal artery.
Turn the mouse in front of you so that its spine is perpendicular to you and its kidney is on the left (Figure 6H).
Move the kidney upward (closer to the liver) using wetted cotton buds to increase the angle between the renal artery and the abdominal aorta.
Stretch the area of the abdominal aorta and renal artery at the junction of these vessels using two forceps. Transfer the tip of the catheter from the aorta to the renal artery within the vessels (Figure 6H).
Try to position the tip of the catheter at the beginning of the renal artery before the bifurcation (Figure 6I) to avoid filling half of the kidney during injection. The kidney should maintain its normal color and blood supply at all stages of the operation, before catheter insertion into the renal artery and solution administration.
When the catheter is inserted into the renal artery, the color of the kidney will change slightly due to incomplete occlusion of the renal artery by the catheter (Figure 6J). Note that the blood continues to flow into the kidney before injection. Minimize blockade of kidney blood flow as much as possible during all procedures.
During drug solution administration, the catheter should be held with tweezers, pressing the walls of the arteries against it, in order to minimize the entry of the injected solution into the abdominal aorta (Figure 6K).
Inject 10–20 µL of solution at a rate of 2–3 µL/s through the catheter. In this case, the kidney should become uniformly light (Figure 6K).
Open the tweezers to allow blood to flow into the kidney (Figure 6L).
Removing the catheter from the vascular system and applying sutures
Return the mouse to its back.
Moisten the femoral artery with saline solution using wet cotton buds.
Remove the catheter from the vessels (Figure 7A). Be careful not to tear the femoral artery.
Pull the upper ligature (to block bleeding), tie it above the puncture, and cut off the excess part of the ligature, leaving 3–4 mm from the knot (Figure 7B).
Inject an antibiotic solution (broad spectrum, for example, penicillin 100 mg/kg) into the abdominal cavity to prevent peritonitis, if the experimental plan allows this.
Sew up the skin on the left hind paw using a continuous suture (Figure 7С).
Place the mouse on the cotton roll in the same way as shown in Figure 4A.
Sew up the muscle above the kidney using a continuous suture (Figure 7D).
Sew up the skin using an interrupted suture (Figure 7E).
Treat surgical sutures with antiseptic, for example, 5% iodine alcohol solution.
Place the mouse in a cage and wait until it recovers from anesthesia.
Place half the cage on top of a heat source to facilitate the maintenance of average normal body temperature.
Figure 7. Removing the catheter from the vascular system and applying sutures. (A) Removing the catheter. (B) Tying the upper ligature. (C) Applying a continuous suture to the skin of the left hind paw. (D) Applying a continuous suture to the muscles above the kidney. (E) Applying an interrupted suture to the skin above the kidney. (F) Active mouse 24 h after surgery.
Until the blinking reflex has returned, eye lubricant (Corneregel 5%) should be provided.
Check the mouse 24 h after surgery. It should maintain normal activity and mobility (Figure 7F).
Examine the animal daily for signs of grooming, general appearance, posture, and locomotor activity.
The main stages of the mouse renal artery catheterization through the femoral artery, as described in sections B, С, D, and E, are presented in Video 1.
Video 1. Main stages of the mouse renal artery catheterization through the femoral artery
Validation of protocol
We initially validated this protocol by renal artery injection of 5% Evans Blue solution using more than three technical replicates. As a result, we obtained stable staining of the target kidney (Figure 8A, B). This effect was particularly noticeable when compared with the contralateral kidney (Figure 8C).
Figure 8. Targeted delivery of 5% Evans Blue solution in saline to the kidney using a renal artery catheterization protocol. (A) Target kidney before and (B) right after renal artery injection of 5% Evans Blue solution in saline. (C) Target and contralateral kidney 5 min after injection.
This protocol has been also validated in the following research articles:
Prikhozhdenko et al. [6]. Target delivery of drug carriers in mice kidney glomeruli via renal artery. Balance between efficiency and safety. Journal of Controlled Release.
Abdurashitov et al. [7]. Optical coherence microangiography of the mouse kidney for diagnosis of circulatory disorders. Biomedical Optics Express.
Gusliakova et al. [8]. Renal Artery Catheterization for Microcapsules’ Targeted Delivery to the Mouse Kidney. Pharmaceutics.
The injection of microcapsule suspensions according to the presented protocol was accompanied by local accumulation of 20% of the entire administered dose of capsules in the target kidney. Most capsules accumulated in the glomeruli of the kidneys [6] but were washed out of the capillary network over time [8]. In this case, a high fluorescent signal remained in the target kidney for several days, most likely due to the diffusion of the encapsulated fluorescent cargo through the polymer walls of the containers into the tissue in the first hours after injection [6,8]. Moreover, the accumulation of the substance in the target kidney in encapsulated form was significantly more effective than in free form. Using a laser speckle contrast analysis system and optical coherence tomography, it was shown that a correctly selected dosage of containers did not lead to irreversible changes in the blood flow of the target kidney, as well as simulating surgery with the introduction of saline [6–8]. In this case, not only the number of capsules administered but also their size is of particular importance when choosing the dosage [8]. Histological analysis also did not reveal morphological changes in the renal tissue after administration of the microcapsule suspension at the correct dose.
General notes and troubleshooting
General notes
The protocol requires the experimenter to have experience in microsurgery.
All experimental procedures on animals must be performed under general anesthesia. The depth of anesthesia should be periodically monitored by the hind limb reflex.
The vessels should be regularly moistened with saline solution at all stages of the operation. Do not let them dry out.
The total duration of the operation should not exceed 45–60 min to avoid the risk of developing thrombosis of the hind limbs.
Do not stretch the thin catheter (Polyurethane intravascular tubing 32ga/.8Fr, 0.13 mm × 0.25 mm) and avoid contact with organic solvents such as alcohol-based antiseptic solutions (at all stages of surgery and catheter preparation). This significantly changes the mechanical properties of the catheter and prevents it from being pushed into the vessels.
The process of catheter removal requires ligation of the femoral artery, and re-catheterization of this artery further upstream is possible, but difficult. At the same time, access to the aorta and renal artery can also be obtained through the right femoral artery if it is necessary to repeat the operation. Nevertheless, it is worth considering that each subsequent operation has a significant impact on the mouse’s organism. Therefore, repeat surgery is not recommended unless there is a significant need.
Troubleshooting
Problem 1: The catheter is stuck and cannot be moved further along the femoral artery into the abdominal aorta.
Possible cause: The catheter rested on or pierced the bend of the artery.
Solution: Do not try to force the catheter through. This will puncture the arterial wall and cause the catheter to exit into the surrounding tissue. The catheter should move smoothly, perhaps with little pressure. If movement is difficult, use one or more of the solutions listed:
Make sure that the mouse's left hind paw and lower ligature are tight enough. The artery must be well straightened to reduce the number of bends. Inject a small volume of saline (30–40 µL) into the artery at a pressure. Then, carefully push the catheter again.
Extend the left hind paw along the body and secure the paw and ligature to reduce the number of bends.
Carefully pull the thin catheter out of the artery 2–4 mm (hold it with tweezers as close to the artery puncture site as possible without squeezing or deforming it). Inject a small volume of saline (30–40 µL) into the artery at a pressure of 20–30 µL. Then, pull the plunger of the syringe back slightly to ensure that arterial blood is entering the catheter, and the tip of the catheter is in the artery. If blood does not flow, continue to gently pull out the catheter until blood flows into it when you pull the plunger of the syringe. Then, inject the solution and push the catheter again as needed.
Problem 2: Blood does not flow into the catheter when the syringe plunger is pulled out.
Possible cause: Most likely, the catheter pierced the artery and exited into the surrounding tissue as a result of excessive pressure on the artery wall in the bend.
Solutions:
Try to clear the tissue along the catheter and find the puncture site of the artery.
If the puncture occurs deep in the abdominal cavity, the animal must be sacrificed straightaway.
If the puncture occurs before the abdominal cavity, you can pull out the catheter, tie a second lower ligature next to the puncture, and try to catheterize the femoral artery again.
Problem 3: The catheter cannot be pushed into the femoral artery; it wrinkles.
Possible cause: Most likely, you slightly stretched the thin catheter during surgery, pulled the catheter out of the femoral artery, or squeezed it hard with tweezers and deformed it.
Solution: Remove the catheter completely from the vessels and replace it with a new one. The catheter must have sufficient mechanical rigidity to navigate arterial bends.
Acknowledgments
This research was supported by the Russian Science Foundation (project no. 23-75-10070, https://rscf.ru/project/23-75-10070/).
Competing interests
The authors declare no competing interest.
Ethical considerations
The laboratory animal studies were performed according to the rules of Saratov State Medical University (Ethics Committee Protocol No. 7, dated 2 February 2021).
References
Takabatake, Y., Isaka, Y. and Imai, E. (2008). Renal artery injection for delivery of biological materials to the glomerulus (Methods in Renal Research Paper). Nephrology. 13(1): 23–26.
Chen, S., Agarwal, A., Glushakova, O. Y., Jorgensen, M. S., Salgar, S. K., Poirier, A., Flotte, T. R., Croker, B. P., Madsen, K. M., Atkinson, M. A., et al. (2003). Gene Delivery in Renal Tubular Epithelial Cells Using Recombinant Adeno-Associated Viral Vectors. J Am Soc Nephrol. 14(4): 947–958.
Dahlqvist, U., Tomic, T. T., Söderberg, M., Stubbe, J., Enggaard, C., Ericsson, A., Zhou, A. X., Björnson Granqvist, A. and William-Olsson, L. (2021). Direct Drug Delivery to Kidney via the Renal Artery. J Visualized Exp.: e3791/61932–v.
Zaw Thin, M., Ogunlade, O., Comenge, J., Patrick, P. S., Stuckey, D. J., David, A. L., Lythgoe, M. F., Beard, P. and Kalber, T. L. (2020). Stem cell delivery to kidney via minimally invasive ultrasound-guided renal artery injection in mice. Sci Rep. 10(1): 7514.
Lorenz, J. N., Lasko, V. M., Nieman, M. L., Damhoff, T., Prasad, V., Beierwaltes, W. H. and Lingrel, J. B. (2011). Renovascular hypertension using a modified two-kidney, one-clip approach in mice is not dependent on the α1 or α2 Na-K-ATPase ouabain-binding site. Am J Physiol-Renal Physiol. 301(3): F615–F621.
Prikhozhdenko, E. S., Gusliakova, O. I., Kulikov, O. A., Mayorova, O. A., Shushunova, N. A., Abdurashitov, A. S., Bratashov, D. N., Pyataev, N. A., Tuchin, V. V., Gorin, D. A., et al. (2021). Target delivery of drug carriers in mice kidney glomeruli via renal artery. Balance between efficiency and safety. J Controlled Release. 329: 175–190.
Abdurashitov, A. S., Prikhozhdenko, E. S., Mayorova, O. A., Plastun, V. O., Gusliakova, O. I., Shushunova, N. A., Kulikov, O. A., Tuchin, V. V., Sukhorukov, G. B., Sindeeva, O. A., et al. (2021). Optical coherence microangiography of the mouse kidney for diagnosis of circulatory disorders. Biomed Opt Express. 12(7): 4467.
Gusliakova, O. I., Prikhozhdenko, E. S., Plastun, V. O., Mayorova, O. A., Shushunova, N. A., Abdurashitov, A. S., Kulikov, O. A., Abakumov, M. A., Gorin, D. A., Sukhorukov, G. B., et al. (2022). Renal Artery Catheterization for Microcapsules’ Targeted Delivery to the Mouse Kidney. Pharmaceutics. 14(5): 1056.
Siqueira, D. E. D. and Guillaumon, A. T. (2019). Angiography for Renal Artery Diseases. In: Angiography. IntechOpen. doi: 10.5772/intechopen.79232.
Ruggiero, N. J. and Jaff, M. R. (2009). Renal artery stenting—which patients will benefit? Nat Rev Cardiol. 6(11): 675–676.
Lu, T., Lin, B., Zhang, Y. p., Zhang, J. h., Luo, J. W., Tang, Y. and Fang, Z. T. (2023). Eighteen cases of renal aneurysms: Clinical retrospective analysis and experience of endovascular interventional treatment. Front Surg. 10: e1106682.
Lai, L. W., Moeckel, G. and Lien, Y. H. (1997). Kidney-targeted liposome-mediated gene transfer in mice. Gene Ther. 4(5): 426–431.
Ullah, M., Liu, D. D., Rai, S., Razavi, M., Choi, J., Wang, J., Concepcion, W. and Thakor, A. S. (2020). A Novel Approach to Deliver Therapeutic Extracellular Vesicles Directly into the Mouse Kidney via Its Arterial Blood Supply. Cells. 9(4): 937.
Article Information
Publication history
Received: Mar 31, 2024
Accepted: Jul 29, 2024
Available online: Aug 16, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,071 | https://bio-protocol.org/en/bpdetail?id=5071&type=0 | # Bio-Protocol Content
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Peer-reviewed
Development and Characterization of Primary Brain Cultures from Japanese Quail Embryos
SZ Shaden Zoabi *
AB Achinoam Blau *
Shai Berlin
(*contributed equally to this work)
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5071 Views: 320
Reviewed by: Sébastien GillotinSilvia Olivera-BravoHong Lian
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Original Research Article:
The authors used this protocol in Communications Biology Mar 2023
Abstract
Cell cultures play a crucial role in neuroscience research, facilitating the elucidation of the complexities of cellular physiology and pathology. The relative simplicity in producing cultures and the accessibility to cells that the cultures provide, in contrast to in vivo settings, allow users to manipulate and monitor cells more easily at higher throughputs and lower costs. These are ideal for screening purposes and electrophysiological characterizations. Despite the prevalence of methodologies for producing brain cultures from various animal models, rodents in particular, approaches for culturing neurons (and glia) from birds are less established or completely absent as in the case of the Japanese quail model. Here, we present a unique culturing protocol for brain cells (e.g., neurons at different maturation levels, such as progenitor cells, excitatory and inhibitory neurons, microglia, and endothelial cells) from entire forebrains of Japanese quail embryos for high-throughput screening of viral vectors in vitro and other various purposes. Following dissection and digestion methods uniquely suited for avian brains, we tailored the growth media and culturing surface to allow the survival of quail brain cultures for more than three weeks in vitro.
Key features
• We introduce a detailed protocol for producing primary brain cultures from quail embryos' forebrains for up to 30 days.
• We show that the cultures support in vitro viral transfections effectively.
• We demonstrate the use of the cultures for rapid (days) screening for suitable viruses for quail brain cells, electrophysiological characterizations, and single mRNA sequencing.
Keywords: Primary cell culture Quail Avian Neurons In vitro Forebrain
Graphical overview
Background
Bird species offer intriguing prospects for neuroscience research due to their sophisticated cognitive abilities and specialized behaviors [1–6]. Despite differences in neuroarchitecture and neuronal densities in avian brains when compared with mammals [4,7], birds demonstrate remarkable and distinctive behavioral performances, including long-distance navigation [8], imprinting, homing, food-caching, and song-learning [9,10], to name a few. Consequently, these provide unique opportunities for comparative studies aiming to elucidate the cellular mechanisms underlying these capabilities [6]. Indeed, these realizations are reflected by the growing interest in avian neuroscience [1,3,9,11–18]. Among the different avian species explored, our focus centers on the domestic Japanese quail (Coturnix japonica). The quail offers several advantages over other birds due to its small size, rapid sexual maturation, and frequent egg-laying; advantages that make this species highly suitable for routine experimentations [17]. Importantly, the ground-dwelling nature of quails makes them ideal for studying birds’ spatial navigation by reducing dimensionalities (from 3D to 2D) [19]. In spite of these advantages and the extensive use of quails in the field of developmental biology [17,20], this species remains relatively underexplored in neuroscience [5,19]. This is partly attributed to the absence of methods and tools to manipulate and examine quail neurons [21], in particular, the lack of efficient viral vectors for neuronal transduction [5,17]. To screen for suitable viral vectors, we choose not to pursue in vivo exploration, as is common practice, due to the considerably large number of animals and time this task necessitates (to overcome animal-to-animal variability, variable injection performances, and other technical limitations). Instead, we envisioned that primary brain cultures would present a more rapid and cost-effective approach for examining multiple viruses in parallel [22].
Primary brain cell cultures constitute a valuable tool in neuroscience [23,24]. Brain cultures faithfully preserve many complex cellular behaviors, electrophysiological properties, and phenotypic traits, as observed in vivo, despite the loss of the precise network architecture and organization in the tissue, as is exclusively found in vivo [23-28]. Nevertheless, the alteration in tissue architecture, which includes the removal of the blood-brain barrier (BBB) and extracellular matrix, proves to be advantageous as it enhances cellular accessibility and reduces time to expression. Indeed, cultures are ideal for high-throughput screening of pharmacology, which should likely translate toward the screening of different viruses [22].
We have initially attempted to produce primary cultures based on standard protocols for culturing cells from the brains of neonatal rodents (such as rats and mice, e.g., [29]) or chicken embryos (e.g., [30,31]). However, these attempts were unsuccessful. Consequently, we decided to optimize the procedure by experimenting with embryos collected from different days in ovo (DIO), modifying the culturing media, and assessing cell growth directly on plastic dishes instead of glass coverslips. After successfully establishing suitable conditions for maintaining viable cultures for several weeks (up to 30 days), we leveraged them for screening several commonly employed viruses in neuroscience research, including adeno-associated viruses (AAV), lentiviruses, and baculovirus. We further harnessed the cultures for the rational engineering of a novel AAV variant, specifically tailored for Japanese quail neurons (denoted AAV1*) [22]. Lastly, we characterized the diverse cellular populations in the cultures through use of electrophysiology, calcium imaging, and single-cell mRNA sequencing [22].
Materials and reagents
Biological materials
Fertilized Japanese quail eggs (Coturnix japonica). The eggs we employ are obtained from our quail colony. Eggs are collected from cages with male and female quails. Nevertheless, there are multiple commercial providers of fertilized quail eggs; however, the number of farms and thereby commercial availability may vary from one country to another.
Reagents
B-27 serum-free supplement (50×) liquid (Gibco, catalog number: 17504-044)
Pen/Strep solution (Sartorius, catalog number: 03-031-1B)
L-glutamine (GlutaMAX) (Gibco, catalog number: 35050-038)
Neurobasal-A medium (Gibco, catalog number: 10888-022)
Papain from papaya latex (Sigma-Aldrich, catalog number: 9001-73-4)
Phosphate buffered saline (PBS) (Sartorius, catalog number: 02-023-5A)
Poly-D-lysine hydrobromide (PDL) (Sigma-Aldrich, catalog number: 27964-99-4)
DNase I set (Geneaid, catalog number: DNS300)
Sylgard 184 silicone elastomer (Sigma-Aldrich, catalog number: 761036-5EA)
DMEM/F-12 (Ham) (Sartorius, catalog number: 01-170-1A)
Solutions
0.2% PDL (see Recipes)
Quail neurobasal medium (QNBM) (see Recipes)
Digestion solution (see Recipes)
Papain stock (see Recipes)
Recipes
Note: Prepare all solutions in a laminar flow cell culture hood.
0.2% PDL (store at -20 °C)
Reagent Final concentration Amount
PDL 2 mg/mL 100 mg
Double-distilled water (DDW) n/a 50 mL
Total n/a 50 mL
Quail neurobasal medium (NBM) (store at 4 °C)
Reagent Final concentration Amount
B-27 2% 5 mL
Pen/Strep 1% 2.5 mL
L-glutamine 0.25% 625 µL
Neurobasal-A medium n/a Complete to 250 mL
Total n/a 250 mL
Digestion solution (prepare on the day of extraction)
Reagent Final concentration Amount
Papain stock 33 U/mL 1 mL
PBS n/a 2 mL
DNase I 57 U/mL 85 µL
Total n/a 3.085 mL
Papain stock
Reagent Final concentration Amount
Papain 10 U/mg 100 mg
PBS n/a 10 mL
Total n/a 10 mL
Laboratory supplies
Spray bottle with 70% ethanol (Bio-Lab Chemicals, catalog number: 000521020500)
Tissue culture dish (60 mm × 15 mm) (Corning, catalog number: 430166)
Tissue culture dish (100 mm × 20 mm) (Corning, catalog number: 430167)
Microtubes, 1.7 mL (Corning, catalog number: MCT-175-C)
Conical bottom tubes, 15 ml (Greiner Bio-One, catalog number: 188261)
Conical bottom tubes, 50 ml (Greiner Bio-One, catalog number: 227270)
Cell strainer 40 μm (LifeGene, catalog number: LG-CSS010040S)
Parafilm (Sigma-Aldrich, catalog number: P7543)
Delicate task Kimwipes (Kimberly-Clark professional, catalog number: 34120)
Minutien Pins, 1 cm, 0.0175 mm tip (Fisher Scientific, catalog number: 2600215)
Equipment
Stereomicroscope system (Zeiss Stemi 2000, Fisher Scientific, catalog number: 10331390)
Microbiological safety cabinet class II (Thermo Scientific, catalog number: 51025757)
Isotemp water bath (PolyScience, catalog number: E12020004)
HERAcell 150i CO2 incubator (Thermo Fisher Scientific, catalog number: 50116047)
Gilson (or equivalent) pipettes and tips (P20/P200/P1000) (Fisher Scientific)
Dumont #5 forceps (Roboz surgical store, catalog number: RS-4905) (Figure 1, #1)
Curved dissecting scissors (Stoelting, catalog number: 52132-11P) (Figure 1, #2)
Plastic Pasteur pipette (blunt tip obtained by trimming pipette ~1 cm from tip) (Sigma-Aldrich, catalog number: 747775 (Figure 1, #3)
Vannas spring scissors (Fine Science Tools, catalog number: 15000-08) (Figure 1, #4)
Iris forceps (Fine Science Tools, catalog number: 11064-07) (Figure 1, #5)
Fine-angled forceps (Stoelting, catalog number: 52102-02P) (Figure 1, #6)
Egg incubator, 37 °C (DMP Engineering, model: INCA 100)
Figure 1. Fine surgical tools required for the removal of eggshell and embryo from egg
Procedure
PDL plate coating
Note: Do this in a sterile laminar flow cell culture hood.
Fill 60 mm Corning dishes with 5 mL of 0.2% PDL solution (see Recipes).
Place the plates in a 37 °C incubator overnight.
Critical: Non-satisfactory coating of the plates can lead to cell dissociation from the surface of the plate and neuronal death causing unhealthy, nonviable cultures.
Collect the PDL solution using a pipette (the solution can be recycled) until plates are empty.
Wash the plates three times using sterile DDW.
Allow residual DDW to evaporate and the plates to dry (leave in the laminar flow cell culture hood).
Seal each plate with parafilm to keep sterile.
Keep coated plates at 4 °C for later use. Plates can be stored for several weeks.
Silicone plate coating
Note: Do this in a sterile laminar flow cell culture hood.
Prepare a 60 mm Corning dish coated with 7 mm silicone, using the Sylgard 184 silicone elastomer kit. The manufacturer’s instructions are included and simple to follow.
Keep overnight at room temperature.
Embryo extraction and brain dissection from quail eggs
Obtain fresh fertilized eggs. Fertilized eggs cannot be distinguished from unfertilized ones at this stage (Figure 2A).
Figure 2. Egg shell removal and embryo extraction from quail eggs. A. Clean quail eggs post-incubation. B. Eggshell penetration. C. Circular eggshell crack, letting the “cap” loose. D. Quail embryo in dish, together with yolk sac and blood vessels. E. Separating embryo from yolk sac and blood vessels using scissors. F. Embryo picked up for transfer with plastic Pasteur pipette.
Store eggs in a suitable incubator at 37 °C with 70%–80% humidity for 7–9 days.
Spray eggshells with 70% ethanol and wipe dry (gently, using Kimwipes), before placing inside the laminar hood.
Note: From this point onward, all procedures are performed in the laminar hood.
Using Dumont #5 forceps (Figure 1, #1), gently penetrate the eggshell in its upper quarter (Figure 2B).
Crack the eggshell with a circular forceps movement, letting the eggshell’s “cap” loose (Figure 2C).
Pour egg content into a 100 mm Corning dish.
Using curved dissection scissors (Figure 1, #2), separate the embryo from the yolk sac and cut the interacting blood vessels (Figure 2D, E).
Transfer the embryo with a blunt plastic Pasteur pipette (Figure 1, #3; Figure 2F) into a 100 mm Corning dish filled with DMEM (Figure 3A).
Figure 3. Embryo forebrain extraction. A. Isolated embryo in a Petri dish, filled with DMEM. B. Embryo decapitation using dissection scissors. C. Silicone-coated Petri dish and fixation pins (dashed box). D. Isolated embryo head in DMEM, pinned to the silicone surface. Inset: closer view of pins (arrowhead). E. Forebrain dissection using forceps and scissors. Inset: closer view of the right forebrain (dashed region). F. One isolated forebrain (without meninges).
Decapitate the embryo with curved dissection scissors (Figure 3B).
Transfer the head of the embryo using a plastic Pasteur pipette to a 60 mm Petri dish coated with silicone and filled with DMEM.
Immobilized the head using Minutien Pins inserted in each eye (Figure 3C, D) and place under a light microscope.
With the Vannas spring scissors (Figure 1, #4), create a mid-sagittal incision from the nape to the eye line (Figure 3E). Peel the soft tissues overlaying the brain with Iris forceps (Figure 1, #5).
Slip the Vannas spring scissors under the forebrain and gently cut through the tissues that are connected to it (Figure 3E, inset).
Using Iris forceps and fine-angled forceps (Figure 1, #5 & #6, respectively), separate the forebrain from the head.
Clean the two hemispheres gently from residual connective tissue and meninges by pealing the meningeal layers with the Iris and the fine-angled forceps (Figure 1, #5 & #6).
Critical: Partial pealing and removal of the meninges can cause fibroblast overgrowth and dominate the culture.
Transfer the clean hemispheres to a 35 mm Petri dish filled with 2 cm of DMEM (Figure 3F) using a plastic Pasteur pipette (Figure 1, #3).
Repeat steps C4–16 until the desired number of hemispheres is obtained. See Troubleshooting.
Tissue dissociation
Place the isolated forebrains in a 15 mL conical tube containing the digestion solution (3.085 mL; see Recipes) using a plastic Pasteur pipette (Figure 1, #3).
Incubate at 37 °C for 1 h (no shaking is needed).
Critical: Incubation of cells for less than 1 h may lead to insufficient dissociation of the tissue, which will result in the appearance of cell clumps in the culture, whereas longer incubations (>1 h) may lead to excessive cell (especially neuronal) death.
Gently remove the solution with a 5 mL pipette without disturbing the precipitated forebrains.
Remove the residual solution with a 1,000 μL plastic tip.
Wash the forebrains by adding PBS to the tube. Repeat three times.
After the third wash, remove the PBS completely using a 1,000 μL plastic tip and replace it with 2 mL of quail neuro-basal medium (NBM) (see Recipes).
Manually dissociate the tissue by gentle trituration with a large, 5 mL plastic pipette tip (×15 times).
Then, triturate the solution gently with a smaller 1,000 μL plastic pipette tip until a homogenous solution is obtained (with no visible tissue fragments).
Place a 40 μm cell strainer on a 50 mL conical tube and prewash it with 1 mL of quail NBM.
Critical: Prewashing the cell strainer prevents cells from sticking to the strainer.
Transfer the entire solution containing the dissociated brains onto the cell strainer placed with a 1,000 μL plastic tip.
After passing the entire solution through the cell strainer, wash the strainer with additional quail NBM to elute the remaining cells from the strainer. The final volume of the filtered (strained) solution should be ~5 mL of quail NBM per one quail’s hemisphere. The optimal density for seeding is 5 × 105 cells/cm2. See Troubleshooting.
Plating
Transfer 5 mL of strained solution onto a sterile 60 mm tissue culture dish that has been pre-coated with 2 mg/mL PDL.
Note: Cells do not adhere to glass coverslips even if the coverslips have been pre-coated with PDL. They clump after several hours and die.
Place the seeded plates in a standard biological cell culture incubator at 37 °C and 5% CO2 for 1 h to ensure cell attachment to the surface of the plates.
After 1 h, gently remove the medium by aspiration and replace it with fresh and pre-warmed quail NBM.
Critical: Aggressive handling and removal of the medium can cause cells to detach from the dish.
Place the plates back in the incubator for 72 h.
After 72 h, replace half the medium with fresh and pre-warmed quail NBM.
Repeat step E4 every other day.
Individual cells should be easily discernible in cultures after one day in vitro (Figure 4A and B, left top and bottom panels). Cells should begin sprouting processes within this time frame. Cellular clumps are an indication of a dying culture (Figure 4A, right panel compared to right panel in B).
Figure 4. Viability of primary cultures grown on glass or plastic after one week in vitro. A, B. Cells grown in vitro at days 1 and 4 post-dissection, on PDL-coated glass coverslips in Petri dishes (A) or directly on the Petri dish (also coated with PDL) (B). The top-right image in A shows a clumping of cells, which does not appear in B. Insets in B show a closer view of the cells’ morphology in culture. Note the development of processes (bottom left) and the appearance of neurons (bottom right). C. MAP2-immunostaining of quail neurons at 14 days in vitro (DIV).
Validation of protocol
This protocol is robust and reproducible. Number of replicates done is >30. Primary neuronal cultures were employed for electrophysiological recordings (patch clamp), calcium imaging, high-throughput viral infection screening, immunohistochemical staining, and single-cell mRNA sequencing experiments (see Figures 1, 2, 3, 4, 6e–g, Suppl. Figure 2 [3]).
General notes and troubleshooting
General notes
This protocol was also found to be suitable for producing primary brain cultures from the forebrains of age-matched chicken embryos (see suppl. Figure 8 in [3]). We therefore assume it may also be suitable for different bird species.
Sources of variability between different cultures may arise from the initial conditions and age of the embryos. Embryos undergoing healthy development are likely to produce more viable cultures, compared to embryos with delayed development. Cultures prepared from poorly developing embryos may yield sparser cultures.
Troubleshooting
Cell density in culture: We recommend dissociating and plating one hemisphere per plate (i.e., one hemisphere per 5 mL of culture medium) (Figure 4A, B; left panels). We noted that this yields favorable conditions for cellular growth, without excessive density of cells. We find that the optimal seeding density for viability of the culture is 5 × 105 cells/cm2. To increase cell density, simply increase the number of hemispheres per plate, for instance plating 1.5 hemispheres per plate. For sparser cultures, dilute the cell suspension in a larger volume of medium. These adjustments may offer strategic means to tailor cell density to experimental requirements, thereby influencing cellular interactions, proliferation rates, and experimental outcomes.
Acknowledgments
Funding: Support was provided by the Rappaport Family Thematic grant (S.B. and Y.G.).
The protocol was first described and validated in Zoabi et al. [3]. A custom-made AAV1 variant (AAV1-T593K) enables efficient transduction of Japanese quail neurons in vitro and in vivo. Communications Biology, 6(1): 337.
Competing interests
The authors declare no competing interests.
Ethical considerations
Animal experiments were approved by the Technion Institutional Animal Care and Use Committee (permit no. IL-157-11-17 and IL-19-10-143) and all experiments strictly followed the approved guidelines.
References
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Article Information
Publication history
Received: May 15, 2024
Accepted: Aug 11, 2024
Available online: Aug 21, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,072 | https://bio-protocol.org/en/bpdetail?id=5072&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Iterative Immunostaining and NEDD Denoising for Improved Signal-To-Noise Ratio in ExM-LSCM
LA Lucio Azzari *
MV Minnamari Vippola
SN Soile Nymark
TI Teemu O. Ihalainen
EM Elina Mäntylä *
(*contributed equally to this work)
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5072 Views: 571
Reviewed by: Xiaoyi ZhengLei GaoDevashish DWIVEDI
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Original Research Article:
The authors used this protocol in Molecular Biology of the Cell Aug 2023
Abstract
Expansion microscopy (ExM) has significantly reformed the field of super-resolution imaging, emerging as a powerful tool for visualizing complex cellular structures with nanoscale precision. Despite its capabilities, the epitope accessibility, labeling density, and precision of individual molecule detection pose challenges. We recently developed an iterative indirect immunofluorescence (IT-IF) method to improve the epitope labeling density, improving the signal and total intensity. In our protocol, we iteratively apply immunostaining steps before the expansion and exploit signal processing through noise estimation, denoising, and deblurring (NEDD) to aid in quantitative image analyses. Herein, we describe the steps of the iterative staining procedure and provide instructions on how to perform NEDD-based signal processing. Overall, IT-IF in ExM–laser scanning confocal microscopy (LSCM) represents a significant advancement in the field of cellular imaging, offering researchers a versatile tool for unraveling the structural complexity of biological systems at the molecular level with an increased signal-to-noise ratio and fluorescence intensity.
Key features
• Builds upon the method developed by Mäntylä et al. [1] and introduces the IT-IF method and signal-processing platform for several nanoscopy imaging applications.
• Retains signal-to-noise ratio and significantly enhances the fluorescence intensity of ExM-LSCM data.
• Automatic estimation of noise, signal reconstruction, denoising, and deblurring for increased reliability in image quantifications.
• Requires at least seven days to complete.
Keywords: Iterative indirect immunofluorescence staining Signal-to-background ratio Expansion microscopy Signal processing Denoising
Graphical overview
Overview of indirect iterative immunofluorescence staining (IT-IF) procedure
Background
Expansion microscopy (ExM) has emerged as a tempting super-resolution microscopy technique for fluorescence imaging of cell and tissue organization. Along with other super-resolution modalities, ExM now extends optical imaging to the nanoscale, yielding unprecedented biological insights within molecularly crowded environments such as the nucleus. In ExM, fixed and gel-embedded samples are isotropically expanded in water by 4× to 10×, allowing imaging beyond the classical diffraction limit of light microscopy and resolving structures in near-molecular resolution. As a result, ExM allows a lateral resolution of ~70 nm with laser scanning confocal microscope (LSCM); the more recently developed iterative ExM including a second gel step allows 10× improvement, achieving a resolution of ~25 nm [2]. Recently, ExM has also been optimized for clinical specimens. The expansion pathology has been speculated to enable routine nanoscale imaging of pathological samples aiding clinical research [3].
However, ExM imaging of immunofluorescence-based samples has drawbacks related to technical difficulties in obtaining the optimal density of the fluorophores. ExM imaging of the samples results in a substantially weaker signal due to the dilution of the fluorophores caused by the sample expansion; the dilution scales to the third power of the (linear) expansion factor. Thus, expansion significantly reduces image brightness and creates noisy images with low signal-to-noise ratio (SNR). Conventional indirect immunostaining can be insufficient, leading to low-density labeling of the targets and, depending on the antibody type, often includes marked off-target staining of the background, also contributing to low image contrast and signal-to-background ratio (SBR) [4,5]. The accessibility of the target epitopes can be reduced, further lowering the image quality [6,7]. Also, super-resolution imaging methods pushing this barrier, such as structured illumination microscopy, pose challenges to the sample properties. When imaging biological samples in the sub-diffraction scale, low labeling density can lead to long acquisition times, higher excitation laser powers, and higher detector sensitivities, leading to increased noise and photobleaching.
We addressed these problems and developed an indirect iterative immunostaining (IT-IF) method for fluorescent labeling of ExM samples. The IT-IF counteracting the expansion-induced decrease in sample fluorescent intensity and signal quality significantly improves the sufficient staining of molecular structures aiding in structural analyses. We argue that the IT-IF approach facilitates nanoscopy, the detection and quantitative analysis of sub-resolution-sized molecular structures, especially within the nucleus. We show how post-image processing through noise estimation, denoising, and deblurring (NEDD) improves the low signal quality in ExM [8] even better than conventional deconvolution and provide step-by-step instructions on how to use the NEDD image processing tool to perform noise reduction on LSCM data. We prove that IT-IF leads to increased signal intensity without compromising the SBR, advancing super-resolution imaging of highly compact intranuclear structures. Finally, we exploit these methods to reveal nanoscopic structural details of nuclear lamina network organization.
Materials and reagents
Reagents
Paraformaldehyde (PFA), 20% (Electron Microscopy Sciences, catalog number: 15713-S)
Primary antibody used in this study: Mouse anti-Lamin A/C (E-1) (Santa Cruz Biotechnology, catalog number: sc-376248)
Secondary antibody used in this study: AlexaTM-conjugated goat anti-mouse IgG (H+L) cross-adsorbed secondary antibody (Alexa FluorTM 488) (Thermo Fisher Scientific, catalog number: A-11008)
TritonTM X-100 (Sigma-Aldrich, Merck, catalog number: X100-100ML)
Bovine serum albumin (BSA) (PAN Biotech, catalog number: P06-139210)
Sodium acrylate (Sigma-Aldrich, Merck, catalog number: 408220)
Acrylamide (0.40 g/mL, 40%) (Bio-Rad Laboratories, catalog number: 1610140)
N,N'-Methylenebisacrylamide (Sigma-Aldrich, Merck, catalog number: 1015460100)
Sodium chloride (Sigma-Aldrich, Merck, CAS #7647-14-5)
N,N,N',N'-Tetramethylethylenediamine (TEMED, 100%) (Sigma-Aldrich, Merck, catalog number: 411019)
Acryloyl X – SE (10 mg/mL in dry DMSO) (Thermo Fisher Scientific, catalog number: A20770)
4-Hydroxy-TEMPO (Sigma-Aldrich, Merck, catalog number: 176141)
Tris base (Sigma-Aldrich, Merck, catalog number: 10708976001)
Proteinase K (New England Biolabs, catalog number: P8107S)
Ammonium persulphate (APS, powder) (Sigma-Aldrich, Merck, catalog number: A3678)
High-melt agarose (1%, w/v) (Thermo Fisher Scientific, catalog number: J61123.22)
Phosphate buffered saline, with Ca2+/Mg2+ (PBS, 10×) (Gibco, Thermo Fisher Scientific, catalog number: J62036.K2)
EDTA, 10 mM (Promega, catalog number: A2631)
Guanidine HCl (8 M) (Thermo Fisher Scientific, catalog number: 24115)
dH2O
Absolute EtOH
Solutions
1× PBS (see Recipes)
Permeabilization solution A (see Recipes)
Permeabilization solution B (see Recipes)
Antibody diluent containing blocking agent (see Recipes)
Tris-HCl, 50 mM, pH 8 (see Recipes)
Sodium chloride, 5 M (see Recipes)
APS, 10% (w/v) (see Recipes)
Monomer solution (see Recipes)
Gelling solution (see Recipes)
Anchoring solution (see Recipes)
Digestion solution (see Recipes)
Recipes
1× PBS
Reagent Final concentration Quantity or Volume
10× PBS 1/10 10 mL
dH2O n/a 90 mL
Total n/a 100 mL
Permeabilization buffer A
Reagent Final concentration Quantity or Volume
1× PBS n/a 50 mL
BSA*
Triton X-100
0.5% (w/v)
0.5% (v/v)
0.25 g
250 µL
Total n/a 50 mL
*Prepare fresh. The prepared solution can be stored at 4 °C and used for two weeks.
Permeabilization buffer B*
Reagent Final concentration Quantity or Volume
1× PBS n/a 50 mL
BSA** 0.5% (w/v) 0.25 g
Triton X-100 0.2% (v/v) 100 µL
Total n/a 50 mL
*Permeabilization buffer B contains a reduced concentration of Triton X-100 for a gentler permeabilization during iteration steps 2–4.
**Prepare fresh. The prepared solution can be stored at 4 °C and used for two weeks.
Antibody diluent containing blocking agent
Reagent Final concentration Quantity or Volume
1× PBS n/a 50 mL
BSA 3% (w/v) 1.5 g
Total n/a 50 mL
Tris-HCl, 50 mM, pH 8
Reagent Final concentration Quantity or Volume
Tris base 50 mM 0.606 g
dH2O n/a 60 mL
Adjust to pH 8.0 with HCl
dH2O to 100 mL
Total n/a 100 mL
Sodium chloride, 5 M
Reagent Final concentration Quantity or Volume
Sodium chloride 5 M (0.292 g/mL) 2.922 g
dH2O to 10 mL
Total n/a 10 mL
APS, 10% (w/v)
Reagent Final concentration Quantity or Volume
APS powder* 10% (w/v) 1 g
dH2O to 10 mL
Total n/a 10 mL
*Store at room temperature (RT) in a desiccator.
Monomer solution*
Reagent (stock concentration) Final concentration Volume
Sodium acrylate (0.38 g/mL)** 0.086 g/mL 2.25 mL
Acrylamide (0.40 g/mL, 40%) 0.025 g/mL (2.5%) 0.58 mL
N,N'-Methylenebisacrylamide (0.02 g/mL, 2%) 0.0015 g/mL (0.15%) 0.75 mL
Sodium chloride (0.292 g/mL, Recipe 6) 0.117 g/mL 4 mL
10× PBS 1× 1 mL
dH2O n/a 0.82 mL
Total n/a 9.4 mL***
*Can be stored at -20 °C for up to 2 months.
**Sodium acrylate may have a variable purity affecting performance. Ensure it appears colorless under room light. If the reagent appears yellow, discard the stock without using it. Store at -20 °C in a desiccator.
***9.4/10 mL (1.06×), the remaining 6% of the volume will be brought up by the initiator, accelerator, and inhibitor as needed (see below).
Gelling solution*
Reagent Volume
Monomer solution** 188 µL
dH2O (or inhibitor 4-Hydroxy-TEMPO)*** 4 µL
TEMED**** 4 µL
APS (10%, w/v) (Recipe 7) 4 µL
Total 200 µL
*Prepare fresh and pipette reagents in the given order on ice.
**Degas the monomer solution after preparation by using a degassing chamber to remove any air bubbles. Air will impair the homogenous polymerization of the gel.
***OPTIONAL replacement of water. 4-Hydroxy-TEMPO solution appears yellow. It slows down the gelling, giving a wider temporal working window.
**** N,N,N',N'-Tetramethylethylenediamine. Store at RT in a desiccator.
Anchoring solution
Reagent (stock concentration) Volume
Acryloyl X – SE (10 mg/mL) 20 µL
PBS (1×) 1980 µL
Total 2,000 µL (2 mL)
Prepare fresh.
Digestion solution
Reagent (stock concentration) Final concentration Volume
Tris-HCl pH 8.0 (50 mM, Recipe 5) 50 mM 1,099 µL
EDTA (10 mM) 1 mM 140 µL
Triton X-100 0.5% (v/v) 7 µL
Guanidine HCl (8 M) 0.8 M 140 µL
Add before use:
Proteinase K 1:100 (8 units/mL) 14 µL
Total n/a 1.4 mL
Laboratory supplies
High-performance cover glasses 22 × 22 mm (D = 0.17 mm) (Carl Zeiss, catalog number: 474030-9020-000)
High-performance cover glasses 18 × 18 mm (D = 0.17 mm) (Carl Zeiss, catalog number: 474030-9000-000)
Scalpels
Spoons or spoon spatulas (stainless steel)
Scissors
Tweezers
Pasteur pipettes
Caliper
Hex nut, 9 mm in diameter
Circular glass coverslip, 1.5 mm thickness, 13 mm in diameter (Marienfeld, VWR, catalog number: MARI0117530)
Cell culture Petri dishes, non-treated, 35 mm in diameter
Cell culture Petri dishes, non-treated, 60 mm in diameter
Cell culture 6-well plates
Parafilm cut to 18 mm × 18 mm pieces to coat cover glasses of similar size
Cyanoacrylate super glue for gluing parafilm to cover glasses
In-house 3D-printed spacers with 350 µm thickness [acrylonitrile butadiene styrene (ABS) plastic], see specification of design for 3D printing from Mäntylä et al. [1]
In-house 3D-printed molds for cutting the gels [polylactic acid (PLA) plastic], see specification of design for 3D printing from Mäntylä et al. [1]
Coverslip cell chamber (Aireka Cells, catalog number: SC15022)
1.5 mL Eppendorf tubes
2.0 mL Eppendorf tubes
P1000 pipette tips
P200 pipette tips
P10 pipette tips
Polystyrene box (size L30 cm × H30 cm × W30 cm)
Tinfoil
Ice
Equipment
Vacuum degassing chamber (e.g., Applied Vacuum Engineering, model: DP3, 3 L)
Vacuum pump
Microwave
Inverted laser scanning confocal fluorescence microscope (e.g., Nikon Instruments, model: Nikon A1R in Ti2 Eclipse)
Software and datasets
Fiji/ImageJ image processing package (open source), Fiji (ImageJ, https://imagej.net/software/fiji/) can be used to analyze acquired microscopy images [9])
Noise estimation, denoising, and deblurring (NEDD) software (open source), MATLAB (v1.0.1, release date Apr 17, 2023). The code has been deposited to GitHub: https://github.com/lucioazzari/NoiseEstimationDenoisingDeblurring
Procedure
Iterative indirect immunostaining of samples
Grow cells to the desired confluency on a 22 × 22 glass coverslip placed in a 6-well plate with 2 mL of growth medium per well.
Note: Prior to culturing the cells, sterilize coverslips by rinsing three times with dH2O and three times in absolute EtOH and air dry for 20 min under UV light inside a laminar hood. Follow cell line–specific instructions in seeding and culturing the cells. If required, coverslips can be protein-coated to enhance cell attachment to the coverslip.
To fix the sample, add 20% PFA to a final concentration of 4% directly into the medium (400 µL in 2 mL of medium) in a laboratory fume hood. Tilt for an even spread and incubate for 10 min at RT. After fixation, discard the PFA-containing medium and wash the sample two times with 2 mL of 1× PBS (with Ca2+/Mg2+). Leave in 2 mL of 1× PBS.
Proceed to iterative indirect immunostaining (IT-IF) by performing the first iteration:
Note: The first iteration corresponds to traditional immunostaining.
Permeabilize the sample for 10 min at RT with 1 mL of the permeabilization buffer A (see Recipes).
Dilute the primary antibody/antibodies (Ab/Abs) of choice according to the manufacturer’s instructions in the antibody dilution solution containing 3% BSA as the blocking reagent. Use the recommended and optimized concentrations.
Note: Careful optimization of the antibody concentration/dilution is required before proceeding to iterative immunostaining.
Transfer the sample-containing coverslip to a clear 6-well plate/35 mm dish for immunostaining by using tweezers. Carefully dip dry the sample on paper to remove excess buffer before adding the primary Ab to avoid uneven spread and dilution of the primary Ab.
Add 100 µL of the primary Ab solution to the coverslip and incubate for 1 h at RT.
After primary Ab incubation, wash once with permeabilization buffer A, once with PBS, and again with the permeabilization buffer A (each wash with 1 mL for 10 min at RT). Use tweezers to dip dry samples into paper and move them to a clean 6-well plate/35 mm dish for the next staining step.
Prepare for the secondary staining by diluting the species-specific and cross-reacted Alexa Fluor 488 -conjugated goat anti-mouse secondary Ab 1:200 to the Antibody diluent solution containing 3% BSA. Protect from light.
For secondary staining, add 100 µL of the secondary Ab solution to the coverslip and incubate for 1 h at RT in the dark.
After secondary Ab incubation, wash the sample once with the permeabilization buffer A for 10 min, with 1× PBS for 10 min, and again with permeabilization buffer A for 10 min at RT (1 mL each) in the dark.
For iterating the staining, repeat primary and secondary immunostaining as described in steps A3b–i four times (four-time iteration) using the same Ab concentration.
Note: Use permeabilization buffer B (see Recipes) for all the subsequent permeabilization steps in iterations 2–4. Permeabilization buffer B contains a reduced concentration of Triton X-100 (0.2%).
After the washes following iteration 4, wash the sample once in PBS and once in dH2O (10 min at RT in the dark). Proceed to anchoring the preceding polymer gel casting.
For an alternative post-gelling immunostaining technique, see General note 1.
Anchoring treatment
Prepare the anchoring solution (see Recipes).
Place the sample on a fresh 6-well plate.
Pipette approximately 150–200 µL of the anchoring solution on the coverslip containing the cell sample.
Incubate the sample for > 6 h at RT.
Note: This reaction can be left overnight.
Wash 2 × 15 min with 2 mL of 1× PBS before proceeding to gelation. Samples can be stored at 4 °C in PBS in the dark before proceeding with the protocol.
Gelling
Note: Tools used in this section are presented in Figure 1.
Cut parafilm to a slightly larger size than the glass coverslip (18 mm × 18 mm) and do not remove the covering paper. Add one drop of glue to the coverslip and stick the parafilm to the glue with its paper-covered side facing up. Press gently and let dry. Since the gel does not adhere to parafilm, the parafilm-mounted coverslip will be used as the bottom of the gelation chamber. See Figure 1A showing the parafilm-mounted coverslip upside down to demonstrate the respective sizes of the parafilm and the coverslips after gluing.
Note: The parafilm prevents the gels from sticking to the top glass slide.
With the covering paper facing up, place the parafilm-mounted coverslip on a 6-well plate lid/60 mm Petri dish and remove the covering paper to expose the clean parafilm surface on the coverslip (Figure 1B).
Place the in-house 3D-printed spacer (Figure 1C) on the exposed parafilm surface of the coverslip (Figure 1D).
Note: This spacer controls the circular shape, size (opening r = 5 mm), and height (350 µm) of the polymer gel in the gelation phase. The size of the polymerized gel is required for calculating the expansion factor after the expansion. By using the in-house printed 3D spacers, the size of the gel is always constant and equals the size of the spacer opening. However, the size of the gel can also be measured using a caliper.
Cool a new 2 mL Eppendorf tube on ice.
Prepare Gelling solution (see Recipes) in the 2 mL Eppendorf placed on ice.
Note: Do not add initiator (APS) yet! One can prepare two samples from one gelling solution aliquot at the same time.
Once ready with the samples, add 10% APS (Recipe 7) to the gelation solution, vortex briefly, and immediately pipette 70 µL of gelation solution into the middle of the opening of the 3D-printed spacer placed on the parafilm-mounted coverslip (Figure 1E). The spacer controls the circular shape, size (opening r = 5 mm), and height (350 µm) of the polymer gel in the gelation phase.
Note: Pipetted volume depends on the sample size, so you might need to scale this if, e.g., tissue samples are being used.
Carefully but rapidly place the sample coverslip containing the immunolabeled cells on top of the gelling solution droplet with cells facing down, i.e., toward the droplet of gelling solution. The cells will be inside the gel after polymerization.
Note: Remove any extra PBS from the sample coverslip by touching it with tissue paper. This prevents PBS from diluting the gel solution.
Add a metal hex nut on top of the sample (Figure 1F).
Protect the sample from light by covering them, e.g., with tinfoil, and let the gel polymerize for 30–45 min at RT.
After polymerization, carefully remove the hex nut and flip the sample (sandwiched coverslips containing the spacer and the polymerized gel in between) so that after flipping, the parafilm-mounted coverslip is on top and the sample-containing coverslip is at the bottom. Remove the parafilm-mounted coverslip by gripping the parafilm edge with tweezers. Discard the parafilm-mounted coverslip. Remove the spacer. A circular gel remains on the sample coverslip with cells located at the bottom (facing the sample coverslip). If needed, use a scalpel to cut and remove any excess gel.
Note: The expansion will increase the gel size by approximately 4×, thus gels over 1 cm will be difficult to handle.
Place the gel (still attached to the sample coverslip) into a fresh well of a 6-well plate with the gel facing up.
The diameter of the polymerized gel is required for calculating the expansion factor after the expansion. By using the in-house-printed 3D spacers with known dimensions, the diameter of the gel is always constant and equals the diameter of the spacer opening (d = 10 mm). However, the diameter or some other size-related parameter of the gel can also be measured after the polymerization by using a caliper, especially if a spacer has not been used for casting the gel (Figure 1L). The expansion factor (equation described in section E) is required for pixel size quantification of the microscopy images.
Note: The gel will start to swell during the next digestion step; thus, it is crucial to measure the original gel size before that step.
Figure 1. Tools used in gel casting, mounting for imaging, and measuring the expanded gel. A. Parafilm piece glued to an 18 mm × 18 mm glass coverslip with super glue (turned upside down to demonstrate respective dimensions of the cut parafilm and the coverslip). B. Parafilm-coated coverslip in a Petri dish with the cover removed. C. In-house 3D-printed plastic gel casting spacer. D. Gel casting spacer placed on a parafilm-coated 18 mm × 18 mm glass coverslip. E. Drop of gelling solution in a gel casting spacer placed on a parafilm-coated 18 × 18 glass coverslip. F. Metal hex nut placed on top of the cell-containing 22 × 22 coverslip placed upside down into the gelling solution pipetted to a gel casting spacer on an 18 × 18 coverslip shown in E. G. In-house 3D-printed plastic gel cutting mold. H. Coverslip cell chamber, unassembled. I. Assembled coverslip cell chamber with a 22 × 22 glass coverslip on the bottom. J. Gel cutting mold containing a gel placed into the coverslip cell chamber containing a 22 × 22 coverslip. K. Sample preparation is finished by adding water on top of the gel to prevent drying and placing another 22 × 22 coverslip with a metal hex nut on top as a weight to keep the assembly in place. Arrows represent places where molten high-melt agar should be pipetted to mount the assembly prior to imaging. L. An example of a digital caliper used to measure the PAA-hydrogel after isotropic expansion in water to determine the expansion factor before proceeding to the gel cutting and mounting to the coverslip cell chamber.
Digestion
Prepare the digestion solution (see Recipes).
Digest the polymerized gel containing the cell sample on a 60 mm Petri dish with the digesting solution by adding at least 10-fold excess volume of digestion buffer (e.g., ~2 mL) overnight (12 h) in the dark at RT.
During the digestion, the gel with the cells inside will detach from the sample coverslip. The coverslip can remain in the Petri dish, but it can also be discarded.
Note: Do not peel the gels off the surface of the glass as this will damage the sample!
After digestion, move the polymerized gel by using a spoon/spatula to a fresh 60 mm Petri dish.
Note: If the coverslip is already detached, you can remove it from under the sample and discard it.
Expansion
Expand the gel sample isotropically by adding an excess volume of dH2O (e.g., 20 mL/60 mm dish). Incubate for 1 h in the dark at RT.
Note: Use at least 10× the final gel volume.
Discard water carefully and replace it with fresh water.
Notes:
The gel has the same refractive index as water. Use caution when removing the water. Avoid vacuum aspiration.
Gel expansion reaches a plateau after the third or fourth wash.
Repeat steps E1–2 at least five times.
After removing the water for the last time, measure the diameter (or other size-related parameters as described in step C12) of the expanded gel with a caliper (Figure 1L).
Calculate the expansion factor using the following equation.
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Note: By casting the gel in the opening (r = 5 mm, height 350 µm) of the in-house printed 3D spacer, a circular gel with known dimensions will be produced. The diameter of the spacer opening can be used as the pre-expansion gel diameter. After the expansion of the polymerized gel, measure the expanded gel diameter and use that as a post-expansion gel diameter to calculate the expansion factor. For traditional ExM, the expansion factor is near 4. Use this factor to determine the real size of your objects, for example during image analysis as described below in Data analysis section.
Sample mounting and microscopy
Note: The tools used in this section are presented in Figure 1.
After the isotropic expansion in water, cut a circular piece of the expanded gel with a 3D-printed mold (Figure 1G). Put the gel with the mold still attached on a new 22 mm × 22 mm coverslip. Place it inside the Aireka coverslip cell chamber (Figure 1H–J).
Place a 13 mm diameter glass coverslip on top of the gel and add a 9 mm hex nut to the top to keep the gel in place during mounting in the next step (Figure 1G–K).
Mount the gel-containing mold in the coverslip cell chamber with melted 1% high-melt agarose. Caution: Hot agarose is a burn hazard!
Note: A volume of 50 mL of high-melt agarose can be melted in a microwave in a glass bottle with a cap only loosely placed on top. Use 800 W power in 30 s intervals and swirl the bottle gently between the intervals. Avoid bubbles. Repeat until completely liquid and clear. Avoid overheating or boiling the agarose in the microwave. Let cool down at RT for 5–10 min. Add slightly cooled melted agarose (≤ 1 mL) to the empty sides between the mold and the chamber (not inside the mold/on top of the sample!) by pipetting with a Pasteur pipette. The agarose should not enter under the gel.
Add dH2O on top of the sample inside the mold to prevent the sample from drying and shrinking. The coverslip and the nut can be left in place and kept there during imaging to ensure that the sample stays in place to avoid drift.
The Aireka coverslip cell chamber fits in a regular 35 mm dish sample holder on a confocal microscopy stage. After placing it, let the temperature stabilize before the imaging. After 15–30 min, acquire images according to your needs.
Note: Imaging can be done according to the analysis requirements.
For imaging, use a water immersion (WI) objective with a high working distance and numerical aperture (N.A.) such as Nikon CFI Plan Apo IR SR 60× WI, N.A. 1.27, with a working distance of 0.17 mm. To acquire images for quantitative analysis, record an optical z-section series of 1,024 × 1,024 pixels (or similar) with a pixel size ≤ 50–70 nm in the x- and y-directions and ≤ 200 nm in the z-direction for optimal resolution and pixel size for mathematical deconvolution or signal processing.
NEDD denoising
Before analyzing the data, our open-access signal processing software NEDD (MATLAB) is recommended to further improve the signal-to-noise ratio of the images acquired from samples prepared with the above protocol for ExM. Our NEDD pipeline in MATLAB assumes the noise to be additive, spatially correlated, and with signal-dependent variance [10].
Note: The NEDD can process any LSCM image data and is not restricted to ExM.
The pseudo-algorithm of the NEDD pipeline can be summarized as follows:
Estimate the noise variance and power spectral density (PSD) of the data.
Apply a variance stabilizing transform (VST) to the data using the noise parameters estimated in the previous step. We use the generalized Anscombe transform [11].
Denoise the stabilized data using the iterative framework introduced in [12]. The denoising filter used within the framework is the RF3D algorithm designed for processing videos (three-dimensional data) corrupted by correlated noise [13].
Apply the inverse VST to return the data to its original intensity domain.
OPTIONAL: Perform deconvolution (deblurring) of the denoised sequence using a regularized Tikhonov deconvolution approach [14].
The software comes with a demo (demo.m), that can be run to demonstrate its usage, which can be called with the command:
noisy,denoised,deblurred]=processData(inputPath,outputPath,noiseModel,optionalParams)
where the inputs are:
inputPath: A string containing the path to the input noisy file. The file must be in single-channel gray-scale multi-page TIFF format.
outputPath: A string containing the path where to store the single-channel gray-scale multi-page TIFF output.
noiseModel: A string that defines the noise model to be assumed for estimation and denoising. It can be either white or colored.
optionalParams: A MATLAB struct variable that contains optional parameters that can be used to adjust, e.g., the denoising filter and the deblurring parameters. More details can be found in the function’s help.
The function will write the processed data into the outputPath. Additionally, the function returns to the workspace:
noisy: The noisy data (M × N × F double array) read from inputPath.
denoised: The denoised data (M × N × F double array).
deblurred: The deblurred data (M × N × F double array)
Data analysis
The application of LSCM in the imaging of ExM gels enables detailed quantitative image analyses, although the resolution is limited by diffraction. The traditional expansion leads to 4–6× enlargement of the samples, enabling visualization beyond the traditional diffraction limit of light microscopy. When performing image analysis or visualization, remember to consider the expansion factor in the pixel size or in the scale bar.
Note: To acquire the correct image scale, divide the voxel dimensions with the expansion factor.
The watery expansion of the gel will induce isotropic expansion, where the sample enlarges in all three dimensions. Thus, a cell nucleus of 20 µm in x/y diameter and 5 µm in z will appear a lot larger, approximately 40–120 µm in x/y and 20–30 µm in vertical direction. This results in long imaging times, where the sample needs to stay in place. However, if any movement occurs during imaging, it can be mathematically corrected in analysis programs. Moreover, the long imaging times might result in bleaching of the sample, as high laser powers are traditionally needed in ExM imaging. This is because the expansion dilutes the antibody-conjugated fluorophores in the sample, reducing the resulting image brightness, where the dilution scales to the third power of the (linear) expansion factor, and thus even 4× expansion can lead to a 64× reduction of the fluorescent signal. The IT-IF technique overcomes this traditional challenge in ExM and induces higher labeling density, leading to significantly improved signal intensity without compromising the SBR, advancing super-resolution imaging of highly compacted and proteinaceous intranuclear structures.
For the quantitative analysis, ensure you have at least three biological replicate samples, and image ~10–20 cells each, depending on the study. Also, due to the sensitivity of the technique and to reduce the statistical variance between the samples, always use predetermined and identical volumes when adding antibodies, washing, or otherwise treating the samples. Any errors or exceptions in the treatment protocol are likely to affect the intensity and SBR of the sample and cause differences between technical replicates of the sample.
To perform analyses, any open-source and free image analysis software such as Fiji/ImageJ [8] can be used. In analyzing the data, you can, for example, measure the total intensity of the sample by first making a maximum intensity z-projection, intensity thresholding and measurement of the region of interest, and normalizing it by subtracting the background intensity measured outside your target.
In Figure 2, we showcase the power of the presented pro-ExM protocol and the analysis of our published NEDD-denoised IT-IF-ExM-LSCM data generated using this protocol for visualization of lamin A/C organization in epithelial cells by using traditional (1×) immunostaining and IT-IF with four iterations (from first to fourth iteration) [1].
The results show how IT-IF combined with ExM-LSCM and NEDD denoising substantially improve the visualization of lamin A/C organization (Figure 2A, 2B, and 2D). We found that the IT-IF pro-ExM protocol results in significantly higher intensity and SBR of the lamin A/C (Figure 2C), and NEDD-denoising further improves the detection of lamin organization in high resolution (Figure 2A, 2B and 2D). Thus, our IT-IF and NEDD protocol provides a significant improvement to the detection and quantification of the structural organization of nuclear lamina in ExM.
Figure 2. Results of nuclear lamin A/C detection in noise estimation, denoising, and deblurring (NEDD)-denoised expansion microscopy–laser scanning confocal microscopy (ExM-LSCM) images following either traditional (1×) or iterative indirect immunostaining (IT-IF). A. Representative grayscale maximum projection LSCM images of nuclear lamin A/C of epithelial Madine Darby canine kidney type II (MDCKII) cells after the first iteration of corresponding traditional (1×) immunostaining (upper panels) and 4× iterated (fourth iteration) indirect immunostaining (IT-IF, lower panels) before (left panels) and after signal denoising (NEDD, right panels). Scale bars, 10 µm. B. Blow-out images showing details of lamin A/C following first and fourth iterations and NEDD denoising. Scale bars, 1 µm. C. Quantification of the fluorescence intensity (left y-axis) and signal-to-background ratio (SBR) (right y-axis) of nuclear lamin A/C and the background after first and fourth iterations. Error bars represent the standard deviation of the mean. One-way ANOVA was used to test for statistical significance. D. Collective blow-up images of the NEDD-denoised lamin A/C IT-IF data after first (upper panel) and fourth iteration (lower panel) from a region of interest indicated by a white box in B for direct comparison. Scale bars, 1 µm.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Iterative immunostaining and denoising protocol:
Mäntylä et al. [1]. Iterative immunostaining combined with expansion microscopy and image processing reveals nanoscopic network organization of nuclear lamina. Molecular Biology of the Cell (Figure 2, panels G–I; Figures 3–5).
Protein retention expansion microscopy protocol:
Tillberg et al. [15]. Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies. Nature Biotechnology (Figures 1–3).
General notes and troubleshooting
General notes
The iterations included in the protocol can be done subsequently without pauses or by dividing the work for four subsequent days (one iteration per day). For the latter, store the samples in 1× PBS at 4 °C overnight in between the iterations. This might be beneficial to enhance washing.
Immunostaining can also be done post-gelling. However, consider that this will require higher reagent volumes and concentrations of antibody solutions and longer incubation times as the diffusion of subjects across the gel takes time.
In a traditional proExM protocol, 4× concentrations of the primary Ab and 2× concentrations of the secondary Abs in comparison to normal immunofluorescence staining are used. Here, carefully optimized 1× concentrations apply.
Troubleshooting
Problem 1: Gel does not polymerize.
Possible causes: APS or TEMED is not working, or sodium acrylate solution might be expired or not pure.
Solutions: Prepare fresh 10% (w/v) APS. 10% (w/v) APS should be aliquoted and stored in a desiccator at -20 °C, as APS is hygroscopic. TEMED is also highly hygroscopic and accumulated water leads to its decomposition. You can also use parafilm to cover the aliquoted tube caps to prevent moisture from getting into the tube.
Problem 2: Gel is not attaching to the cells and/or part of the sample remains attached to the coverslip.
Possible cause: Crosslinking by using the anchoring solution is incomplete.
Solutions: Extend the anchoring time or prepare a fresh anchoring solution. Acryloyl-X is a succinimidyl ester-based linker. In anchoring, PFA-fixed and immunostained molecules undergo amine-acryloyl conversion to enable their covalent anchoring to the polymeric gel. Make sure that you use only clear Acryloyl-X solutions. Pure sodium acrylate solution is clear, whereas contaminated solution will have a yellow tint. Store Acryloyl-X as undiluted aliquots. Succinimidyl ester decomposes rapidly in water, especially in pH > 7; thus, use dry and fresh DMSO to prepare the stock solutions (DMSO is hygroscopic).
Problem 3: Gel is polymerizing too quickly, e.g., during pipetting.
Possible causes: Too high accelerator (TEMED) and/or initiator (APS) concentrations.
Solutions: Work in a laboratory with a less moist atmosphere. Prepare gelling solutions with less TEMED/APS. Make sure you keep all the solutions and tubes on ice. Add APS only before pipetting to the sample. If the gel is still polymerizing too quickly, replace water with 4-Hydroxy-TEMPO. This will slow down the reaction but not prevent it.
Problem 4: Only parts of the gel expand during expansion in water.
Possible cause: Uneven gelling solution following inadequate mixing.
Solution: Mix your gelling solution thoroughly before adding APS.
Problem 5: Gel drifts during LSCM imaging.
Possible causes: The sample is not mounted properly, or the temperature is higher during imaging causing movement of the agar used for mounting.
Solution: Apply only high-melt agar for mounting the samples. Keep samples under a weight (glass coverslip and a hex nut) during imaging. Check the temperature of the microscopy room/incubator. Imaging should be done at RT and in stable conditions. Let the sample stabilize on the microscopy stage for at least 15 min before imaging.
Problem 6: Antibody background is too high after iterations.
Possible causes: Poor quality of secondary antibodies or off-target binding of the primary antibody.
Solution: Carefully optimize the antibody concentration before iterations. Iteration works for most of the antibodies. However, polyclonality might present challenges. Try another and prefer monoclonal antibodies.
Acknowledgments
This work was supported by the Research Council of Finland under the award numbers 308315 and 314106 (T.O.I.), 332615 (E.M.), and 336357 (PROFI6—TAU Imaging Research Platform [L.A., M.V.]. The authors acknowledge Biocenter Finland, and Tampere University Tampere Imaging Facility for the services. This IT-IF-ExM protocol was originally described and validated in Mäntylä et al. [1] Molecular Biology of Cell (2023), doi: 10.1091/mbc.E22-09-0448.
Graphical overview was created with BioRender.com.
Competing interests
The authors declare no competing interests.
Ethical considerations
This work did not use human or animal subjects and has no ethical considerations.
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Azzari, L. and Foi, A. (2014). Indirect Estimation of Signal-Dependent Noise With Nonadaptive Heterogeneous Samples. IEEE Trans Image Process. 23(8): 3459–3467.
Maggioni, M., Sanchez-Monge, E. and Foi, A. (2014). Joint Removal of Random and Fixed-Pattern Noise Through Spatiotemporal Video Filtering. IEEE Trans Image Process. 23(10): 4282–4296.
Dabov, K., Foi, A., Katkovnik, V. and Egiazarian, K. (2008). Image restoration by sparse 3D transform-domain collaborative filtering. SPIE Proceedings: e766355.
Tillberg, P. W., Chen, F., Piatkevich, K. D., Zhao, Y., Yu, C. C., English, B. P., Gao, L., Martorell, A., Suk, H. J., Yoshida, F., et al. (2016). Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies. Nat Biotechnol. 34(9): 987–992.
Article Information
Publication history
Received: Jun 14, 2024
Accepted: Aug 8, 2024
Available online: Aug 30, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cell Biology > Cell imaging > Super resolution imaging
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Pharyngeal Pumping Assay for Quantifying Feeding Behavior in Caenorhabditis elegans
MS Muniesh Muthaiyan Shanmugam
Pankaj Kapahi
Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5073 Views: 401
Reviewed by: Chiara AmbrogioRupkatha BanerjeeIstvan Stadler
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Original Research Article:
The authors used this protocol in eLIFE Sep 2023
Abstract
C. elegans is a well-established nematode model organism, with 83% of its genes conserved in humans with translation potential. C. elegans is translucent, with clearly defined cellular organization, and robustly identifiable under a microscope, being an excellent model for studying feeding behavior. Its neuromuscular pharyngeal pump undergoes a pumping motion that can be quantified to study feeding behavior at specific treatment conditions and in genetically modified worms. Understanding the evolutionarily conserved feeding behaviors and regulatory signals is vital, as unhealthy eating habits increase the risk of associated diseases. The current protocol was developed to identify and study evolutionary conserved signals regulating feeding behavior. The protocol described here is very robust in calculating the pumping rate (pumping per minute) as it directly counts the pharyngeal pumping for 30 s. This protocol uses basic laboratory instrumentation, such as a stereomicroscope with an attached camera and a computer with a video program that can be used to count manually. The advantages of studying C. elegans feeding include understanding the genetic basis of feeding regulation, dysregulation of feeding behavior in a disease model, the influence of toxic or environmental substances in feeding behavior, and modulation of feeding behavior by pharmacological agents.
Key features
• Quantifies pharyngeal pumping, which can be used to study up/downstream signaling in feeding regulation.
• Uses a phenotype (pharyngeal pumping) that is easy to score.
• Requires only a stereomicroscope with a camera to record the pharyngeal pumping, which can be counted manually.
Keywords: C. elegans Pharyngeal pumping Feeding Worms Microscopy Stereomicroscope Eating behaviors
Background
Feeding is a crucial and complex behavior that ensures the intake of nutrition for survival and healthy living across animal species [1]. With the recent availability of calorie-dense foods and an increase in unhealthy eating habits [2], it has become necessary to understand feeding behavior and the mechanisms that thoroughly regulate it. Discovery of the mechanisms regulating feeding behavior in primitive model organisms offers a faster understanding with translation potential to humans. Caenorhabditis elegans is a tiny translucent nematode worm, with only 1 mm in length, and a robust feeding behavior that is easy to quantify. These worms have a short life cycle (duration for the larval worms to reach the reproductive age, or young adults) of 3.5 days at 20 °C and a lifespan of approximately 30 days, making them cost-effective to maintain in a laboratory [3]. With an 83% gene homology to humans and a high translation potential, C. elegans is an excellent model organism for genetic manipulations to study various biological processes and discover previously unexplained signaling pathways [4]. The worms feed on bacteria through their pharynx, and their feeding rate depends on the neuromuscular pumping action of their pharyngeal muscles. The worm's intake of bacterial food is facilitated by two pharyngeal movements, namely pumps and isthmus peristalses. Pumping occurs due to cycles of coordinated contraction and relaxation of pharyngeal muscles, which can be measured to determine the feeding rate. Isthmus peristalsis movement involves the opening of the posterior lumen, followed by closure in an anterior-to-posterior wave [5,6].
Extensive understanding of C. elegans feeding behaviors exists, and with availability of food ad libitum, the pharyngeal pumping rate is the limiting step for food intake [5–9]. The method outlined here is robust because it directly counts the pharyngeal pumping movement using a manual counter (Figure 1). To do this, a low-magnification stereomicroscope with an attached camera can be used to record the pumping of pharyngeal muscles in an unrestrained C. elegans in vivo for further analysis. Since the pumping rate is very high and difficult to follow with the naked eye, video player software can be used to slow down the speed, making it possible to count the pumps manually using a clicker counter [10]. However, implementing this method can be laborious for collecting and analyzing large datasets or conducting high-throughput analysis. Recently, Bonnard et al. [11] developed an automated system to measure unrestrained worms' pharyngeal pumping and foraging behavior, which can be implemented for conducting high-throughput analysis and screening. Further, a microfluidic device was developed to measure the pumping rate on restrained worms in microfluidic channels for high-throughput analysis [12]. Other methods for quantifying feeding behavior include measuring the reduction in bacterial density in liquid culture (bacterial clearance assay) [13,14], intestinal fluorescence of GFP-labeled bacteria, and BODIPY dye [6]. Compared to the alternative methods mentioned above, direct counting of pharyngeal pumping requires less standardization with basic equipment that commonly exists in any C. elegans laboratory and is quicker if a large number of samples is not involved.
Figure 1. Step-by-step visual guide for quantifying pharyngeal pumping in C. elegans to study feeding behavior. Refer to the representative Video 1.
Materials and reagents
Biological materials
Required C. elegans strains (N2 wild-type and glod-4 mutant). The C. elegans strains used in this study were obtained from the Caenorhabditis Genetic Center (CGC) in Minneapolis, USA, and a few other/necessary strains can be obtained from the National Bioresource Project, Tokyo, Japan. Please refer to wormbook.com for basic worm culture techniques
Bacteria E. coli OP50-1. Obtained from Caenorhabditis Genetic Center (CGC), Minneapolis, USA
Reagents
Sodium chloride (Sigma-Aldrich, catalog number: S9888)
BactoTM peptone (BD Diagnostic Systems, catalog number: 211677)
Agar (BD DIFCOTM, catalog number: 214510)
Calcium chloride (Sigma-Aldrich, catalog number: C3881)
Magnesium sulfate (Sigma-Aldrich, catalog number: 208094)
Cholesterol (Sigma-Aldrich, catalog number: C8667)
Streptomycin sulfate (Sigma-Aldrich, catalog number: S1567)
Nystatin (Chem Impex, catalog number: 00816)
Potassium phosphate monobasic (Sigma-Aldrich, catalog number: P0662)
Potassium phosphate dibasic (Sigma-Aldrich, catalog number: P3786)
Luria-Bertani (LB) broth (BD DifcoTM, catalog number: 244610)
Serotonin hydrochloride (Sigma-Aldrich, catalog number: H9523)
Solutions
1 M calcium chloride (autoclave to sterilize)
1 M magnesium sulfate (autoclave to sterilize)
5 mg/mL cholesterol in ethanol
100 mg/mL streptomycin sulfate (filter with 0.2 μm filter to sterilize)
10 mg/mL nystatin in ethanol
Potassium phosphate buffer: Mix 868 mL of potassium phosphate monobasic (1 M) and 132 mL of potassium phosphate dibasic (1 M), pH 6 (autoclave to sterilize)
0.5 M serotonin hydrochloride stock
Note: Double-distilled or MilliQ water is used as solvent unless specified.
Nematode growth medium (NGM) plates (see Recipes)
LB broth (see Recipes)
OP50-1 culture (see Recipes)
Recipes
Nematode growth medium (NGM) plates
Prepare 3 g of sodium chloride, 2.5 g of peptone, and 25 g of agar in 970 mL of water, then autoclave the solution (15 psi pressure at 121 °C for 30–60 min) to sterilize it. Once the solution has reached a bearable temperature (bearable when the container touches the ventral side of the arm below the hand), add 25 mL of potassium phosphate buffer and 1 mL each of 1 M magnesium sulfate, 1 M calcium chloride, cholesterol, streptomycin, and nystatin in a laminar flow hood. After thorough mixing, pour 25 mL of the solution into 10 cm Petri plates and allow it to cool inside the laminar flow hood overnight and solidify. Spread 500 µL of 5× concentrated OP50-1 bacterial culture on the NGM plates and incubate at 37 °C for 16 h. After 16 h, store the plates at 4 °C until used. Although the plates can be stored at 4 °C for a month if properly stored in a sealed box with reduced perspiration, use the plates within a week.
LB broth
Dissolve 25 g of LB in 1 L of water in a 5 L conical flask and autoclave to sterilize the media. The autoclaved LB media can be stored at 4 °C for up to a month or until its sterility is compromised by outside contaminants.
OP50-1 culture
Inoculate a colony of the OP50-1 E. coli strain in LB broth (1 L of LB broth prepared as in Recipe 2) containing streptomycin sulfate (working concentration of 100 μg/mL) using a sterile inoculating loop. OP50-1 obtained from CGC should be cultured, maintained, and stored at -80 °C. Streak the OP50-1 in an LB agar plate to isolate a single colony (refer to basic microbiological technique for maintenance of bacterial strain). Incubate the culture at 37 °C at 250 rpm in a bacterial incubator (shaking the culture at 250 rpm improves oxygenation, resulting in better bacterial growth) for 16 h. After 16 h, concentrate the bacterial culture five times by centrifugation at 4,000× g for 10 min in a 50 mL tube; remove the necessary supernatant (40 mL). Vortex (use maximum speed setting in the vortexer) the pellet to disperse the bacterial pellet, which can then be used to spread on the NGM plates (500 μL of bacterial culture/NGM plate).
Laboratory supplies
Glass conical flasks (5 L)
Measuring cylinder (1,000 mL and 500 mL)
Spatula
Disposable pipette (50 mL, 25 mL, and 10 mL) and motorized pipette controller (Accuhelp, model: PH01-B)
Pipette tips and micropipette (Eppendorf)
10 cm disposable sterile plastic Petri plates (standard Petri dishes) (Celltreat® Scientific Products, catalog number: 229695)
Inoculating loop (UltraCruz®, model: sc-200265)
Equipment
Autoclave (any basic or benchtop autoclave that can reach 15 psi and 121 °C for 30 min)
Weighing balance (Ohaus, model: Pioneer PX163 analytical and precision balance)
Laminar flow hood
Stereomicroscope (Leica, model: M165 FC)
Computers for recording the pharyngeal pumping video and analysis (A basic computer with Windows operating system that runs Window Media Player or/and VLC media player)
Handheld manual counter clicker
Bacterial shaker incubator set to 37 °C and >250 rpm
Centrifuge (Eppendorf, model: 5810 R)
Overhead projector (OHP) sheets or multifunction transparency film (example: Optiazure, multifunction transparency film 8.5 × 11 inch)
WormStuff worm pick with platinum wire (Genesee Scientific, model: 59-AWP)
Worm incubator at 20 °C
Bacterial incubator at 37 °C (to grow OP50-1 on the NGM plates; Recipe 1)
Vortexer
Software and datasets
Leica microscope software LAS V4.12
Window media player
VLC media player
Microsoft Excel
GraphPad Prism (any version)
Procedure
Preparation of worms
Note: Worms are synchronized by timed egg-laying.
Transfer 35 young-adult hermaphrodite worms (per plate) to fresh NGM plates containing OP50-1 to lay embryos on the NGM plates. [Choose the number of plates based on the number of treatment groups for the experiment. Each group should have at least 30–40 worms (N = 30–40) for data collection.]
Incubate the plates at 20 °C for 30 min.
After 30 min, remove the adult worms from the plates. They can be transferred to another plate for further use or discarded.
Incubate the plates with embryos at 20 °C until the worms reach the young-adult stage (approximately 65 h from timed egg-laying).
For treatment, such as 5 mM serotonin, overlay the NGM plate containing OP50-1 bacteria (grown on 25 mL of solidified media) with the 250 μL of 500 mM serotonin stock solution. Rotate the NGM plate to ensure the stock solution spreads across the entire surface. Allow the spread stock solution to air dry in the laminar flow hood and diffuse into the agar to achieve the desired concentration.
A necessary number of young-adult worms (N = 30–40) are transferred to either a treatment plate or control plate without the drug and incubated for an appropriate amount of time [e.g., for N2 vs. glod-4 genetic mutant (Figure 2), capture videos of appropriately aged worms (no treatment necessary), 1 h incubation for serotonin (Figure 3) and 24 h for MG-H1 [10] (methylglyoxal-derived hydroimidazolone)].
Note: Basic worm maintenance techniques are found in WormBook in the chapter “Maintenance of C. elegans” [15].
Data collection
We used a Leica M165 FC stereomicroscope to capture video recordings of pharyngeal pumping using a 2× objective lens with brightfield illumination. Any stereomicroscope with a 2× objective lens and a camera to record movies can be used to record pharyngeal pumping videos.
To capture the worms' live activity, keep the exposure per frame very low, around 5 ms, and record movies for 40 s (Video 1).
Choose a healthy, actively moving worm on the bacterial lawn, and start recording the videos. Do not record the worms that were harmed while picking (worms that cannot move in a sinusoidal waveform, worms that are not moving, and worms that are severely injured, like being split open near the vulva) or while transferring during treatment (if necessary, use eyelash picks to pick and transfer the worms). Worms outside the bacterial lawn should not be considered for data collection and analysis.
Since the worms are unrestrained, they will move freely. Therefore, move the plate to keep the worm within the objective lens's field of view. To exert less stress on the hand while freely moving the plate to accommodate the worm in the field of view, place the Petri plates on the transparent plastic sheet (OHP sheets) and move the sheet accordingly to move the plate around (Figure 4).
The obtained pharyngeal pumping recording can be analyzed as described below.
Data analysis
Run the video obtained from the Leica M165 FC stereomicroscope on Windows Media Player Legacy software at 0.25 times the original speed (open the video → right-click → Enhancements → Play speed settings → reduce the speed to 0.25) and count manually using a handheld counter clicker until the video reaches 30 s of recording.
Enter counted values in either Microsoft Excel or GraphPad Prism for further statistical analysis. A Student t-test can be used to compare two experimental groups, and a one-way ANOVA can be used to compare more than two groups. In the examples provided with this protocol, we utilized a Student t-test to compare the experimental groups (Figures 2: N2 vs. glod-4, and Figure 3: untreated control vs. serotonin treatment). Please follow the instructions in the software manual to perform statistical analysis.
Validation of protocol
This protocol has been used and validated in the following research article: Muthaiyan Shanmugam et al. [10]. Methylglyoxal-derived hydroimidazolone, MG-H1, increases food intake by altering tyramine signaling via the GATA transcription factor ELT-3 in Caenorhabditis elegans—eLife (Figure 2). The data below compares the pharyngeal pumping of N2 wild-type worms with that of glod-4(gk189) mutant worms. The glyoxalase enzyme (glod-4) is responsible for detoxifying methylglyoxal (MGO), which non-enzymatically interacts with biomolecules to form advanced glycation end-products (AGEs). The absence of the glyoxalase enzyme leads to increased accumulation of AGEs. MG-H1, a type of MGO-derived AGE, has been shown to increase feeding behavior in our research.
Figure 2. Glyoxalase glod-4 mutant worms exhibit increased pharyngeal pumping. Quantification of pharyngeal pumping (number/30 s) in N2(wt) and glod-4(gk189) mutants at different stages of adulthood from day 1 of the young-adult stage (65 h after egg laying). Student t-test. ****p < 0.0001 and **p < 0.01. # represents number. Figure reprinted from Muthaiyan Shanmugam et al. [10]. The Creative Commons Attribution License permits unrestricted use of this article with proper acknowledgment to the authors and source.
Serotonin is a neurotransmitter that plays a vital role in regulating various behaviors in C. elegans, such as locomotion, pharyngeal pumping, or egg-laying [16]. We validated our method by quantifying pharyngeal pumping after treating day 1 young-adult N2 wild-type worms with 5 mM of serotonin for 1 h. The results demonstrate a significant increase in pharyngeal pumping following 1 hour of serotonin treatment (Figure 3).
Figure 3. Serotonin neurotransmitter increases pharyngeal pumping. Quantification of pharyngeal pumping after 5 mM serotonin treatment for 1 h on N2 wild-type worm (day 1 young-adult worms, 65 h after egg laying). Equal solvent volume (in this case, water, as serotonin hydrochloride is dissolved in water) was added over the control NGM plates containing OP50-1. Student t-test. ***p < 0.001. # represents number.
The literature consistently shows that the pharynx pumps at a rate of 200–300 pumps per minute when food is available [5,6,14]. Our calculated pumping rate, as shown in Figures 2 and 3, aligns with the rates reported in the literature, further validating our approach.
General notes and troubleshooting
The pharyngeal pumping rate varies with the worm's development [10] (Figure 2), so it is important to collect data from different treatment groups or genetic backgrounds at the exact developmental stage to compare them accurately. One way to achieve this is to synchronize the worm groups individually at a specific time rather than simultaneously, to ensure the exact developmental stage during data collection.
When designing data collection for large sample sizes or multiple test groups, it is important to consider the technician’s or researcher’s fatigue that may result from the technique used. It typically takes about a minute to locate an unrestrained worm, focus, and begin recording a 40 s pumping video, of which 30 s of video is available for manual counting to acquire data. Depending on the penetrance of the observed phenotype, it is necessary to collect data from at least 30 individual worms per group, which requires 40–50 min, depending on the fatigue experienced. Planning short resting periods during data collection between treatment groups is wise. Alternatively, when comparing multiple treatment or genetic groups, the video recording of worms from each group can be staggered. This means that 10 worms from each group are recorded, followed by worms from the other groups, and this cycle is repeated until the desired number of worms from each group has been recorded, thus minimizing any variations caused by the recording process.
To account for variations induced by lab temperature fluctuations, prepare three worm plates for each treatment group. Record data from only 10 worms per plate, while the other plates are incubated at 20 °C.
The video file generated by the Leica software LAS V4.12 is generally very large, which makes transferring raw data to another computer difficult and time-consuming. If necessary, the VLC media player can be used to record the desktop for 40 s when a worm is focused and projected using the Leica software LAS V4.12 in live mode (Video 1); while doing so, set 50 frames per second in the VLC media player option. This generates pumping videos with a smaller file size, which makes transferring files easier.
Video 1. Representative video recording of an N2 wild-type worm as described in the protocol
To ensure that the unrestrained worms on the NGM plate containing OP50-1 remain in the field of the objective lens, it is necessary to move the NGM plate constantly. However, constantly holding and moving the NGM plate can exert considerable stress on the hand. One way to reduce this stress is to place the NGM plate on a transparent plastic sheet, like an overhead projector (OHP) sheet, and slide the sheet as the worms move to keep them in the field of the objective lens (Figure 4).
Figure 4. Picture showing the use of a transparent plastic sheet to move the NGM plate without exerting stress on the hand by holding the plate
Acknowledgments
This work was supported by grants from NIH (R01AG061165 and R01AG068288) and the Larry L Hillblom Foundation (2021-A-007-FEL) to P.K. We would like to express our appreciation and acknowledge to Dr. Michael Petrascheck, The Scripps Research Institute, for his valuable insights that enhanced our research, and to Karishma Patel, The Buck Institute for Research on Aging, for her assistance in obtaining the image for Figure 4.
Competing interests
The authors declare no competing interests.
References
Wu, Q., Gao, Z. J., Yu, X. and Wang, P. (2022). Dietary regulation in health and disease. Signal Transduction Targeted Ther. 7(1). doi.org/10.1038/s41392-022-01104-w.
Imamura, F., Micha, R., Khatibzadeh, S., Fahimi, S., Shi, P., Powles, J. and Mozaffarian, D. (2015). Dietary quality among men and women in 187 countries in 1990 and 2010: a systematic assessment. Lancet Glob Health. 3(3): e132–e142.
Meneely, P. M., Dahlberg, C. L. and Rose, J. K. (2019). Working with Worms: Caenorhabditis elegans as a Model Organism. Curr Protoc Essent Lab Tech. 19(1): e35.
Lai, C. H., Chou, C. Y., Ch'ang, L. Y., Liu, C. S. and Lin, W. C. (2000). Identification of Novel Human Genes Evolutionarily Conserved in Caenorhabditis elegans by Comparative Proteomics. Genome Res. 10(5): 703–713.
Avery, L. and You, Y. J. (2012). C. elegans feeding. WormBook. doi/10.1895/wormbook.1.150.1.
David, R., Song, B., Trojanowski, N. and You, Y. J. (2012). Methods for measuring pharyngeal behaviors. WormBook. doi/10.1895/wormbook.1.154.1.
Fang-Yen, C., Avery, L. and Samuel, A. D. T. (2009). Two size-selective mechanisms specifically trap bacteria-sized food particles in Caenorhabditis elegans. Proc Natl Acad Sci USA. 106(47): 20093–20096.
Avery, L. and Shtonda, B. B. (2003). Food transport in the C. elegans pharynx. J Exp Biol. 206(14): 2441–2457.
Seymour, M. K., Wright, K. A. and Doncaster, C. C. (1983). The action of the anterior feeding apparatus of Caenorhabditis elegans (Nematoda: Rhabditida). J Zool. 201(4): 527–539.
Muthaiyan Shanmugam, M., Chaudhuri, J., Sellegounder, D., Sahu, A. K., Guha, S., Chamoli, M., Hodge, B., Bose, N., Roberts, C., Farrera, D. O., et al. (2023). Methylglyoxal-derived hydroimidazolone, MG-H1, increases food intake by altering tyramine signaling via the GATA transcription factor ELT-3 in Caenorhabditis elegans. eLife. 12: e82446.
Bonnard, E., Liu, J., Zjacic, N., Alvarez, L. and Scholz, M. (2022). Automatically tracking feeding behavior in populations of foraging C. elegans. eLife. 11: e77252.
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Article Information
Publication history
Received: Apr 1, 2024
Accepted: Jul 1, 2024
Available online: Aug 30, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Neuroscience > Behavioral neuroscience > Sensorimotor response
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A Full Good Manufacturing Practice–Compliant Protocol for Corneal Stromal Stem Cell Cultivation
MS Mithun Santra
YH Yen-Michael S. Hsu
SK Shaheen Khadem
SR Sydney Radencic
MF Martha L. Funderburgh
OS Onkar B. Sawant
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Published: Vol 14, Iss 18, Sep 20, 2024
DOI: 10.21769/BioProtoc.5074 Views: 428
Reviewed by: Alessandro DidonnaShun Yu Jasemine YangGabriel Campolina-Silva
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Cited by
Original Research Article:
The authors used this protocol in Stem Cell Research & Therapy Jan 2024
Abstract
Corneal scarring, a significant cause of global blindness, results from various insults, including trauma, infections, and genetic disorders. The conventional treatment to replace scarred corneal tissues includes partial or full-thickness corneal transplantation using healthy donor corneas. However, only 1 in 70 individuals with treatable corneal scarring can undergo surgery, due to the limited supply of transplantable donor tissue. Our research focuses on cell-based strategies, specifically ex vivo–expanded corneal stromal stem cells (CSSCs), to address corneal scarring. Preclinical studies have demonstrated the efficacy of CSSC treatment in reducing corneal inflammation and fibrosis, inhibiting scar formation, and regenerating native stromal tissue. Mechanisms include CSSC differentiation into stromal keratocytes and the expression of regenerative cytokines. Here, we present a good manufacturing practice (GMP)-compliant protocol to isolate and expand human CSSCs. This method paves the way to produce clinical-grade CSSCs for transplantation and clinical trials.
Key features
• This protocol utilizes surgical skills to dissect human corneal tissues for CSSC isolation.
• The yield and features of CSSCs rely on donor tissue quality (freshness) and have donor-to-donor variability.
• Up to 0.5 billion CSSCs can be generated from a single cornea specimen, and cells at passage 3 are suitable for treatment uses.
Keywords: Corneal opacities Corneal stromal stem cells Anterior limbal stroma Good manufacturing practices Standard operating protocol
Graphical overview
Background
Damage to the cornea, caused by physical trauma, chemical burns, infections such as microbial or viral keratitis, or genetic disorders like corneal dystrophies and keratoconus, initiates a complex cascade of inflammatory and fibrotic reactions [1,2]. These events lead to tissue damage and remodeling, which involves the generation of stromal fibroblasts and myofibroblasts. These cells produce a repair-type extracellular matrix (ECM) to heal the wound. However, excessive deposition of ECM, including type-III collagen and fibronectin extradomain A (EDA-Fn), leads to the formation of haze and opacities, eventually resulting in corneal scarring that can compromise the corneal clarity and integrity, causing visual impairment [3]. While superficial scarring can often heal within months, deep or dense scars may develop and cause permanent vision loss.
Corneal scarring is the fourth leading cause of global blindness, affecting approximately 2 million people [4]. Each year, over 350,000 children are affected by this condition [5,6]. This is a major concern, as it disproportionately affects younger individuals and low-income rural communities due to factors such as communicable diseases, higher risk of injury, and limited access to treatment options [7]. Various options are available to treat scarred corneas, including pharmacological treatments such as steroids, immune modulation, and grafting with donor corneal materials [8]. Pharmacological therapies offer moderate correction for corneal haze and mild opacities. Corneal transplantation, such as penetrating and lamellar keratoplasty, is the standard for moderate-to-severe scarring and opacities occurring along the visual axis. However, the limited global supply of transplantable donor corneas, immune response (depending on the histocompatibility context of the host that could affect the long-term graft survival), risk of graft rejection, and potential surgical complications restrict the widespread use of keratoplasty [7,9].
Our research team investigated the use of human corneal stromal stem cells (CSSCs) to reduce corneal inflammation and fibrosis; the treatment prevented corneal scarring and restored corneal clarity in pre-clinical models of stromal injury created by mechanical debridement, alkali burns, or liquid nitrogen [10–17]. Our team follows a specific set of protocols to isolate primary donor CSSCs through microdissection of the anterior limbal stroma and then culture them using a stem cell growth medium called JM-H [10,18,19]. These CSSCs can be propagated up to 50–70 doublings before undergoing senescence and changes in cell features [20]. They express stem cell markers like ATP-binding cassette transporter superfamily G member 2 [ABCG2], nestin, Pax6, and Bmi-1, undergo stromal keratocyte differentiation, express anti-inflammatory tumor necrosis factor-stimulating gene 6 [TSG-6] and transforming growth factor b3 [TGFb3], and anti-fibrosis microRNAs (hsa-miR-29a and 381) [10,12,14,17,18].
In order to apply these cells in clinical settings, it is necessary to generate them using a good manufacturing practice (GMP)-compliant protocol. Regulatory approval for cell therapy is challenging, with only a small number gaining approval in comparison to pharmaceutical drugs [21]. Quality, safety, and effectiveness are extremely important for clinical trials, guided by good clinical practice principles [22,23]. To ensure regulatory compliance and consistent results, a quality management system that encompasses quality assessment and quality control is crucial for cell therapy [23]. We have developed GMP-compliant protocols for clinical-grade CSSC production, which laid the groundwork for human testing in clinical trials.
Materials and reagents
Biological materials
Donor corneas
GMP Reagents
Dulbecco’s modified Eagle medium (DMEM/F12) (Thermo Fisher Scientific, Gibco, catalog number: 12634-010)
Phosphate buffered saline (PBS) (Thermo Fisher Scientific, Gibco, catalog number: 10010-023)
DMEM (1 g/L D-glucose, with GlutaMAX) (Thermo Fisher Scientific, Gibco, catalog number: 10567-014)
Ham’s F12 (Thermo Fisher Scientific, Gibco, catalog number: 31765-035)
BioWhittaker antibiotic-antimycotic (100×) (Lonza, catalog number: 17-745E)
Insulin-transferrin-selenium (100×) (Thermo Fisher Scientific, Gibco, catalog number: 41400-045)
Flexbumin (25%) (BioSupply, catalog number: 00944-0493-01)
2-Phospho-L-Ascorbate (Tocris Biosci, catalog number: 5778)
Recombinant human epidermal growth factor (EGF) (Thermo Fisher Scientific, Gibco, catalog number: PHG6045)
Recombinant human platelet-derived growth factor-BB (PDGF-BB) (R&D, catalog number: 220-GMP-050)
Dexamethasone (Dex) (USP micronized) (Spectrum, catalog number: DE121)
Human serum (Innovative Res, catalog number: ISER-36670)
Collagenase (Collagenase NB6) powder (Nordmark, catalog number: N0002779)
Fibronectin coating substrate (MedChemExpress, catalog number: HY-P70593G)
TrypLE Select CTS (Thermo Fisher Scientific, Gibco, catalog number: A12859-01)
CryoStor CS10 (serum-free DMSO 10%) (BioLife Solutions, catalog number: 210373)
Trypan blue (0.4%) (Invitrogen, catalog number: T10282)
Hydrochloric acid (HCl) (Sigma-Aldrich, catalog number: H9892)
Solutions
Recombinant human EGF (see Recipes)
Recombinant human PDGF (see Recipes)
Dexamethasone (Dex) solution (see Recipes)
Collagenase NB6 solution (see Recipes)
GMP fibronectin (see Recipes)
CryoStor CS5 (see Recipes)
GMP stem cell culture medium (see Recipes)
Recipes
Recombinant human EGF
Store the lyophilized EGF at -20 °C under desiccation until reconstitution. Prior to opening, briefly centrifuge the vial of lyophilized EGF at 400× g for 10–15 s to bring the contents to the bottom. Reconstitute EGF powders (500 mg) with sterile distilled water (1 mL volume) to a concentration of 0.5 mg/mL. Further dilute to 10 mg/mL with sterile DMEM/F12 added with 0.1% Flexbumin (carrier protein), aliquot at 0.5 mL, and store at -80 °C until use or at 2–8 °C for one month.
Reagent Final concentration Quantity
Recombinant human EGF 10 mg/mL 100 mL from 0.5 mg/mL stock
DMEM/F12 1× 5 mL
Flexbumin 0.1% (v/v) 20 mL from 25% stock
Recombinant human PDGF
Store the lyophilized PDGF-BB at -20 °C under desiccation until reconstitution. Prior to opening, briefly centrifuge the vial of lyophilized PDGF-BB at 400× g for 10–15 s to bring the contents to the bottom. Reconstitute PDGF-BB powders (50 mg) with sterile hydrochloric acid (HCl, 4 mM) with 0.1% Flexbumin (carrier protein) to a concentration of 10 mg/mL, aliquot at 0.5 mL, and store at -80 °C until use or at 2–8 °C for one month.
Reagent Final concentration Quantity
Recombinant human PDGF-BB 10 mg/mL One vial of 50 mg
HCl 4 mM 5 mL
Flexbumin 0.1% (v/v) 20 mL from 25% stock
Dexamethasone (Dex) solution
Store the lyophilized Dex at -20 °C under desiccation until reconstitution. Prior to chemical preparation, wear PPE and a suitable eye protection device as Dex has a risk of causing eye damage/irritation. Thaw Dex at room temperature for 10–20 min. Weigh approximately 50 mg of powder and calculate the amount of distilled water to make a stock concentration of 10 mM with reference to the molecular weight 392.47. Further dilute to 10 mM with sterile DMEM/F12, aliquot at 100 mL volume, and store at -80 °C until use or at 2–8 °C for one month.
Reagent Final concentration Quantity
Dexamethasone 10 mM 5 mL from 10 mM stock
DMEM/F12 1× 5 mL
Collagenase NB6 solution
Store the lyophilized collagenase powder at 2–8 °C under desiccated conditions. Dissolve 100 mg of powder with 10 mL of sterile PBS (cGMP) to make a stock concentration of 10 mg/mL. Aliquot the solution to 0.15 mL in sterile 2 mL Eppendorf tubes and store at -20 °C. Prior to tissue digestion, thaw the collagenase aliquot at room temperature for 10–15 min and add 1.35 mL of DMEM/F12 with 0.1% Flexbumin and 1× antibiotics/antimycotic to make a working concentration of 1 mg/mL.
Reagent Final concentration Quantity
Collagenase NB6 powder 1 mg/mL 0.15 mL of 10 mg/mL stock
DMEM/F12 1× 1.35 mL
Flexbumin 0.1% (v/v) 6 mL
Antibiotics/antimycotic 1× 15 mL from 100× stock
GMP fibronectin
Store lyophilized fibronectin at -20 °C under desiccation for a maximum of 2 years. Thaw the powder at room temperature for 10–20 min. Prior to opening, briefly centrifuge the vial at 400× g for 10–15 s to bring the contents to the bottom. Add 1 mL of injection water to 500 mg of fibronectin powder to make a stock concentration of 500 mg/mL, aliquot to 50 mL volume, and store at -80 °C for up to a year or at 2–8 °C for a week.
Fibronectin coating: Dilute GMP fibronectin at 500 mg/mL stock with sterile PBS to 10 mg/mL working concentration. Apply 1 mL per TCF25 (tissue culture flask, 25 cm2) and coat at room temperature for 15–30 min. Remove the solution, and the surface is ready for cell seeding.
Reagent Final concentration Quantity
Fibronectin 10 mg/mL 20 mL of 500 mg/mL stock
PBS 1× 1 mL
CryoStor CS5
Store CryoStor CS10 at 2–8 °C. Prior to use for CSSC cryopreservation, dilute CS10 with GMP culture medium (1:1 v/v) to make a working CS5 solution with 5% DMSO and keep at room temperature for immediate cell storage or at 2–8 °C for one week. The step of cell freezing requires the use of a controlled temperature apparatus (e.g., Mr. FrostyTM freezing container with isopropanol).
Reagent Final concentration Quantity
GMP culture medium 0.5× 0.5 mL
CryoStor CS10 0.5× 0.5 mL
GMP stem cell culture medium (GMP SCCM)
Thaw the frozen reagents (including recombinant human EGF, recombinant human PDGF-BB, and Dex) at room temperature for approximately 10–15 min or at 2–8 °C overnight. Mix the thawed solutions by gently inverting the Eppendorf vial a few times, followed by a brief spinning with a benchtop micro-centrifuge.
To prepare SCCM with 500 mL volume, mix 300 mL of DMEM (1,000 mg/L D-glucose) and 200 mL of Ham’s F12. Add reagents with the required volume, filter-sterilize, and bring the culture medium to room temperature before use.
Reagents Final concentration Volume
DMEM (1 g/L D-glucose, with GlutaMAX) 60% (v/v) 290 mL
Ham’s F12 40% (v/v) 190 mL
BioWhittaker antibiotic-antimycotic (100×) 1× 5 mL
Insulin-transferrin-selenium (100×) 0.5× 2.5 mL
2-Phospho-L-Ascorbate (100 mM) 0.5 mM 2.5 mL
Recombinant human EGF (10 mg/mL) 10 ng/mL 0.5 mL
Recombinant human PDGF-BB (10 mg/mL) 10 ng/mL 0.5 mL
Dexamethasone, USP micronized (10 mM) 10 nM 0.5 mL
Flexbumin (25%) 0.1% (v/v) 2 mL
Human serum (100%) 2% (v/v) 10 mL
Notes:
All reagents used in this GMP culture formulation adhere to the GMP standards. During the development of the GMP-compliant formulation, we tested different GMP substitutes for the research-grade chemicals/reagents that are used in the lab-based culture medium [19]. This was done to ensure that the formulation meets the GMP standard. The corresponding reagents were comprehensively evaluated against the original research-grade components through rigorous cellular and molecular assays using at least three primary donor CSSC batches. After evaluating multiple GMP-grade reagents, we meticulously developed an optimized formulation that closely mirrors the research-grade culture medium.
It is worth noting that human serum can be lipid- and protein-rich, depending on the donors. This can cause the culture medium to appear cloudy with the presence of insolubilities, although it is sterile. For this reason, we suggest that the final medium should be filter-sterilized before use.
Laboratory supplies
Cell strainer (40 and 70 μm pore size) (Fisher Scientific, catalog numbers: 22362537 and 22362548)
Cryovials (1.8 mL volume) (Fisher Scientific, catalog number: 1050026)
Tissue culture flask (TCF) 25 cm2 (Corning, catalog number: 430168), 75 cm2 (Genesse Scientific, GenClone, catalog number: 25-209); tissue culture dish (TCD) 30 mm (Falcon, catalog number: 353001), 60 mm (Falcon, catalog number: 353002), 100 mm (diameter) (Genesse Scientific, GenClone, catalog number: 25-202); tissue culture plate (TCP) 6-well (Fisher Scientific, catalog number: EB012927)
Sterile syringe (10 mL volume) (BD, catalog number: 305482)
Sterile centrifuge tubes (15 mL and 50 mL volume) (VWR, catalog numbers: 525-1069 and 525-1014)
Sterile microcentrifuge tubes (1.5 mL and 2 mL volume) (Fisher Scientific, catalog numbers: 05-408-129 and 02-681-321)
Sterile syringe filters (0.22 mm pore size) (Nalgene, catalog number: 726-2520)
Equipment
Biological safety cabinet (Class II type A2/B2) (Baker, model: SteriGARD III Advance)
Phase-contrast microscope with 4×, 10×, and 20× objectives (Invitrogen, model: EVOS XL Core)
Stereomicroscope for dissection (0.8 to 16×) (AMScope, model: SM-1 LED 1445)
Humidified CO2 incubator (Thermo, model: HERAcell 150i)
Temperature-controlled benchtop centrifuge (Thermo, catalog number: 75004381)
Automated cell counter (Invitrogen, model: CountessTM)
Colibri toothed forceps
Vannas curved scissors (6 mm blade) for corneal procedures
Scalpel handle #3
Sterile surgical blade #10 (Integra Miltex carbon steel sterile surgical blades, model: 4-110)
Germinator glass bead sterilizer (Fisher Scientific, model: Microbead Sterilizer B1305-FIS)
Rotator (set at 30 rounds per minute, rpm)
Mr. FrostyTM freezing container (with isopropanol) (Nalgene Cryo freezing container, model: 5100-0001)
Liquid nitrogen cell storage (Thermo, model: Thermolyne CY509975)
Digital balance (Denver Inst Co., model: TR2102)
Procedure
Donor corneas
Donor selection: Donor assessment will be performed by a certified eye bank (Eversight Eye Bank, Cleveland, OH, Chicago, IL, Ann Arbor, MI, and Clark, NJ). To be considered for donation, donors should be younger than 60 years old, with no history of cancer or drug use. Informed consent from the next of kin of all deceased donors is necessary. The corneas must be suitable for clinical use and free of any known disease, injury, or inflammation. Additionally, the donor endothelial cell count should be greater than 2,000 cells/mm2. Negative serology results for transmissible diseases (including but not limited to human immunodeficiency virus, hepatitis B and C, and syphilis) are also required. The time from death to tissue recovery and preservation must be less than 12 h. The time from preservation to arrival at the cell facility should not exceed 5 days.
Note: It is important that one of the donor inclusion criteria is that the corneal endothelial cell count should be greater than 2,000 cells/mm2. A low corneal endothelial cell count may be associated with stromal edema and an altered stromal environment. This could affect the viability and cell features of native stromal cells, including possibly the CSSCs at the limbal stroma.
Cornea harvest and delivery: Donor cornea harvest will be performed under sterile conditions by a trained technician from a certified eye bank. Corneas are preserved in Optisol GS (Bausch & Lomb) or in other equivalent FDA-approved storage solutions under hypothermic storage (2–8 °C) conditions, without freezing, for shipment to the cell facility. For shipment, each cornea is placed in a primary container (sterile moist chamber) (Figure 1A), then in a temperature-controlled shipping box (at 2–8 °C).
Receiving corneal tissue: On the arrival of shipment, confirm that the primary moist chamber is without cracks or leakage. Before opening, wipe clean the container with 70% alcohol.
Corneal tissue sterilization: Using sterile forceps from the Germinator glass bead sterilizer, and after cooling, transfer corneal tissue to a TCD60 with 6–8 mL of DMEM/F12 containing 1× antibiotic/antimycotic at room temperature and rinse two times for 10 min each (Figure 1B).
Figure 1. Processing steps of donor corneal tissue. (A) Donor corneal tissue stored in a primary container (sterile moist chamber) for delivery. (B) Tissue sterilization in DMEM with 2% antibiotics. (C) Removal of conjunctiva/tenons on the anterior side of donor cornea. (D) Clearing of all tissues on the posterior side.
Preparation prior to cell isolation
Collagenase NB6 working solution: Thaw collagenase aliquot at room temperature for 10–15 min and add 1.35 mL of DMEM/F12 with 0.1% Flexbumin and 1× antibiotics/antimycotic (at room temperature) to make a working concentration of 1 mg/mL.
Note: Avoid repeat freeze-thaw cycles of collagenase, as this process damages the physical protein structure and affects the enzymatic activities.
Fibronectin coating of culture surface: Dilute GMP fibronectin at 500 mg/mL stock to 10 mg/mL working concentration. Apply 1 mL per TCF25 and coat at room temperature for 15–30 min. Remove solution, and the surface is ready for cell seeding.
Corneal tissue processing
Prepare 4–5 TCD100 with sterile PBS (up to 15 mL). Transfer corneal tissue to the first TCD100. Under stereomicroscopy magnification 0.8–2×, remove conjunctiva/tenons on the anterior side with a pair of toothed forceps and curved Vannas micro-scissors (Figure 1C).
Note: Ensure the complete removal of conjunctiva and tenons to avoid contaminating fibroblasts in the CSSC culture. This step can be assisted by further scraping the anterior surface using a #10 surgical blade.
Using a #10 surgical blade, scrape to remove corneal and conjunctival epithelium.
Note: A complete removal of epithelium is necessary to avoid epithelial cells overgrowing the CSSC in culture. The use of dispase can be included but no GMP reagent is available.
On the posterior side, clear all tissues (including iris, corneal endothelium, and trabecular meshwork tissues) with a #10 surgical blade (Figure 1D).
After rinsing the corneal tissue in the second TCD100 with sterile PBS (up to 15 mL), position the corneal tissue with the posterior side facing up. Remove the central cornea using a circular trephine (8 mm diameter), leaving the peripheral cornea (approximately 0.5–1 mm width) and the limbus (a transition from the transparent cornea to the opaque scleral tissue) undisturbed (Figures 2A–C).
Figure 2. Steps of isolating anterior limbal stroma . (A) Whole adult cornea (~12 mm diameter) without the central region after removal by a circular surgical trephine. (B–C) Cut the corneal rim into quadrants. (D–F) Trimming of limbal strip with ~0.5 mm on scleral side and 1–1.5 mm on corneal side. (G–I) Making vertical cuts to separate anterior limbal stroma from the rest of tissue. (J) Separation of anterior limbal stroma. (K) Cuts of anterior limbal stromal strip into small blocks for efficient collagenase digestion. *denotes the anterior limbal stroma.
Anterior limbal stroma isolation
Cut the corneal rim into quadrants with a #10 surgical blade.
At the exterior side of the limbus, make a vertical cut to remove the sclera, leaving approximately 0.5 mm (width) scleral tissue with the limbus (Figures 2D–F).
After rinsing the limbal strip quadrants in the third TCD100 with sterile PBS, position the limbal strip slightly vertical and trim to obtain the anterior 1/3 to 1/2 of the thickness (Figures 2G–J).
Note: CSSCs are located at the anterior limbal stromal region near the basement membrane [18,20].
After rinsing the anterior limbal tissue in the fourth TCD100 with sterile PBS, cut the tissue into small blocks, approximately 1 mm3 in size (Figure 2K).
Transfer tissue blocks to the 2 mL tube with 1 mg/mL collagenase NB6 solution (see Recipes).
Digest tissues for 6–8 h at 37 °C with slow rotation at 30 rpm.
Note: At the last hour of digestion, triturate the digest with a P1000 pipette tip 5–10 times to assist cell dislodging from the loosened stromal matrix by collagenase activity. Return the mixture to 37 °C incubation for the last hour of digestion.
After digestion, triturate the lysate with a P1000 or P200 (depending on the extent of tissue digestion).
Pass the digest through a 70 μm cell strainer to obtain a single-cell suspension, wash the cell strainer with 10 mL of PBS, and collect into a 50 mL centrifuge tube.
Centrifuge at 250× g in a swinging-bucket rotor for 5 min at room temperature.
Gently resuspend the pellet in 3 mL of GMP SCCM (see Recipes) at room temperature and transfer to a 15 mL centrifuge tube.
Note: The pellet should not be visible, as not many cells can be obtained from a single anterior limbal stroma. If a visible pellet is noticed, it could mean that the limbal stromal matrix has not been digested properly. Check that the cell strainer is being used correctly and repeat the collagenase digestion for 1–2 h followed by trituration to ensure that cells are well dislodged. Then, repeat the filtration process through the cell strainer to obtain a single-cell suspension.
Spin again at 250× g in a swinging-bucket rotor for 5 min at room temperature.
Resuspend the pellet in GMP CSSC (5 mL of volume) and seed to the fibronectin pre-coated TCF25.
Place the primary culture (passage 0, P0) in a humidified CO2 incubator at 37 °C with 5% CO2 balanced with air.
Primary donor CSSC culture at P0
Under phase-contrast microscopy, check the primary cell clusters (with 4–10 cells) that usually appear within 3–7 days (Figure 3). They should be homogeneously small-sized cells. Mark 2–3 clusters that appear early in the P0 culture and monitor their confluence status. Once sufficient CSSC clusters are detected, remove the culture medium and replenish with fresh GMP SCCM. Repeat medium change every 48–72 h.
The clusters expand with increasing cell numbers and become confluent at the center region (Figure 3). When those marked clusters reach 50%–70% confluence and with increasing cell–cell contacts, the P0 culture should be sub-passaged by trypsinization. If no clusters appear after 14 days, the primary culture is considered to have failed and should be discarded.
Notes:
Our GMP medium formulation has been developed based on research published earlier [18,10] to support the growth of donor CSSCs. Although other cell types such as endothelial cells and neutrophils may briefly attach to the culture surface, they do not proliferate or form detectable colonies.
On days 3–4, if no cells adhere to the culture surface, add 1–2 mL of fresh medium without discarding the used medium to replenish the nutrients and growth factors. Such supplements can improve cell viability and attachment.
Multiple CSSC clusters/clones can be generated during P0 culture, and they have different initiation times, cell density, morphology, homogeneity, and growth rate. Early-formed clones tend to contain more cells and be more confluent than late-formed clones. To monitor cell growth and avoid over-confluence in early clones, it is recommended to mark the first few clones that appear in the early stages of P0 culture. Once these marked colonies reach approximately 50%–70% confluence or increased cell–cell contact and packing, it is necessary to sub-passage the culture. This will prevent stem cell changes that may result from high confluence, e.g., cell differentiation and the loss of stem cell properties.
The quality and yield of primary CSSCs can be affected by various factors. These factors include the freshness and viability of the donor corneal tissue, sub-optimal conditions during tissue processing, and poor cell survival or cell damage after the tissue’s enzymatic digestion. Additionally, the transition from an in vivo matrix environment to an in vitro condition with different substrate adhesion can be a crucial factor in determining cell survival and behavior. Therefore, it is not uncommon to have no viable CSSCs in the P0 culture. Even though the early clusters can form and be detected, this does not guarantee the continuous survival and growth of primary cells. In fact, primary cells may undergo early senescence or cell death due to poor adaptability to the new environment.
Figure 3. Primary good manufacturing practice (GMP)-corneal stromal stem cell (CSSC) cluster (TPF-23-29) at P0 and cell propagation. Phase-contrast micrographs showing the cell cluster with 6–8 cells detected at day 3 post-seeding of primary CSSCs freshly isolated from anterior limbal stroma. The expansion of the same cluster at day 5, 7, and 10 showed an exponential increase in stem cell number. At day 12, the clonal expansion generated colonies with >500 cells. Magnified image at day 12 (bottom right) shows cells with moderate cell–cell contact at approximately 70% confluence. This stage is suitable for sub-passaging and cryopreservation. Scale bar, 50 μm.
P0 sub-passaging, viable cell count, and cryopreservation
When early cell clusters at P0 reach 50%–70% confluence with increasing cell–cell contacts (Figure 3), cells are sub-passaged by trypsinization:
Remove the medium by aspiration.
Add 5 mL of sterile PBS to each TCF25, rinse, and discard.
Add 1 mL of TrypLE Select CTS and allow brief trypsinization for approximately 1 min at 37 °C.
Dislodge cells from the culture surface by rocking the flask side to side or pipetting 5–10 times.
Collect cell suspension into a 15 mL conical centrifuge tube. Rinse TCF25 with 5–8 mL of sterile PBS and collect to the cell suspension.
Centrifuge at 250× g for 5 min at room temperature in a swinging-bucket rotor.
Resuspend the cell pellet in GMP SCCM (1 mL) and mix well to single-cell suspension.
Mix 10 mL of cell suspension with 10 mL of 0.4% Trypan blue for cell counting and viability check.
Calculate total cell yield. Take 2,000 cells for P1 culture in a new TCF75 pre-coated with GMP fibronectin (see Note). Add 10–12 mL of GMP SCCM to each TCF75. For even seeding, gently rock the flask from side to side. Incubate the flask at 37 °C with 5% CO2. Refresh medium every 2–3 days. Examine cell distribution and growth confluence under the phase-contrast microscope.
Calculate the number of P0 cells and number of aliquots for cryopreservation (see Note).
Centrifuge the cell suspension at 250× g for 5 min at room temperature in a swinging-bucket rotor.
Prepare cryovials (n = 6 each cell batch) labeled with cell ID, quantity, and date of preparation.
Prepare cryopreservation medium by diluting CS10 with GMP SCCM (1:1 vol/vol) to CryoStor CS5 (see Recipes). Add 3 mL of cryopreservation medium to resuspend the cell pellet.
Put 0.5 mL of cell suspension to each labeled cryovial.
Freeze cryovials in a controlled freezing apparatus at -80 °C from overnight to 24 h.
Transfer frozen vials to liquid N2 and store at the gas phase position.
Notes:
The cell seeding density is suggested after our evaluation with more than 15 primary CSSC batches. The cells propagated with the GMP SCCM for 5–7 days in a TCF75 without reaching more than 80% confluence. However, this may vary from case to case. Some stem cell batches may grow faster and others not. Hence, it warrants an optimization based on the suggested seeding density.
In Funderburgh et al. [20], authors showed that healthy primary CSSCs have 50–70 doublings before undergoing senescence, apoptotic cell death, or adverse cell features [20]. This represents that CSSCs can generally be sub-passaged up to three times to Passage 3 without detectable cell changes.
It is advisable to freeze P1 cells at 2,000–6,000 cells/cryovial so that the cells after thawing can be seeded up to 3× TCF75 to generate increasing cell quantity for P3 experiments.
Thawing CSSCs
When thawing CSSCs, allow at least one passage of recovery and expansion before plating for experiments. Since the cells should be used within the first three passages, it is advisable to freeze more cell aliquots at P1 so that the thawed P2 cells can be recovered and expanded for p3 experiments.
Prepare fibronectin-coated culture surface (see Recipe 5).
Per cryovial, prepare a 15 mL conical centrifuge tube with 9 mL of GMP SCCM.
Remove the cryovial from liquid N2 storage and float it in a 37 °C water bath until a tiny ice clump is left.
Transfer 1 mL of GMP culture medium to the cryovial, mix by pipetting up and down, and transfer the cell suspension to the prepared 15 mL tube with culture medium (10 mL) at room temperature.
Mix well and centrifuge the tube at 250× g for 5 min at room temperature.
Remove supernatant and add 1 mL per TCF75 (~2,000 cells per mL) to resuspend the pellet by carefully pipetting up and down under single-cell suspension.
Add 10–12 mL of GMP SCCM to each fibronectin-coated TCF75 and transfer cell suspension at 1 mL per TCF75.
To evenly distribute the cells, gently rock the flask from side to side.
Put the flasks inside the CO2 incubator at 37 °C.
Examine cell growth and distribution under phase-contrast microscopy the next day. After having visible cell colonies, perform GMP medium change every other day.
Notes:
Good CSSC will give clonal expansion from single cells in 3–5 days.
During propagation, stem cell differentiation from CSSCs to corneal stromal keratocytes, the expression of stem cell markers [e.g., ABCG2 (ATP-binding cassette super-family G member 2), Pax6, nestin, Bmi-1 (B lymphoma murine leukemia viral insertion region 1)], in vitro quality control, and in vivo efficacy tests in reducing corneal scar formation in animal models of corneal stromal injury can be executed. Details of these assays are given in our early studies [10,15,16,19].
Validation of protocol
This protocol has been used and validated in the following research article:
• Santra et al. [19]. Good manufacturing practice production of human corneal limbus-derived stromal stem cells and in vitro quality screening for therapeutic inhibition of corneal scarring. Stem Cell Res Ther. 2024 Jan 8;15(1):11. DOI: 10.1186/s13287-023-03626-8.
The GMP protocol is optimized from the previous donor CSSC protocol using research-grade chemicals and reagents. It was validated in the following publications:
Du et al. [18]. Stem Cells; DOI: 10.1634/stemcells.2004-0256
Du et al. [24]. Invest Ophthalmol Vis Sci; DOI: 10.1167/iovs.07-0587
Du et al. [25]. Stem Cells; DOI: 10.1002/stem.91
Basu et al. [10]. Sci Trans Med; DOI: 10.1126/scitranslmed.3009644
Hertsenberg et al. [12]. PLoS One; DOI: 10.1371/journal.pone.0171712
Shojaati et al. [26]. Stem Cells Trans Med; DOI: 10.1002/sctm.17-0258
Shojaati et al. [11]. Stem Cells Trans Med; DOI: 10.1002/sctm.18-0297
Khandaker et al. [15]. Exp Eye Res; DOI: 10.1016/j.exer.2020.108270
Weng et al. [14]. Eye Vis; DOI: 10.1186/s40662-020-00217-z
Jhanji et al. [16]. Int J Mol Sci; DOI: 10.3390/ijms23136980
Yam et al. [17]. J Adv Res; DOI: 10.1016/j.jare.2022.05.008
General notes and troubleshooting
General notes
Primary CSSC colonies typically manifest within 3 to 5 days. Clonal growth on this selective medium is a unique characteristic of stem cells. Although other cell types, such as endothelial and neutrophils, may attach, they do not survive as colonies. CSSC clones, which have uniformly small-sized cells, can be identified visually.
If no colonies appear within 14 days, the culture is deemed poor or without growth, and it should be terminated by adding diluted bleach solution before disposal.
GMP stem cell culture medium (SCCM) can be stored at 4 °C for a maximum of one week.
Troubleshooting
Problem 1: A major challenge faced is the transition of CSSC to fibroblast types, particularly in serum and growth factor–supplemented conditions. The fibroblasts are different from the small-sized CSSCs and have a heterogenous slender to stellate shape. However, these fibroblast-like cells can arise within the CSSC clones (Figures 4C and D).
Solution: If there is mild fibroblast growth among CSSC clusters, perform a brief trypsinization using pre-warmed TrypLE Select for less than a minute’s incubation at 37 °C. This will dislodge the primary CSSCs, which can be collected for the next passaged culture. This will cause a substantial loss of CSSCs, but the cell purity/enrichment can be improved. If fibroblasts persist substantially, terminate the culture (Figure 4E). Additionally, the incomplete removal of limbal epithelium (refer to step C2) will result in the formation of epithelial cell colonies (Figure 4F). They usually grow faster than CSSCs, and the culture should be terminated.
Problem 2: The preservation of corneal tissue in Optisol GS has not been optimized for the maintenance of CSSC viability. Prolonged storage of donor tissues in Optisol may have unexpected adverse effects on CSSCs.
Solution: It is recommended to process the corneal tissues as soon as possible upon receiving the samples, to avoid any potential harm to CSSCs.
Figure 4. Potential contaminating cell types in primary good manufacturing practice (GMP)-corneal stromal stem cell (CSSC) culture. Phase-contrast micrographs showing (A) proper CSSC cluster with clonal expansion at P0 and (B) the growth of small-sized proliferating cells at P1. (C) Growth of fibroblast-like cells can occur in CSSC culture. (D) Under higher magnification, they display an heterogenous slender to stellate shape. (E) After sub-culture, the fibroblasts overgrow the CSSCs. (F) In case the limbal epithelium is not completely removed, epithelial colonies are formed due to the fast-growing limbal epithelial stem cells. Scale bar 120 μm.
Acknowledgments
The authors would like to thank the support from NIH U01 [EY035252], the Hillman Foundation, and unrestricted funds from Research to Prevent Blindness and the National Eye Institute [P30, EY008098]. We also thank the Eversight Eye Bank for the procurement of transplantation-grade donor corneas with informed consent. Research activities at Eversight are supported by the Eppley Foundation for Research, Connecticut Lions Eye Research Foundation, Albert G. and Olive H. Schlink Foundation, Lowell F. Johnson Foundation, William G. and Helen C. Hoffman Foundation, and the Louise H. and David S. Ingalls Foundation. We thank Ophthalmology Core Facility – Image Acquisition & Analysis (led by Lathrop K, Ph.D.), and Billig I, Ph.D. for the assistance in experiments and coordination. Hsu YMS was supported by Hillman Senior Fellowship for Innovative Cancer Research.
Graphical overview created with BioRender.com under license UV26P7FRKJ, accessed on Apr 14, 2024.
Competing interests
The authors declare that they have no competing interests.
Ethical considerations
The use of donor corneas followed the tenets of the Declaration of Helsinki and was approved by The University of Pittsburgh, Committee for Oversight of Research and Clinical Training Involving Decedents (CORID), Protocol #161, “Stem cells for corneal engineering & regeneration” renewal approved on May 9, 2023; and the Human Stem Cell Research Oversight (hSCRO), Protocol ES-07-019-R, “Corneal Stem Cell Therapy” renewal approved on May 15, 2023.
References
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Hertsenberg, A. J., Shojaati, G., Funderburgh, M. L., Mann, M. M., Du, Y. and Funderburgh, J. L. (2017). Corneal stromal stem cells reduce corneal scarring by mediating neutrophil infiltration after wounding. PLoS One. 12(3): e0171712.
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Article Information
Publication history
Received: Apr 30, 2024
Accepted: Aug 11, 2024
Available online: Sep 6, 2024
Published: Sep 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
Stem Cell > Adult stem cell > Mesenchymal stem cell
Cell Biology > Cell isolation and culture > Cell growth
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5,075 | https://bio-protocol.org/en/bpdetail?id=5075&type=0 | # Bio-Protocol Content
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This is a correction notice. See the corrected protocol.
Peer-reviewed
Correction Notice: Expression and Purification of Recombinant Human Mitochondrial RNA Polymerase (POLRMT) and the Initiation Factors TFAM and TFB2M
AH An H. Hsieh §
SR Sean D. Reardon
JM Jubilee H. Munozvilla-Cabellon
JS Jiayu Shen
SP Smita S. Patel
TM Tatiana V. Mishanina
(§ Technical contact)
Published: Aug 20, 2024
DOI: 10.21769/BioProtoc.5075 Views: 276
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After official publication in Bio-protocol (https://bio-protocol.org/en/bpdetail?id=4892&type=0), we found errors that should be corrected:
• In the Recipes section under Part I: POLRMT purification buffers, the note under the 5% PEI solution should say “Adjust pH to 7.9…” instead of “7.0” to match the pH of all the other buffers.
• In the Procedure section for Part I: POLRMT Purification, under part D. POLRMT AS precipitation, there is a math error that should be corrected to “______mL lysate × 0.351 g/mL…” instead of “3.51 g/mL”.
• In step 11 of part E. POLRMT DEAE gravity column, The concentration speed should say “Concentrate POLRMT by centrifuging the Amicon filtration units at 1,300× g” instead of “1,900× g”. Having a lower speed will prevent protein precipitation issues and keep the centrifugation speed consistent with the later steps.
• In section Part III: TFB2M purification, under G. TFB2M concentration, step 3 says “Start centrifuging the TFB2M sample at 2,500× g and gradually decrease the speed to 1,300× g if precipitate starts forming.” We have seen, however, that simply starting at a low speed of 1,300× g is better to prevent precipitation. The sentence should be changed to “Start centrifuging the TFB2M sample at 1,300× g.”
Article Information
Publication history
Received: Aug 14, 2024
Published: Aug 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Visualization and Analysis of Neuromuscular Junctions Using Immunofluorescence
YH You-Tian Hsieh
SC Show-Li Chen
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5076 Views: 611
Reviewed by: Marion HoggIshita ChandelJosé M. Dias
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Original Research Article:
The authors used this protocol in Journal of Cachexia, Sarcopenia and Muscle Aug 2018
Abstract
The neuromuscular junction (NMJ) is an interface between motor neurons and skeletal muscle fibers, facilitating the transmission of signals that initiate muscle contraction. Its pivotal role lies in ensuring efficient communication between the nervous system and muscles, allowing for precise and coordinated movements essential for everyday activities and overall motor function. To provide insights into neuromuscular disease and development, understanding the physiology of NMJ is essential. We target acetylcholine receptors (AChR) by immunofluorescence assay (IFA) with α-bungarotoxin (BTX; snake venom neurotoxins binding to AChR) to visualize and quantify the NMJ. Changes in AChR distribution or structure can indicate alterations in receptor density, which may be associated with neuromuscular disorders or conditions that affect synaptic transmission. This protocol provides the methodology for isolating and longitudinally sectioning gastrocnemius muscle for AChR-targeted IFA for confocal microscopy and performing quantitative analysis of NMJs.
Key features
• Visualizes and quantifies NMJs using α-bungarotoxin.
• Utilizes high-resolution confocal microscopy for detailed imaging.
Keywords: Neuromuscular junction Acetylcholine receptor Immunofluorescence assay α-bungarotoxin
Graphical overview
Schematic workflow of this protocol (figure created in BioRender.com)
Background
The neuromuscular junction (NMJ) serves as a chemical synapse between the axon terminal of a motor neuron and the postsynaptic region on a muscle fiber [1]. At the NMJ, the motor neuron's axon terminal releases acetylcholine (ACh) into the synaptic cleft. This neurotransmitter then binds to nicotinic acetylcholine receptors (AChR) on the muscle fiber membrane, causing an influx of sodium ions and subsequent depolarization of the muscle cell membrane. This depolarization initiates an action potential that travels along the muscle fiber, ultimately leading to muscle contraction. The NMJ is vital for voluntary muscle movement and is precisely regulated to ensure accurate motor control [2].
Structural instability of NMJ can cause several neuromuscular diseases and may occur in the aged population to cause loss of muscle strength and mass [3]. For instance, we have recently found that the nuclear receptor interaction protein (NRIP) is an AChR-interacting protein, functioning as a scaffold to stabilize the AChR complex and playing a physiological role in the neuromuscular system [1]. Without NRIP, the efficacy of synaptic transmission and AChR clustering may decline, leading to the loss of nerve supply to NMJs (denervation) and impaired motor function, as observed in NRIP knockout mice (Figure 3C and D in Chen et al. [4]). Additionally, Lrp4 regulates the AChR clustering to guarantee proper NMJ assembly [5]. Conditional knockout of muscle Lrp4 causes a significant decline in AChR cluster size and number [6]. Besides muscle disorder, nerve injury also causes a decrease in AChR cluster and density. Researchers have used a sciatic nerve crush injury model in mice to simulate nerve damage and observed significant fragmentation of the NMJ and a notable decrease in the AChR cluster area [7].
NMJ analysis is a crucial tool in neuromuscular disease research. By combining high-resolution imaging with molecular studies, researchers gain a comprehensive understanding of NMJ physiology, which aids in developing effective treatments for amyotrophic lateral sclerosis (ALS), spinal muscular atrophy (SMA), and other neuromuscular diseases. Key visualization techniques include immunofluorescence assays (IFA) with confocal microscopy, electron microscopy, and super-resolution microscopy. IFA with confocal microscopy is preferred for its ability to produce high-resolution 3D images, enabling detailed examination of NMJ structures. α-bungarotoxin, a snake venom toxin, is commonly used to bind acetylcholine receptors for precise visualization. Despite its origin, α-bungarotoxin is safe in labs due to non-toxic concentrations. Confocal microscopy enhances IFA by reducing background noise and increasing image clarity, surpassing traditional fluorescence microscopy. While electron microscopy offers ultra-high resolution, its extensive sample preparation makes it less practical. Super-resolution microscopy provides nanometer-scale visualization but is limited by the complexity and specialized equipment needed. Overall, IFA with confocal microscopy balances high-resolution imaging with ease of use and specificity, making it indispensable in neurobiological research and treatment development for neuromuscular diseases.
Materials and reagents
High-profile microtome blades (Leica, catalog number: 14035838383)
Glass insert 70 mm (Leica, catalog number: 14047742497)
Micro slides (Muto Pure Chemicals, catalog number: GA13-7626R)
O.C.T (VWR, catalog number: 95057838)
Aluminum foil (Kirkland, catalog number: RK611)
10× phosphate-buffered saline (PBS) (Omics Bio, catalog number: IB3012)
Sucrose (Sigma-Aldrich, catalog number: S0389-500G)
Glycine, powder (Omics Bio, catalog number: BT5031)
Paraformaldehyde powder (Sigma-Aldrich, catalog number: P6148-500G)
Alexa-594-conjugated α-bungarotoxin (Life Technologies, catalog number: B13423)
DAPI fluoromount-G (SouthernBiotech, catalog number: 0100-20)
Microscope cover glass (Shinetech Inc, catalog number: GA11-2450)
1.5 mL clear microcentrifuge tubes (Corning, catalog number: MCT-150-C)
2,2,2-tribromoethanol (avertin) (Sigma-Aldrich, catalog number: T48402-5G)
Horse serum (Gibco, catalog number: 16050-114)
BSA albumin fraction V (BioFroxx, catalog number: 4240GR100)
Anti-synaptophysin primary antibody (Abcam, catalog number: ab32127)
Anti-neurofilament primary antibody (Abcam, catalog number: ab8138)
488-conjugated donkey anti-rabbit secondary antibody (Jackson ImmunoResearch, catalog number: AB_2340620)
Solutions
1× PBS (see Recipes)
4% PFA (see Recipes)
2.5% avertin (see Recipes)
0.1 M glycine solution (see Recipes)
30% sucrose (see Recipes)
Blocking buffer (see Recipes)
Recipes
1× PBS
Reagent Final concentration Amount
10× PBS n/a 200 mL
Deionized H2O n/a 1800 mL
Total n/a 2000 mL
Note: The 1× PBS solution should be prepared fresh and used within seven days.
4% PFA
Reagent Final concentration Amount
Paraformaldehyde powder 4% 4 g
1× PBS n/a 100 mL
Total n/a 100 mL
Note: To prepare 4% PFA solution, add 4 g of paraformaldehyde powder to 90 mL of 1× PBS, stir, and heat to 60–65 °C until dissolved. Add a few drops of 5 N NaOH to clear the solution (5 N NaOH is used to facilitate the dissolution of paraformaldehyde powder), cool to room temperature, add 1× PBS to 100 mL, and adjust the pH to 7.4. The solution can be prepared in advance and stored in a -20 °C freezer for long-term storage. Thaw the solution 30 min before use to ensure it is fully liquefied.
2.5% avertin
Reagent Final concentration Amount
2,2,2-tribromoethanol 2.5% 250 μL
1× PBS n/a 9.75 mL
Total n/a 10 mL
Note: The 2.5% avertin solution can be prepared in advance and stored in a 4 °C refrigerator for up to one month.
0.1 M Glycine solution
Reagent Final concentration Amount
Glycine powder 0.1 M 0.75 g
1× PBS n/a 10 mL
Total n/a 10 mL
Note: To prepare a 0.1 M glycine solution, add 0.75 g of glycine to 10 mL of 1× PBS in a 15 mL conical centrifuge tube and shake until the solution is clear. There is no need to adjust the pH of the solution. The solution can be prepared in advance and stored in a 4 °C refrigerator for up to two months.
30% sucrose
Reagent Final concentration Amount
Sucrose 30% 15 g
1× PBS n/a 50 mL
Total n/a 50 mL
Note: To prepare a 30% sucrose solution, add 15 g of sucrose to 50 mL of 1× PBS and shake until the solution is clear. The solution can be prepared in advance and stored in a 4 °C refrigerator for up to one month.
Blocking buffer
Reagent Final concentration Amount
BSA albumin fraction V 2% 0.2 g
1× PBS n/a 9.5 mL
Horse serum 5% 500 μL
Total n/a 10 mL
Note: To prepare the blocking buffer, add 500 μL of horse serum to 9.5 mL of 1× PBS in a 15 mL conical centrifuge tube, shake, and then add 0.2 g of BSA albumin fraction V into the solution. Shake until the solution is clear.
Equipment
P2, P200 and P1000 micropipettes
Laser scanning confocal microscope (Zeiss, model: LSM880)
Staining jar (Shineteh Instruments, catalog number: GB11-15)
Slide racks (Shineteh Instruments, catalog number: GB11-15S)
Liquid nitrogen insulation barrels (Shineteh Instruments, catalog number: IE1-D6000)
Cryostat (Leica, catalog number: CM3050)
4 °C, -20 °C freezers [Sampo, catalog number: SR-C61G(K3)]
-80 °C freezers (Panasonic, catalog number: MDF-U74V)
Laminar flow cabinet (CHUAN, catalog number: TBH-520M)
Orbital shaker (TKS, catalog number: OS-701)
Disposable syringe with needle (Terumo, catalog number: MDSS01S2613)
Disposable syringe (Terumo, catalog number: MDSS20ES)
Iris scissors (Shineteh Instruments, catalog number: ST-S009)
Fine curve point tweezers (Shineteh Instruments, catalog number: ST-NO7)
Super fine point tweezers (Shineteh Instruments, catalog number: ST-NO3)
Iris forceps (Shineteh Instruments, catalog number: ST-I210)
Lab mat (Dogger, catalog number: D4EG-031055)
FalconTM 15 mL conical centrifuge tubes (Thermo Fisher, catalog number: 14-959-53A)
Immuno-pathology staining kit (PEOPLE LIFE TEK CO., LTD., catalog number: H198099910)
Software and datasets
ImageJ image processing and analysis software (https://imagej.net/ij/)
ZEN microscopy software (https://www.zeiss.com/microscopy/en/products/software/zeiss-zen.html)
GraphPad Prism 8.0.2
Procedure
Mouse dissection
Set up the dissection area with a 40 cm × 50 cm lab mat in a clean and well-lit workspace.
Restrain the mouse as shown in Graphical overview. Insert the disposable syringe with a needle into the lower-right quadrant of the abdomen to avoid puncturing internal organs. Gently and slowly inject 0.5 mL of 2.5% avertin (Figure 1) (volume of avertin per mass: 0.2 mL/10 g body weight).
Note: Alternatively, mice can be anesthetized with isoflurane using a gas anesthesia instrument, instead of using avertin.
Wait 5 min until the mouse is completely unconscious.
Note: Tap the tail of the mouse to confirm that it does not respond to stimuli at all.
Gently place the mouse with its ventral side facing upward. It is recommended to pin the mouse to prevent movement during dissection. Use scissors to make a midline incision through the skin from the lower abdomen to the chest. Continue cutting through the muscle layers to expose the ribcage. Then, carefully cut through the ribs on both sides to open the chest cavity and expose the heart.
Gently lift the heart using iris forceps and create a small incision (0.5–1.5 mm wide) in the right atrium with iris scissors. Insert a perfusion needle (connected to the perfusion syringe) into the left ventricle, ensuring the needle tip penetrates the ventricular wall to prevent leakage (Figure 2).
Figure 1. Avertin intraperitoneal injection. Gently restrain the mouse, insert the needle into the lower abdomen slightly to the right of the midline, and slowly inject 0.5 mL of 2.5% avertin solution. Scale bar, 3 cm.
Figure 2. Schematic figure of mouse perfusion. Gently restrain the mouse. Create a 0.5–1.5 mm incision with iris scissors. Perfuse with 1× PBS at a steady rate (10–15 mL/min) using a perfusion syringe with a needle inserted into the left ventricle. The effluent will exit the heart from the incision at the right atrium. (created in BioRender.com)
Begin perfusing with PBS at a steady rate (10–15 mL/min). This step clears the blood from the circulatory system. You will see the liver and other organs blanch as the blood is replaced with PBS.
Continue perfusing until the effluent (fluid exiting the incision in the right atrium) runs clear, indicating that the blood has been adequately removed.
Remove the skin and main organs in the abdominal cavity including stomach, liver, kidneys, small and large intestine, and spleen to facilitate the subsequent dissection steps.
To expose the muscle and knee, remove the skin from the area around the hindlimb by lifting the skin with fine-point tweezers and cutting the fascia between the skin and muscle with iris scissors.
Separate the hindlimbs from the body by cutting the knees with iris scissors.
Incise the Achilles tendon at the ankle (Figures 3 and 4); then, use super fine point tweezers to pull the severed tendon toward the knee (Figures 3 and 5).
Note: The Achilles tendon is the tendon shared by the soleus and gastrocnemius muscle.
Figure 3. Schematic image illustrating the posterior view of the hindlimb of mice, highlighting key muscles and Achilles tendon. The knee is indicated with a black box, and the Achilles tendon is highlighted in purple at the back of the ankle. The quadriceps (QUAD) are located at the front of the thigh. The tibialis anterior (TA) runs along the front of the lower leg. The gastrocnemius (GAS) and soleus (SOL) muscles form the bulk of the calf. (created in BioRender.com)
Figure 4. Incision of the Achilles tendon. Use iris scissors, carefully sever the Achilles tendon (yellow arrow). Scale bar, 1 cm.
Figure 5. Precise tendon grasp for gastrocnemius isolation. Delicately grasp the severed tendon (blue arrow) using super fine point tweezers and gently draw it toward the knee. Do not pull too hard or too fast to prevent damaging the muscle structure. Scale bar, 1 cm.
Cut the tendon in the knee region with iris scissors to separate the muscle from the hindlimb.
Remove the soleus muscle and use fine curve point tweezers to lift the fascia, a thin transparent membrane that sits on the surface of the muscle, from the gastrocnemius muscle. Then, cut it with iris scissors (Figure 6).
Note: The gastrocnemius muscle was chosen for the experiment because it is large enough for cryosectioning. Alternatively, soleus and tibialis anterior muscle are also acceptable (see General note section).
Figure 6. Detachment of soleus muscle. Carefully extract the soleus muscle (blue arrowhead) and remove any residual fascia surrounding the gastrocnemius muscle with iris scissors. Scale bar, 1 cm.
Preparation for cryosection
Subsequent to their separation from the mouse, submerge the gastrocnemius muscles in 4% paraformaldehyde in a 15 mL conical centrifuge tube and shake on the orbital shaker for 2 h at room temperature for fixation.
Wash the muscle tissues three times with approximately 10 mL of 1× PBS in the 15 mL conical centrifuge tube, each time for a duration of 10 min, at room temperature.
Incubate the muscle tissue with rotation in 10 mL of 30% sucrose in a 15 mL conical centrifuge tube at 4 °C overnight to facilitate dehydration.
Place the muscle on a clean lab mat for subsequent experiments.
Make a cube mold with aluminum foil (Figure 7) and fill it with O.C.T. compound until it is 70% full.
Figure 7. Representative image of a cube mold made from aluminum foil for embedding the muscle tissue in the O.C.T compound. Scale bar, 1 cm.
Using forceps, carefully position the muscle tissue longitudinally in the center of the cube mold. Ensure that the entire tissue is submerged in the O.C.T compound.
Note: It is strongly recommended to mark the side that aligns with the intended cutting plane to avoid sectioning in the wrong direction. The intended cutting plane should be parallel to the longitudinal axis of the muscle.
Using forceps, carefully hold the cube mold and immerse it into the liquid nitrogen in the liquid nitrogen barrel. To ensure there is no direct contact between the O.C.T. compound and the liquid nitrogen, do not submerge the mold with liquid nitrogen.
Note: Placing the mold with dry ice, instead of liquid nitrogen, is acceptable for the O.C.T to solidify.
Once the O.C.T. compound has solidified, transfer it to a cryostat for immediate sectioning or store it at -80 °C for future use.
Set the cryostat temperature to -20 °C and install a new or sharp cryostat blade and anti-roll plate.
Remove the aluminum foil from the O.C.T.-embedded muscle tissue block and place it into the cryostat chamber.
Mount the O.C.T. block on the specimen holder of the cryostat with the muscle tissue oriented longitudinally (parallel to the direction of sectioning).
Trim the block face with a thickness of 50 μm using the cryostat blade until the muscle tissue is exposed and a smooth surface is achieved.
Start sectioning the muscle tissue slowly and carefully into slices with a thickness of 30 μm each. Maintain a steady hand and consistent speed to produce even sections.
Note: The reason for cutting such thick sections is to preserve the integrity of the muscle fibers and neuromuscular junctions (NMJs), which can be compromised in thinner sections. This preservation ensures that the fine details of the NMJ are more easily observable.
After ensuring the section is not curled or folded, attach the section with the pre-labeled slide. The section will adhere to the slide due to the temperature difference. (The slides are kept at room temperature throughout the entire procedure.)
Continue cutting and collecting sections until you have the desired number of sections.
Note: At least three sections should be placed on one slide.
After sectioning, wrap the tissue in aluminum foil and store it at -80 °C for future use.
Quenching (background fluorescence reduction)
Wash the slide carrying the sectioned tissue with approximately 250 mL of 1× PBS three times for 10 minutes each at room temperature to eliminate the O.C.T. compound. The slide-washing procedure entails placing the slides onto the slide rack, fully submerging the slide rack into 1× PBS in the staining jar, and then positioning the staining jar on the orbital shaker, shaking at 60 rpm. It is strongly recommended not to shake at speeds exceeding 60 rpm, as this may cause the tissue to detach from the slides.
Take the slide out of the slide rack.
Note: See the troubleshooting section about signal weakness when staining with antibodies for neurofilament or synaptophysin to visualize axonal innervation to NMJs.
Apply 200 μL of 0.1 M glycine solution onto each section and allow it to incubate for 40 min at room temperature, ensuring complete coverage of the tissue by the liquid. This step helps reduce tissue autofluorescence and quench free aldehydes left over from fixation with paraformaldehyde. This step does not require shaking.
Dump the glycine solution from the slide.
Place the slide back in the slide rack and submerge the slide rack in approximately 250 mL of 1× PBS in the staining jar.
Place the staining jar onto the orbital shaker and shake at 60 rpm, three times for 10 minutes each at room temperature to eliminate any residual glycine.
Probe primary antibody (optional)
Take out the slide from the slide rack and place it in the immuno-pathology staining kit.
Apply 200 μL of blocking buffer onto each section and allow it to incubate for 1 h at room temperature, ensuring complete coverage of the tissue by the liquid. This step reduces nonspecific binding.
Dump the blocking buffer from the slide and place the slide in the immuno-pathology staining kit.
Combine anti-synaptophysin primary antibody and anti-neurofilament primary antibody, each diluted at a 1/200 ratio in blocking buffer. Apply 200 μL of this solution onto each section.
Incubate in an immuno-pathology staining kit overnight at 4 °C. This step does not require shaking.
Note: The immuno-pathology staining kit must be humidified. Placing a wet paper towel in the immuno-pathology staining kit is feasible.
Dump the primary antibody solution from the slide.
Place the slide back in the slide rack and submerge the slide rack in approximately 250 mL of 1× PBS in the staining jar.
Place the staining jar onto the orbital shaker and shake at 60 rpm, three times for 10 min each at room temperature.
Probe with α-bungarotoxin and secondary antibody
Take out the slide from the slide rack and place it in the immuno-pathology staining kit.
Apply 200 μL of Alexa-594-conjugated α-bungarotoxin diluted in 1× PBS at a 1/1,000 dilution onto each section and incubate for 2 h at room temperature in darkness. This step does not require shaking.
Note: Add 488-conjugated donkey anti-rabbit secondary antibody to the α-bungarotoxin solution at a 1/1,000 dilution rate. There is no need to apply the secondary antibody if section D has not been conducted.
Remove the α-bungarotoxin solution and proceed to wash the slide three times at room temperature with approximately 250 mL of 1× PBS with shaking on the orbital shaker at 60 rpm for 10 min each time.
Mounting
Take out the slide from the slide rack and place it in the immuno-pathology staining kit.
Add 20 μL of DAPI fluoromount mounting solution onto each section for nucleus staining.
Hold the slide gently, then slowly lower the coverslip onto the slide by gradually tilting it down over the drop of mounting medium. This technique allows the mounting medium to spread evenly and avoids introducing bubbles once the coverslip is in place.
Seal the edges of the coverslip with nail polish.
Air-dry the slide by placing it in a laminar flow cabinet for 20 min.
Visualizing NMJs
Switch on the confocal microscope and ensure that the connected computer is also powered on.
Open the Zen software on the computer. Once the software is running, activate the laser by following the manufacturer's guidelines.
Mount the slide steadily on the stage of the microscope.
Switch to locate mode.
Use the 20× objective lens to scan the sample and identify the region containing the NMJ. The appearance of the NMJ is characterized by red, pretzel-like clusters, which extend less than 15 μm along the z-dimension.
Switch to Acquisition mode within the Zen software.
Configure the imaging parameters as detailed in Table 1. This includes settings for laser intensity, detector gain, and scan speed, among others.
Table 1. Confocal microscopy image snapping condition
αBTX DAPI
Detection wavelength range 566–759 nm 405–481 nm
Scan mode stack, frame
Scaling X 0.277 μm
Scaling Y 0.277 μm
Scaling Z 0.700 μm
Lasers 100% (optimal) 10% (optimal)
Gain 550 (optimal) 520 (optimal)
Objective plan-apochromat 20×/0.8 M27
Direction single
Z-stack slices interval 0.7 μm
Define the Z-stack region by setting the first and the last plane to specify the scanning range. This step is crucial for obtaining a comprehensive three-dimensional image of the NMJ.
Click the Start Experiment button to initiate the scanning process. The system will begin capturing the 3D image based on the defined Z-stack region and the set acquisition parameters.
The figures are processed by orthogonal projection in the XY, XZ, and YZ planes (Figure 8A), and then exported into TIFF format using ZEN software.
Figure 8. Representing data of co-staining of neuromuscular junctions (NMJs) and DAPI. (A) NMJs are labeled by Alexa-594-conjugated α-bungarotoxin, and the image is taken by confocal microscopy. The figure is processed with orthogonal projection in the XY, XZ, and YZ planes. Arrows of different forms point to specific NMJs on three planes. The green and white solid arrowheads indicate two distinct NMJs that are parallel to the XY plane. Although the NMJs pointed to by the white and yellow hollow arrowheads appear to be joined together in the XY plane, their independence can still be discerned by examining the YZ plane figure. Scale bar, 20 μm. A cleared representation of the NMJ in three-dimensional space is demonstrated in Video 1. (B) The comparison of normal NMJ and fragmented NMJ. The upper panel shows a normal morphology of an NMJ, characterized by a pretzel-like structure. The lower panel demonstrates a fragmented NMJ, which exhibits a decrease in area. This decrease will be discussed in the data analysis section. Both NMJs in the panels are parallel to the XY plane. Scale bar: 20 μm. (C) Representative images of co-staining of NMJ, neurofilament, and synaptophysin show a comparison of a fully innervated NMJ (left panel), a partially innervated NMJ (middle panel), and a denervated NMJ (right panel). A nerve terminal and motor endplate were deemed “fully innervated” if they overlapped by 50% or more. An overlap between 20% and 50% classified NMJs as “partially innervated.” Motor endplates without any coverage were identified as “denervated” NMJs [8]. Scale bar, 20 μm.
Video 1. Representation of the three-dimensional (3D) structure of the neuromuscular junction in 3D space
Data analysis
Image type conversion
Launch the ImageJ software.
Go to File > Open and select the image file you want to work with.
Press type in the list and select 8 bit to convert the image.
Scale setting
Select the Straight-Line tool from the toolbar (it looks like a diagonal line).
Draw a line on the image along a known distance (for example, if you have a scale bar on your image, draw the line along the length of the scale bar).
Go to Analyze > Set scale.
In the Set Scale dialog box, the Distance in pixels will be automatically filled in based on the length of the line you drew.
Enter the actual distance that the line represents in the Known distance field (e.g., 10 if the scale bar represents 10 μm).
Select the appropriate unit of length from the Unit of Length field (e.g., micrometers, millimeters, etc.).
If you want the scale to be applied to all future measurements in this image, check the Global option.
Click OK to set the scale.
Threshold adjustment
Go to Image > Adjust > Threshold to open the Threshold dialog box.
The Threshold dialog box displays a histogram of pixel intensities and allows you to set the lower and upper threshold values using sliders.
Adjust the sliders to include the range of pixel intensities that you want to highlight. Pixels within this range will be highlighted in red, indicating the thresholded area. Ensure the connectivity of all pixels.
Press the Apply bottom; this will convert the highlighted regions into a binary image.
NMJ area measurement
Click on the Wand tool in the ImageJ toolbar (it looks like a magic wand) (Figure 9).
Figure 9. Representative example of neuromuscular junction (NMJ) quantification. (A) The red frame indicates the ROI manager dialog box that appears after pressing Ctrl + T. The yellow frame indicates the result window displaying area data after clicking Measure in the ROI manager dialog box. The orange arrow points to the selected NMJ after threshold adjustment. The blue and green arrowheads point to the straight-line tool and the wand tool, respectively. (B) Schematic figure that illustrates the formula for calculating NMJ density. Calculating the NMJ density is another method for analyzing NMJ physiology. The NMJ number is counted under the 10× objective lens of the microscope. The NMJ density is calculated by dividing the NMJ number by the muscle section area in mm2.
Click inside the region you want to analyze. The Wand tool will automatically select the contiguous area with similar pixel values.
Press Ctrl + T to add the selection to the ROI Manager. This will open the ROI Manager dialog box showing all selected regions.
In the ROI Manager dialog box, select the region of interest from the list (Figure 9).
Click Measure in the ROI Manager dialog box. This will open the results window, displaying the measurements of the selected region, including the area. (If there is a hollow area in the NMJ, its area needs to be deducted.)
NMJ area statistic
Calculate the mean value of the NMJ area in mice. Having at least 30 NMJs per mouse and the inclusion of at least three mice is critical: according to the central limit theorem, the distribution of sample means will be approximately normally distributed if the sample size is large enough (typically n ≥ 30). This normality is crucial for the validity of the T-test, which assumes a normally distributed data set. Moreover, including at least three mice ensures that the findings are not specific to a single mouse but are generalizable across multiple individuals.
Set up a column table of GraphPad Prism 8.0.2 software and paste the data for the area of the NMJs.
Go to Analyze in the menu bar and choose t-test (and nonparametric tests) in the submenu.
Select unpaired t-test. If needed, select options for assuming equal variances or not (e.g., Welch's correction).
Click OK. GraphPad Prism will provide the t-value, degrees of freedom, p-value, and confidence intervals.
Validation of protocol
Two published papers can validate the measurements of NMJs:
Our previously published paper, “Muscle-restricted nuclear receptor interaction protein knockout causes motor neuron degeneration through down-regulation of myogenin at the neuromuscular junction” [4], provides a representative image and quantification of NMJ area in gastrocnemius muscle in Figure 3.
The methods for quantifying the NMJ area and calculating the percentage of innervated endplates are demonstrated in Figure 6 of Wong and Martin's 2010 publication, “Skeletal muscle-restricted expression of human SOD1 causes motor neuron degeneration in transgenic mice” [9].
General notes and troubleshooting
General notes
The limitation of this protocol is that it still requires the dissection of mice, so it is impossible to monitor the physiological status of the NMJ in living mice in real-time.
Soleus and tibialis anterior muscles are acceptable for this protocol; however, they are relatively small, making it challenging during cryosectioning and difficult to visualize a sufficient number of NMJs, especially in the soleus. The best way to visualize NMJs in these muscles is to perform a whole-mount immunofluorescence experiment. This method does not require cryosectioning but involves extensive steps.
The signal intensities of NMJs are an unfavorable parameter for researchers to analyze, as signal intensity varies due to the three-dimensional conformation.
Troubleshooting
Cause Solution
Tissue damage during dissection Excessive force or improper handling. Practice gentle handling techniques and keep tissues moist with PBS during dissection.
Tissue cracking while cryosectioning Inappropriate freezing or blade dullness.
a. Use less liquid nitrogen to avoid flooding the mold.
b. Use a new blade for cryosectioning.
Weak or no staining signal Insufficient concentration of α-BTX applied.
a. Increase the concentration of α-BTX applied.
b. When staining with antibodies for neurofilament or synaptophysin, permeabilize tissues with 0.1% Triton X-100 in PBS before the quenching procedure.
c. Increase the detector gain during the scanning procedure of confocal microscopy.
Acknowledgments
This work was supported by National Science and Technology Council [NSTC 112-2320-B-002-004, NSTC 112-2320-B-002-009]. The mice were bred in the Transgenic Mouse Core Facility in National Taiwan University Center for Genomic Medicine. We also express our gratitude to the Imaging Core at the First Core Labs, National Taiwan University College of Medicine, for their technical assistance in image acquisition and analysis; and Dr. Hsin-Hsiung Chen for his advice for the techniques. This protocol was derived from our previous work [4].
Competing interests
The authors declare no competing financial interests.
References
Tsai, L., Chen, I., Chao, C., Hsueh, H., Chen, H., Huang, Y., Weng, R., Lai, T., Tsai, Y., Tsao, Y., et al. (2021). Autoantibody of NRIP, a novel AChR‐interacting protein, plays a detrimental role in myasthenia gravis. J Cachexia Sarcopenia Muscle. 12(3): 665–676.
Rodríguez Cruz, P. M., Cossins, J., Beeson, D. and Vincent, A. (2020). The Neuromuscular Junction in Health and Disease: Molecular Mechanisms Governing Synaptic Formation and Homeostasis. Front Mol Neurosci. 13: e610964.
Ang, S. T., Crombie, E. M., Dong, H., Tan, K. T., Hernando, A., Yu, D., Adamson, S., Kim, S., Withers, D. J., Huang, H., et al. (2022). Muscle 4EBP1 activation modifies the structure and function of the neuromuscular junction in mice. Nat Commun. 13(1): 7792.
Chen, H., Tsai, L., Liao, K., Wu, T., Huang, Y., Huang, Y., Chang, S., Wang, P., Tsao, Y., Chen, S., et al. (2018). Muscle‐restricted nuclear receptor interaction protein knockout causes motor neuron degeneration through down‐regulation of myogenin at the neuromuscular junction. J Cachexia Sarcopenia Muscle. 9(4): 771–785.
Weatherbee, S. D., Anderson, K. V. and Niswander, L. A. (2006). LDL-receptor-related protein 4 is crucial for formation of the neuromuscular junction. Development. 133(24): 4993–5000.
Wu, H., Lu, Y., Shen, C., Patel, N., Gan, L., Xiong, W. C. and Mei, L. (2012). Distinct Roles of Muscle and Motoneuron LRP4 in Neuromuscular Junction Formation. Neuron. 75(1): 94–107.
Kosco, E. D., Jing, H., Chen, P., Xiong, W. C., Samuels, I. S. and Mei, L. (2022). DOK7 Promotes NMJ Regeneration After Nerve Injury. Mol Neurobiol. 60(3): 1453–1464.
Mejia Maza, A., Jarvis, S., Lee, W. C., Cunningham, T. J., Schiavo, G., Secrier, M., Fratta, P., Sleigh, J. N., Fisher, E. M. C., Sudre, C. H., et al. (2021). NMJ-Analyser identifies subtle early changes in mouse models of neuromuscular disease. Sci Rep. 11(1): 12251.
Wong, M. and Martin, L. J. (2010). Skeletal muscle-restricted expression of human SOD1 causes motor neuron degeneration in transgenic mice. Hum Mol Genet. 19(11): 2284–2302.
Article Information
Publication history
Received: Apr 30, 2024
Accepted: Aug 11, 2024
Available online: Sep 12, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Basic technology > Tissue dissection
Neuroscience > Neuroanatomy and circuitry > Fluorescence imaging
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5,077 | https://bio-protocol.org/en/bpdetail?id=5077&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Construction and Application of a Static Magnetic Field Exposure Apparatus for Biological Research in Aqueous Model Systems and Cell Culture
JV Jana Vučković
HG Hakki Gurhan
BG Belen Gutierrez
JG Jose Guerra
LK Luke J. Kinsey
IN Iris Nava
AF Ashley Fitzpatrick
FB Frank S. Barnes
KT Kelly Ai-Sun Tseng
Wendy S. Beane
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5077 Views: 330
Reviewed by: Willy R Carrasquel-UrsulaezYoungchan KimSusovan Chowdhury
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Original Research Article:
The authors used this protocol in Science Advances Jan 2019
Abstract
With the growth of the quantum biology field, the study of magnetic field (MF) effects on biological processes and their potential therapeutic applications has attracted much attention. However, most biologists lack the experience needed to construct an MF exposure apparatus on their own, no consensus standard exists for exposure methods, and protocols for model organisms are sorely lacking. We aim to provide those interested in entering the field with the ability to investigate static MF effects in their own research. This protocol covers how to design, build, calibrate, and operate a static MF exposure chamber (MagShield apparatus), with instructions on how to modify parameters to other specific needs. The MagShield apparatus is constructed of mu-metal (which blocks external MFs), allowing for the generation of experimentally controlled MFs via 3-axial Helmholtz coils. Precise manipulation of static field strengths across a physiologically relevant range is possible: nT hypomagnetic fields, μT to < 1 mT weak MFs, and moderate MFs of several mT. An integrated mu-metal partition enables different control and experimental field strengths to run simultaneously. We demonstrate (with example results) how to use the MagShield apparatus with Xenopus, planarians, and fibroblast/fibrosarcoma cell lines, discussing the modifications needed for cell culture systems; however, the apparatus is easily adaptable to zebrafish, C. elegans, and 3D organoids. The operational methodology provided ensures uniform and reproducible results, affording the means for rigorous examination of static MF effects. Thus, this protocol is a valuable resource for investigators seeking to explore the intricate interplay between MFs and living organisms.
Key features
• A comprehensive roadmap, suitable for undergraduate to advanced researchers, to construct an apparatus for in vitro and in vivo experiments within uniform static magnetic fields.
• Designed to fit inside standard incubators to accommodate specific environmental conditions, such as with cell culture, in addition to stand-alone operation at room temperature.
• Requires two DC power supplies and 3D printer access for the Helmholtz coils, Plexiglass and mu-metal foil for the partition, and a milli/Gaussmeter for calibration.
• Requires ordering a custom mu-metal shell from a commercial resource (using provided schematics), where lead times for delivery can vary from 2 to 4 months.
Keywords: Quantum biology Magnetic field exposure Xenopus laevis Planaria HT-1080 fibrosarcoma cells Magnetic field manipulation Static weak magnetic fields Tissue growth Regeneration
Graphical overview
MagShield apparatus. This protocol offers a detailed guide on the construction and use of a static magnetic field exposure chamber (top right) for the investigation of magnetic field effects on biological processes. It consists of a mu-metal enclosure (top left), the design and calibration of 3-axial Helmholtz coils (top middle), and considerations for its use with aquatic (Xenopus embryos and larvae and adult planarians), cell culture model systems, and near-zero fields.
Background
Life evolved within the Earth’s natural geomagnetic field, which varies from 25 to 65 μT. However, in the modern environment, anthropogenic magnetic fields (MFs) generated by electronic devices and high-voltage power lines are now ubiquitous [1,2]. Exposures range from very low (e.g., 0.1–3 μT) for household appliances to extremely high (from 1.5 to 7 T) for clinical magnetic resonance imaging (MRI) devices [3–7]. With emerging technologies, new avenues for human exposure to non-ionizing radiation are continually introduced [8,9]. Additionally, the number of diagnostic and therapeutic uses for MFs is growing. The widely used MRI diagnostic relies on static MFs to provide detailed anatomical and physiological information [6,7]. Therapeutically, electromagnetic fields (EMFs) are used for pain management during rehabilitation and with musculoskeletal diseases such as neuropathy and fibromyalgia [10–14]. Research has demonstrated that exposure to even weak MFs can affect biological systems by altering free radical formation—believed to result from MF interactions with spin dynamics [15–21]. Weak MFs have been shown to change reactive oxygen species (ROS) signaling and alter cellular outcomes in vivo [19,22], and thus they represent a potential means to control host defense/immunological responses, regenerative stem cell proliferation, and cancer progression [23–28]. Despite increased interest in uncovering the underlying mechanisms, standardized methods for experimental exposure are lacking and few model systems have been used for in vivo studies (for a recent review, see [29]).
Using this protocol, the authors (a collaboration of engineers, developmental biologists, and geneticists) have successfully investigated MF effects in several different model systems. In vitro, we showed that exposure to static MFs of 0.5 μT and 600 μT MFs inhibited HT-1080 fibrosarcoma cell growth in culture, while 300 and 400 μT increased growth [16,17]. In vivo, we demonstrated that static weak MF exposure of regenerating planarians was able to manipulate stem cell proliferation and gene expression (where 200 μT decreased and 200 μT increased new tissue growth) via changes in superoxide accumulation after injury [19,22]. Our current efforts aim to investigate hypomagnetic field (nT) effects in the vertebrate model organism, Xenopus laevis. The use of a single standardized exposure protocol increases the ease of mechanistic comparisons across organisms and prevents confounding effects from the variance in MFs arising from incubators and standard laboratory equipment (hypothesized to contribute to experimental reproducibility issues even in non-MF experiments, which themselves could benefit from this type of controlled environment). This protocol provides step-by-step instructions on assembling a single 16 (w) × 16 (h) × 16 (d) in (40.64 cm × 40.64 cm × 40.64 cm) mu-metal MagShield apparatus for static MF exposure assays using 60 mm Petri dishes or 96-well plates, with a partition and two 3-axial Helmholtz coils to run controls and experiments concurrently. We focus here on the room temperature aquatic model systems Xenopus and planarians, as well as temperature-sensitive cell culture requiring placement in standard CO2 incubators with internal dimensions of 18.5 (w) × 23.9 (h) × 22.7 (d) in (46.99 cm × 60.71 cm × 57.66 cm). However, the apparatus could be easily adapted to other model systems (including zebrafish, C. elegans, Drosophila, and organoids), as well as extremely low–frequency EMF exposure.
Materials and reagents
Mu-metal enclosure (to block external MFs), ordered commercially using provided schematics (Mu-Shield Company, custom order). Enclosure inner dimensions: 16 (w) × 16 (d) × 16 (h) in (40.64 cm × 40.64 cm × 40.64 cm). Mu-metal thickness: 0.125 in (3.175 mm). See also Notes 1 and 2
Mu-metal partition (in order to run controls and experiments at different field strengths simultaneously):
Plexiglass sheet (Marketing Holders, ASIN: B08G5DD77N); dimensions: 16 (w) × 16 (h) in (same size as the side of mu-metal enclosure); sheet thickness: 1/8 in (3.175 mm)
Mu-metal foil with adhesive (PST on 1-side) (Magnetic Shield Corporation, item number: MUT002-8); foil length: 6 ft (1.8288 m); foil width: 8 in (20.32 cm) (need a total of 16 in wide but not sold wider than 8 in); foil thickness: 0.002 in (0.051 mm)
Two 3D printed Helmholtz coil base frames (STL file of 3D model provided as Supplemental File S1). Base frames were printed with 2.85 mm diameter PLA filament using an Ultimaker S3 printer with a 0.25 mm nozzle. Base frame outer dimensions: 14 cm × 14 cm × 14 cm. Distance between zip-tie anchors: 7 cm
Enameled copper wire for electrical applications, 17 AWG/1 lb (Emtel, model: EMTHERM200, ASIN: B08NW3YVL3). Wire length: 161 ft (49.0728 m). Wire thickness: 17 AWG diameter (1.15 mm)
Power leads (banana plug to alligator clip), 4 black and 4 red (for connecting the power supplies to the Helmholtz coils) 14 AWG (Zhenyu, item model number: ZY12091, ASIN: B0BPL4C29T). Lead silicone outside diameter: 3.5 mm. Lead length: 6 ft
Grounding lead (banana plug to alligator clip), 1 green, 18 AWG (DigiKey, Pomona Electronics, part number: 501-2514-ND, product number: EM1166-24-5#). Lead length: 2 ft
Note: The grounding lead does not have to be green, although we highly recommend that you use a separate color from the red/black power leads to clearly delineate your grounding lead.
Optional: For hypomagnetic field exposure, two small corrugated cardboard shipping boxes, 4 in × 4 in × 4 in (10.16 cm in all dimensions) for secondary mu-metal shielding (SUNLPH, ASIN: B09QCXPXY1)
Laboratory supplies
Safety glasses
Gloves
Scissors/metal shears
Ruler or tape measure
Zip-ties (wire management for Helmholtz coils)
Electrical tape (to secure lead connections)
Tape/adhesive (duct tape for the partition and laboratory labeling tape for wires is recommended)
Wire strippers
Wire cutters (if not already a function included with the wire strippers)
Sandpaper (may be needed when fitting Plexiglass partition)
Empty welled plates/Petri dishes (for positioning samples in the exact center of the Helmholtz coils)
Note: The width of the plasticware matters as they must fit in between the two rows of a Helmholtz coil pair (~6 cm). We recommend recycling used plasticware for positioning, such as 6/12/24-welled plates or 60 mm Petri dishes.
Empty pipette tip boxes/welled plates (for positioning the Helmholtz coils in the center of each chamber)
Notes:
The size of the plasticware does not matter for positioning the Helmholtz coils.
We recommend recycling used plasticware for positioning.
Optional: For cell culture/incubator placement, polytetrafluoroethylene (PTFE) tape (such as Teflon tape)
Equipment
Two DC power supplies, 300 W dual output, 30V/5A 30V/5A (Jameco, Mastech, part number: 301947, model: HY3005D-2-R)
Note: The dual output power supply is suitable for Helmholtz coils using x- and y-axes only. To use the z-axis as well, a triple output power supply (such as Mastech HY3005D-3-R, Jameco, part number: 301955) should be used.
DC milli/Gaussmeter (for experiments at 0–199 μT) (AlphaLab, Inc., model: MGM, sensor alignment: standard, bandwidth: 3 Hz)
DC Gaussmeter (for experiments at 200–1,000 μT) (AlphaLab, Inc., model: GM1-HS)
Procedure
This protocol requires a “pre-step” (Section A) of ordering the mu-metal enclosure. While waiting for this to arrive, the Helmholtz coils can be constructed (Section C), although this is not required. Once the mu-metal enclosure has arrived, the partition may be added (Section B). After all this is complete (Sections A–C), the MagShield apparatus can be assembled and calibrated (Section D) prior to use (Section E). The instructions are purposely comprehensive, despite the comparatively straightforward nature of constructing the apparatus. In particular, attention has been focused on providing explanations for biologists lacking engineering/physics backgrounds. Once all materials needed have been obtained, the entire apparatus can be constructed in a single day if desired. As an example, the hypomagnetic apparatus shown below was constructed by a team of mostly undergraduate students (with graduate student participation), and the initial test experiments began at the end of the day.
Critical: Read through the entire protocol before beginning.
Ordering the mu-metal enclosure (Figures 1-2 and Supplemental File S2)
This protocol provides schematics (Figure 1 and Supplemental File S2) for constructing a 16 × 16 × 16 in enclosure (inside measurements) using mu-metal of 0.125 in thickness. The enclosure's features include accessibility ports, a latch system for the door, and support feet for stability. Most biology labs lack the necessary equipment for welding, and thus it is recommended to order the enclosure from a commercial company. See Notes 1 and 2.
Figure 1. Schematic for MagShield apparatus enclosure. Schematic diagrams for constructing the 16 × 16 × 16 in enclosure out of 0.125 in thickness mu-metal. See Supplemental File S2 for complete details.
We custom ordered the mu-metal enclosure (Figure 2) from The MuShield Company (Londonderry, NH) according to the provided schematics (although any company with the same capabilities will work).
Note: Mu-metal blocks external MFs, allowing researchers to precisely control experimental conditions.
Critical: Please note that construction times can be lengthy; for our supplier, the average delivery time was 12 weeks.
Figure 2. Mu-metal enclosure. As unpacked directly from the supplier, before the addition of the partition. (A) Inside. (B) Front outside. Note the door latches (asterisks), access ports at the top (arrows), and the access ports at the bottom (arrowheads).
Adding the mu-metal partition (Figures 3–4)
It is important to always run a control concurrent with the experiment. Variations in local MF exposure of organisms prior to the assay could result in differences in experimental outcomes (and have in our experience). To run experiments simultaneously, a mu-metal-covered partition is created in the center of the mu-metal enclosure to produce two chambers that can have different MF strengths.
Note: You need the mu-metal enclosure (Section A) in order to complete this step.
Double-check the inner dimensions of the mu-metal enclosure (Figure 2A) and the dimensions of the Plexiglass sheet (Figure 3B–C), as dimensions may vary slightly during manufacturing. All dimensions should be 16 in (except thickness).
Notes:
The thickness of the Plexiglass is only required for the stability of the partition.
If the inner enclosure dimensions are not exactly 16 in, a Plexiglass sheet of the exact dimensions will be required. If the inner enclosure dimensions are only slightly smaller than 16 in, sandpaper may be used to reduce the Plexiglass sheet to the exact dimensions.
Caution: Plexiglass that is slightly (1 mm) larger in overall dimensions than the enclosure’s inner dimensions is preferable to Plexiglass that is slightly smaller. If the Plexiglass is larger, it can be sanded down to fit exactly, while Plexiglass that is smaller should not be used (as it will leave gaps in the shielding between the two chambers).
Figure 3. Constructing the mu-metal partition. (A) Mu-metal foil showing a corner where the adhesive packing is peeled off (arrow). (B) Plexiglass sheet. (C) View of Plexiglass sheet thickness 1/8 in (3.175 mm). (D) Cutting the mu-metal foil with adhesive backing into 16 in lengths. (E) First 16 in strip of foil attached to half of the Plexiglass sheet.
Cut the mu-metal foil (Figure 3A) by laying it out on a flat surface. Using a ruler/tape measure and a pencil, mark 16 in lengths on the mu-metal foil. Cut the foil along the marked lines using scissors or metal shears (Figure 3D). One partition should require four 16 in lengths (each 8 in wide), two for each side. Be precise as mu-metal foil is delicate and can easily become damaged.
Caution: You may wish to wear gloves to protect yourself from sharp edges.
Place the Plexiglass sheet on a flat surface and confirm mu-metal foil strips are the correct length. Peel off the backing paper to reveal the adhesive on the foil and cover the Plexiglass fully on both sides (Figure 3E). Cover any sharp edges from the foil with tape.
Notes:
If the edges of the Plexiglass are not smooth, sand them before attaching mu-metal foil.
If the mu-metal foil gets bent, the creases will be permanent. It can still be used, but then the dimensions will be off, and you will require more foil to cover the partition. While tiny, minor creases may be unavoidable, major creases as seen in Figure 3E should be avoided if possible.
Critical: The mu-metal foil must cover the entire surface of both sides, as any uncovered area will allow bleed through of the MF from one chamber into the other.
Place the foil-covered mu-metal partition in the middle of the enclosure so that it bisects the enclosure, making two equal-sized chambers (Figure 4A). Make sure the partition is securely in place.
Note: The fit should be tight enough that the partition will stay in place without being held.
Figure 4. Installing the mu-metal partition. (A) Correct placement (arrow) of the foil-covered Plexiglass sheet, bisecting the mu-metal enclosure. (B) Taping the partition to the enclosure for extra stability. (C) Final installation of the mu-metal partition, showing the tape (arrowheads) connecting the partition to the enclosure.
For extra security, use tape (such as duct tape) to attach the partition to the enclosure at the top and bottom of both sides (Figure 4B–C).
Critical: It is important to avoid any gaps between the partition and the enclosure to ensure complete shielding between the two chambers.
Notes:
You will need to confirm that the MF generated in one chamber is not influencing the field strength in the other chamber. See step D9.
If there are no gaps but the partition does not provide full shielding between the two chambers (see section D), thicker mu-metal or more layers should be used. However, as a single layer of 3.175 mm mu-metal provides more than 99% attenuation, additional layers should not be necessary. See also Notes 1c and 6.
Constructing the Helmholtz coils (Figures 5–6 and Supplemental Files S1 and S3)
Download and print the 3D model (Figure 5A and Supplemental File S1) of the base frames for the 3-axial Helmholtz coils. You will need to print two bases (one for controls and one for experiments). The provided STL file produces a base frame of 14 (w) × 14 (d) × 14 (h) cm, which is suitable for assays using a single 60 mm (or 35 mm) Petri dish or 96-well plate.
Note: The breakaway support structures connecting the zip-tie anchors are shown still attached in Figure 5B and removed in Figure 5C. Make sure these supports are removed prior to winding.
Figure 5. Helmholtz coil anatomy. (A) 3D model of the base frame. (B) Actual 3D printed base frame, with breakaway supports (arrowheads) still attached. (C) Close-up of the 3D printed base frame with breakaway supports removed. Note the raised zip-tie anchors (arrows) for correct positioning of coil rows and wire management. (D) Diagram of 3-axial Helmholtz coil showing the three coil pairs for the three axes (x in blue, y in red, and z in grey).
Note: Only two axes (x- and y-axis) are required for this protocol.
The base frame, made from non-conducting material [polylactic acid (PLA) filament], allows for three Helmholtz coil pairs (one pair each for the x, y, and z axes, Figure 5D). For this protocol, only the x and y pairs are used. Each coil pair is comprised of two rows (left and right) on the same axis (arrows in Figure 5D) that are positioned at a specific distance apart in parallel orientation. The 3D model provided includes zip-tie anchors (Figure 5C) for both correct positioning of coil rows and wire management. For this size base frame, the correct distance between anchors is 7 cm. Each face/side of the base frame has eight zip-tie anchors: four for each axis (the axes are perpendicular to each other).
Critical: The space between the two rows of each coil pair must be separated by a distance (h) equal to the radius (R) of the coil.
Note: The zip-tie anchors in the provided 3D model file are positioned exactly at this distance.
Wind one coil pair onto the first side (axis) of one base frame.
Temporarily secure the copper wire to the base frame, centered over the appropriate zip-tie, with a small piece of tape/adhesive. Leave approximately 4 in (~10 cm) of wire free at the beginning for attaching to the power supply leads (arrow in Figure 6A).
Critical: Make sure to always keep your wire centered over the zip-tie anchor so that the final distance of the two rows will remain the correct distance apart.
Note: It does not matter whether you start with the left or right row of the coil pair.
Wrap the copper wire completely around the base frame for one full revolution, until you are back where you started. This is considered one winding. Continue until you reach a total of 50 windings for the first row of that coil pair.
Critical: Make sure your wire stays centered over the appropriate zip-tie anchors on each face of the base frame.
Cross your wire to the other side of the coil pair (asterisk in Figure 6B) to begin the second row/coil of that pair. Both rows of each coil pair are made from one continuous length of copper wire.
Critical: The winding for the second coil needs to follow the same direction of winding as the previous one.
Continue until you reach 50 windings for the second row of that coil pair.
Secure the end of the wire to the frame temporarily with a small piece of tape/adhesive to keep it in place. Cut the wire, leaving approximately 4 in (~10 cm) of wire free at the end (in addition to the 50 windings) to attach to the power supply leads (Figure 6A).
Note: The tape secures the wire until all coil pairs for that base frame are finished. Do not attach with zip-ties (as seen in Figure 6) until all coil pairs are wound.
Critical: Make sure that, when finished, the two rows of the coil pair are parallel to each other and centered over their respective zip-tie anchors.
Repeat steps C3a–e for the two other coil pairs on that same base frame.
Note: Only two perpendicular axes (x- and y-axis) are needed for this protocol. Winding the third coil pair for the z-axis is optional.
Use wire strippers to remove the insulation from both ends of the copper wire (arrowhead in Figure 6C) of each coil pair. Make sure the wire ends are clean and exposed.
Secure each coil row to the base frame using zip-ties through each zip-tie anchor, cutting off the unused “tail” of each zip-tie (Figure 6D) and removing any tape that was temporarily holding the wire to the frame.
Note: We recommend that on the two “open” faces of the base frame, you zip-tie the middle of each coil row (as seen at the top of Figure 6D) for stability and to ensure proper spacing is maintained.
Repeat the entire process (C3a–h) for the other base frame.
Figure 6. Winding the coils. (A) Free end of the copper wire (arrow) for one coil, where 4 in (~10 cm) of wire is left unwound at each end for attaching to the power supply. (B) View of one face of a completed coil, showing the crossing of the copper wire (asterisk) between the left and right coil pairs. (C) Free end of one coil, showing the stripped insulation (arrowhead) required for a strong connection to the power supply lead. (D) Completed Helmholtz coil with three coil pairs (x, y, and z) that have been zip-tied (white clasps) to the base frame via the anchors and had the zip-tie “tails” cut off.
Assembly and calibration of the MagShield apparatus (Figures 7–9)
Place the mu-metal enclosure on a stable, level surface that also has enough room for the power supplies.
Note: This should be the final placement for the apparatus.
Position the power supplies to one side of the enclosure. Each power supply will provide current to a single Helmholtz coil (either the control or the experimental). A single power supply connects to one Helmholtz coil via four leads: one positive (red) and one negative (black) lead will connect to each coil pair on both x and y axes.
Connect the leads to the Helmholtz coils (Figure 7).
Caution: Steps D3–7 should be done without power to the Helmholtz coil to ensure safety.
Take one Helmholtz coil and attach the alligator clip of the red lead to one of the exposed wire ends of a coil pair (arrow in Figure 7A). Attach the black lead to the other end of the wire of that same coil pair (which will be on the other row).
Note: Each coil pair forms a circuit with one positive and one negative input.
Secure the alligator clips to each wire end using electrical tape (arrowhead in Figure 7A).
Critical: Ensure the connections are secure and insulated to prevent short circuits.
Tape the red and black leads together (for lead management/organization) as shown in Figure 7D. Label the banana plug ends of each lead as either x- or y-axis based on that coil pair’s orientation (refer to the diagram in Figure 5D). General laboratory labeling tape can be used.
With another set of red/black leads, repeat steps D3a–c for the remaining coil pair on that same Helmholtz coil.
Repeat the entire process (D3a–d) for the other Helmholtz coil.
Figure 7. Placement of the coils, samples, and leads. (A) The alligator clip end of a red lead is attached to the exposed end of one coil pair (arrow). The alligator clip of a black lead is attached to the other end of that same wire (middle of image). To ensure a good connection, secure the clip to the wire with electrical tape (arrowhead). (B) A taped stack of used pipette tip boxes is used to position the Helmholtz coil in the center of each partition. (C) A complete Helmholtz coil with a stack of used plasticware (welled plates) to position the samples (Petri dish) in the center of the coil base frame. (D) Final positioning of both Helmholtz coils, one in each partition. Note in the lower-right corner where a pair of red and black leads clipped to each end of a single coil pair have been taped together (green tape).
Position the Helmholtz coils in the enclosure.
In each chamber, create a platform using a stack of empty plastic pipette tip boxes or non-conductive welled plates that will position one Helmholtz coil exactly in the center of that chamber (Figure 7B).
When placed on the platform, the center of the Helmholtz coil should correspond to the center of the chamber. This is required for the uniformity of the MF exposure.
Tape the final stack of tip boxes/plates together for platform stability.
Note: Alternately, a separate stage using a non-conducting material such as cardboard or plastic can be designed if more stability for the Helmholtz coil is needed/desired.
Connect the Helmholtz coils to the power supplies (Figure 8). Ensure the power supplies are turned off and unplugged before starting the process.
After each Helmholtz coil (wound base frame) is placed on its platform, feed all the leads from one Helmholtz coil through the most convenient access port. Leads can be fed through either the top (Figure 8A) or bottom (Figure 8B–C) ports.
Critical: It is imperative to ensure that the Helmholtz coils themselves do not make contact with the mu-metal of the enclosure's sides.
The four leads from a single Helmholtz coil connect to a single power supply. Each power supply has two groups of terminals, one group for each axis of a single Helmholtz coil.
Note: If using the optional z-axis, there will be six leads from each Helmholtz coil that will all connect to a single power supply with three terminals.
Identify one set of positive (red) and negative (black) terminals on one power supply and attach the corresponding colored leads (via the banana plugs) from one axis of one Helmholtz coil (Figure 8D).
Attach the other set of leads for the remaining axis on that base frame to the other set of terminals on that same power supply.
Repeat steps D5a–d with the other power supply and the remaining Helmholtz coil (wound base frame).
Figure 8. Connecting to the power source. (A) Leads from a Helmholtz coil fed through an upper access port (arrowhead). (B–C) Views of a set of leads fed through a lower access port (yellow arrows), shown from the outside (B) and the inside (C) of the enclosure. (D) Power supplies with attached leads. Red arrow shows axis labeling of each lead. (E) Alligator clip end of ground lead (asterisk) attached to an upper port on the enclosure. (F) Probe end of a milli/Gaussmeter, taped to the stage in the correct orientation for that make of probe, during a test of field strength.
Ground the MagShield apparatus by attaching the green lead to any of the green terminals on either power supply via the banana plug end and then attaching the alligator clip to one of the unused access ports on the mu-metal enclosure (asterisk in Figure 8E). Only one ground lead is required.
Create two raised stages using stacks of empty Petri dishes or welled plates that will position your samples exactly in the center of each Helmholtz coil (Figure 7C). The raised stage can be taped together for stability.
Calibrate the MagShield apparatus.
Plug in and turn on the power supplies and determine your working voltage and current settings by using the milli/Gaussmeter to measure the MF produced at the center of the Helmholtz coil. Tape the probe end to the stage for accurate readings (Figure 8F).
Caution: Monitor the power supplies for any unusual behavior or overheating. If any issues arise, turn off the power supply immediately.
Critical: The orientation of the meter’s probe is crucial for an accurate reading, as the meter will still display an (incorrect) reading in the wrong orientation. Consult the user manual of the milli/Gaussmeter for guidance on proper usage for your meter.
Critical: If your meter is not correctly calibrated, then your MagShield apparatus will not be either. Make sure that your milli/Gaussmeter has a current calibration record/certificate and that routine maintenance, including recalibration, is performed as directed by the manufacturer. Most manufacturers recommend at least annual meter calibration.
The first step when measuring the field strength of the coils needs to be a measurement of each coil axis separately, while the other coils are not connected to the power supply. For example, supply power to the x-axis coil pair only and record the field strength produced.
Note: The probe should remain taped to the stage at the center of the Helmholtz coil (where your experiments will be placed) for all field strength measurements.
Change the power supply settings (according to manufacture guidelines) until you record a field strength at (or as close as possible) to your desired MF strength.
Repeat steps D8b–c for the other axis coil pair (such as the y-axis).
Critical: The orientation of the meter’s probe will need to be changed with each different axis; refer to your specific probe’s user manual for proper orientation.
Note: If using all three axes, repeat for the y-axis coil pair as well.
After initial measurements of each individual coil axis, measure the field strength produced by the Helmholtz coil (wound base frame) by supplying power to all axes simultaneously. Measure the field strength recorded when the probe is in each orientation specific to each axis. For example, with power to both x- and y-axes, record the MF strength with the probe in both the x and then y orientations. There should not be any significant differences; however, it is important to confirm the field strength when all the coils are connected.
Change the power supply settings for each axis until you record your desired field strength for the entire Helmholtz coil (i.e., with power to all axes).
Repeat steps D8a–f for both your control and your experimental chambers until you record your desired MF strength. Document your specific power supply settings for future reference.
To confirm that the MF generated in one chamber is not influencing the field strength generated in the other chamber, take measurements with the milli/Gaussmeter of the ambient field strength in one chamber (without power) while simultaneously increasing the field strength in the other chamber (with power). There should be no fluctuations of field strength recorded, indicating a lack of bleed through.
Once the MagShield apparatus is completed and calibrated (Figure 9 and Supplemental File S3), we recommend performing an initial trial run of both Helmholtz coils at the desired field strengths without samples lasting for the total time of your assay. During this trial run, multiple field strength and temperature checks should be conducted (at the center of each Helmholtz coil) to ensure that environmental and exposure conditions remain stable.
Figure 9. Completed MagShield apparatus. Shown with enclosure safety latches securing the door shut and power supplies positioned next to the mu-metal enclosure so that the leads can reach the coils inside.
General basic operation of the MagShield apparatus
Note: These general instructions are for room-temperature aquatic model systems where the organisms are free swimming/moving (such as planarians). This requires the use of both the x-axis and y-axis coil pairs to ensure a uniform field regardless of the animal’s position in the dish.
Inspect the wires, leads, and connections, to ensure they are in good condition and free from damage/wear.
Carefully turn on the power supplies, ensuring that the output is at your predetermined settings (see Section D).
Caution: Monitor the power supplies for any unusual behavior or overheating. If any issues arise, turn off the power supply immediately.
Critical: Ensure that there are no loose connections, exposed wires, or other safety hazards.
Use the milli/Gaussmeter to confirm your field strengths at the beginning of each experiment for both coils. Tape the probe end to the top of the stage to get accurate measurements.
Critical: You should repeat all steps of Section D, Step 8 each time you measure field strength.
Note: Controls are typically run at or near 45 μT, as Earth normally averages 25–65 μT. It is recommended to designate one power supply/Helmholtz coil set as your control set.
Watch for any changes in current/voltage in the first few minutes, ensuring that they remain stable.
Note: Fluctuations in current/voltage suggest a loose connection.
Place your samples on the Helmholtz coil stage. Samples will only be exposed to a uniform field if they are placed in the center of the Helmholtz coil. There are limits to the number of samples that can be accommodated. Only one 60 mm Petri dish or 96-well plate should be placed in each chamber.
Caution: The power supply should be disconnected and off for the safety of the person placing the sample in the center of the Helmholtz coil.
Critical: Do not place samples in the outer wells of a 96-well plate, as their field strength exposure will not be comparable to those in the center.
At the end of each experiment, always confirm with the milli/Gaussmeter that the field strengths of each Helmholtz coil have remained the same.
Critical: Trials where field strengths did not remain constant should not be used and suggest a problem with the power supply or the connections between the power supply and the coils (the leads).
Make sure to turn off both power supplies at the end of each experiment.
Caution: Always exercise caution when working with electrical equipment and follow safety guidelines and manufacturer's instructions.
Modifications for cell culture (Figure 10)
Note: These modifications are for environmentally controlled model systems where the MagShield apparatus needs to be housed inside an incubator. For organisms that are not freely swimming, such as cell culture, the use of a single axis is required. In our experiments with HT-1080 cells, we observed significant effects on cell growth and mitochondrial calcium levels when the MFs were perpendicular to the cell culture flask compared to when they were parallel (see Validation, Section B). As a result, we chose to only utilize the y-axis of the Helmholtz coils in some of our experiments.
The provided mu-metal enclosure schematics are designed so that the MagShield apparatus will fit inside standard CO2 incubators with internal dimensions of 18.5 (w) × 23.9 (h) × 22.7 (d) in (Figure 10). Most standard incubators come with ports for access to the outside.
Figure 10. Modified MagShield setup for cell culture. View of MagShield apparatus inside a carbon dioxide (CO2) incubator used for cell culture, as previously reported [17]. This modification is required to maintain specific temperature, humidity, and gas composition needed to support the growth of cells in culture. Modifications include additional environmental sensors and considerations to prevent gas leaks while accommodating lead access to the power supplies.
The length of the leads may need to be changed so that they can extend through the incubator port to the power supplies to the Helmholtz coils. We used test leads with banana-to-banana connectors that are 60 in long to extend through the incubator port to connect to the power supplies for the Helmholtz coils.
Cell culture experiments will typically require the use of different sensors to monitor the environmental conditions inside the MagShield apparatus.
For example, it is essential for cell culture experiments that there is a CO2 monitor to verify proper air circulation (even though vents are designed into the enclosure).
This protocol is adaptable to accommodate various sensor types (temperature, CO2, humidity, etc.).
We addressed the need for cable access by opening a hole in the silicon plug of the port, which was sized precisely to accommodate the leads. Furthermore, we took precautionary measures by wrapping the leads with Teflon tape to prevent any gas leaks from the incubator. This approach ensured that the leads could enter the incubator without compromising its integrity, thereby allowing for the safe and effective operation of the MagShield apparatus within the incubator environment.
Modifications for hypomagnetic fields (Figure 11 and Supplemental File S4)
Note: These modifications are for experiments testing the effects of near-zero MF exposure. An additional mu-metal-covered container is required.
For experiments in hypomagnetic, “near-zero” fields (< ~10 μT), increased mu-metal protection from surrounding MFs is required, necessitating the use of an additional mu-metal-covered container inside the MagShield apparatus for experiments.
Only a single power supply and Helmholtz coil (wound base frame) is required for near-zero experiments, which is used for controls.
Note: It is possible to run near-zero assays without the use of any Helmholtz coils, where controls are placed in “normal” environmental conditions.
Caution: Due to the high variability of environmental conditions, we strongly recommend the use of the MagShield apparatus to shield external MF and a Helmholtz coil to generate an average Earth normal MF (45 µT) for all control conditions.
Cover a 4 in × 4 in × 4 in cardboard shipping box with mu-metal foil for secondary containment (red arrow in Figure 11A).
Critical: Make sure that all surfaces, including the lid, are covered in mu-metal to provide a complete secondary MF barrier.
Caution: The edges of the mu-metal foil are sharp. The sharp edges of the mu-covered container can be covered in tape (Figure 11B) to prevent injuries.
Figure 11. Hypomagnetic field exposure setup. (A) Interior of MagShield apparatus showing a Helmholtz coil in the left partition for Earth-normal controls, and an additional mu-metal-covered container for “near-zero” (< ~10 μT) hypomagnetic fields in the right partition (red arrow). (B) Mu-metal covered near-zero containers shown with tape covering sharp edges. (C) Photo of the hypomagnetic field exposure setup, with a single power supply and Helmholtz coil for controls, and additional mu-metal container for near-zero experiments.
Alternately, since Helmholtz coils are not needed for hypomagnetic experimental conditions, larger secondary containers can be used, such as the 3D-printed secondary containment options shown in Supplemental File S4.
Critical: Make sure that all surfaces of the non-conductive container, including the lid and the inside, are covered in mu-metal foil to provide a complete secondary MF barrier, using tape to cover any sharp edges from the foil.
Place the smaller mu-metal container in the experimental chamber of the MagShield apparatus in place of the experimental Helmholtz coil (Figure 11C).
Using the milli/Gaussmeter, validate that in the experimental chamber the MF is at the desired near-zero field strength inside the smaller mu-metal container (when inside the MagShield apparatus). Then, validate that in the control chamber the control MF inside the Helmholtz coil is also accurate.
Note: Even though no Helmholtz coil is needed, you should still record the field strength in the experimental chamber with the milli/Gaussmeter probe in the correct orientation for all three axes (x, y, and z) to ensure that the field is truly near zero.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Van Huizen, et al. [22]. Weak magnetic fields alter stem cell–mediated growth. Science Advances. 5: eaau7201. doi:10.1126/sciadv.aau7201.
Gurhan et al. [17] Effects induced by a weak static magnetic field of different intensities on HT-1080 fibrosarcoma cells. Bioelectromagnetics 42(3): 212–223. doi: 10.1002/bem.22332.
Gurhan et al. [16]. Impact of weak radiofrequency and static magnetic fields on key signaling molecules, intracellular pH, membrane potential, and cell growth in HT-1080 fibrosarcoma cells. Scientific Reports. 13, 14223. https://doi.org/10.1038/s41598-023-41167-5.
Gurhan et al. [15] Weak radiofrequency field effects on chemical parameters that characterize oxidative stress in human fibrosarcoma and fibroblast cells. Biomolecules. 13(7): 1112. https://doi.org/10.3390/biom13071112.
Kinsey et al. [19]. Weak magnetic fields modulate superoxide to control planarian regeneration. Frontiers in Physics. 10: 1–20. https://doi.org/10.3389/fphy.2022.1086809.
Planarian regeneration experiments (Figure 12)
Amputation scheme (Figure 12A): Previously, we investigated static MF effects on planarian regeneration only for trunk fragments (with heads and tails removed) that have two amputation planes (one above and one below the pharynx/feeding tube) [19,22]. To determine the effects on fragments with a single amputation plane, we investigated three other fragment types. Schmidtea mediterranea (5–7 mm in size) were amputated just above the pharynx to generate both a) head fragments that will undergo posterior regeneration of both pharynx and tail, and b) posterior fragments (which include both the pharynx and tail) that will regenerate a new head. Additionally, we amputated animals just below the pharynx to produce c) tail fragments that will undergo anterior regeneration of both a new head and pharynx.
Figure 12. Effects of static magnetic fields on planarian regeneration. Regenerating fragments analyzed for blastema size (new tissue growth, visualized as the white region at the wound site) after three days of exposure to 500 µT (activating), 200 µT (inhibiting), or 45 µT (control) field strengths, using the MagShield apparatus as described in this protocol at 20 °C (room temperature). (A) Diagram of cuts. Adult planarians were transected by scalpel at one of two different amputation planes (dotted red lines), one above and the other below the pharynx (feeding tube), producing three different fragment types. (B) Head fragments, regenerating their pharynx and tail. (C) Posterior fragments, regenerating their head. (D) Tail fragments, regenerating their pharynx and head. (E) Quantification of B–D. Anterior is up. Single solid arrows: control blastema size. Double solid arrows: increased tissue growth. Empty arrows: inhibited tissue growth. Scale bars = 100 µm. n ≥ 5. Error bars = SEM. * p < 0.05, *** p < 0.001, **** p < 0.0001.
Our previous studies found that weak static MF effects were dependent on field strength, with increased stem cell–mediated regeneration peaking at 500 μT and decreased regeneration peaking at 200 μT. For our three new fragment types, regenerates were exposed to static MFs at either control (45 μT), 200 μT, or 500 μT field strengths. MF exposure occurred from within 5 min of amputation until Day 3 of regeneration, when blastema (new tissue) growth at the wound site was analyzed for all fragment types (Figure 12B–E). Blastema size was measured as a percentage of total regenerate size, to account for worms/fragments of different sizes.
We found that compared to controls, 500 μT static MF exposure resulted in blastemas that were significantly larger for all three fragment types (Figure 12E): head fragments (500 µT, p = 0.0255), posterior fragments (500 µT, p = 0.000975), and tail fragments (500 µT, p = 0.000596). Similarly, our results demonstrated that 200 µT exposure produced regenerates with significantly inhibited regeneration in all fragment types (Figure 12E): head fragments (200 µT, p = 0.000465), posterior fragments (200 µT, p = 9.65E-06), and tail fragments (200 µT, p = 4.727E-05).
Together, these data indicate that similar to fragments with two amputation planes, the regeneration of planarians with a single wound site can also be modulated by weak static MF exposure. Furthermore, MF regulation of stem cell–mediated regeneration, in a field strength-dependent manner, can both increase and decrease new tissue growth regardless of the number of new tissues/organs that must be regenerated. Additionally, these data suggest that MF effects on new tissue growth are consistent regardless of the level of the injury along the anterior-posterior axis.
Statistics and quantification: Specific sample sizes (n) that correspond to the total number of worms across all replicates and the number of independent replicates or trials (N) were as follows. For 45 µT exposures: n = 12, N = 4 for head fragments; n = 31, N = 4 for posterior fragments; and n = 15, N = 2 for tail fragments. For 500 µT exposures: n = 12, N = 2 for head fragments; n = 15, N = 2 for posterior fragments; and n = 10, N = 1 for tail fragments. For 200 µT exposures: n = 13, N = 2 for both head fragments and posterior fragments; and n = 5, N = 1 for tail fragments. The magnetic lasso tool in Photoshop (Adobe) was used to generate total pixel counts of the blastema (white tissue at the wound site) and the total regeneration (entire worm including blastema). Blastema size is reported as a percentage of total body size: (blastema size/body size) × 100. Significance was calculated using a two-tailed Student’s t-test with unequal variance (Microsoft Excel or GraphPad Prism 10), where p < 0.05 was considered statistically significant. Each experimental group was normalized against its corresponding control group, which was maintained at 45 µT (average Earth normal).
Cell culture experiments (Figure 13)
Cell growth (Figure 13A): The impact of static MF exposure on the growth rates of HT-1080 cells fibrosarcoma cells over a four-day period, ranging from 0.5 to 600 µT was analyzed. Notably, growth rates showed an initial increase from 0.5 to 400 µT, followed by a decline at 600 µT. Growth rates at 0.5 and 600 µT were significantly lower by 9% (p < 0.001) compared with the control group at 45 µT (average Earth normal); conversely, at 300 and 400 µT, growth rates exceeded controls by 23% (p < 0.001) and 28% (p < 0.001), respectively.
Figure 13. Effects of static magnetic field intensity on fibrosarcoma cells. HT-1080 cell line exposed to a range of 0.5–600 µT static MFs, with 45 µT as controls (“C” bar on graphs), using the modified MagShield apparatus for cell culture in a 37 °C CO2 incubator. (A) Effects on cell growth. Cell growth rates expressed as a function of static MF exposure (mean ± SD) with the fields perpendicular to the flask bottom; n = 12, N = 3 for each group. (B) Effects on hydrogen peroxide (H2O2) levels. H2O2 concentrations as a function of static MF exposure (mean ± SD); n = 63, N = 3 for each group. (C) Effects on mitochondrial calcium (Ca2+) levels. Mitochondrial Ca2+ concentrations as a function of static MF exposure (mean ± SD) for fields oriented at 90° with respect to the plane of the cell flask bottom; n = 63, N = 3 for each group. * p < 0.05, ** p < 0.01, and *** p < 0.001. Error bars = standard deviation (SD).
Hydrogen peroxide (H2O2) levels (Figure 13B): Amplex Red reagent assay was used to measure the production of H2O2 in HT‐1080 fibrosarcoma cells. Concentrations of H2O2 exhibited a non-linear response corresponding to the intensity of the applied MF. Specifically, at 100, 200, 500, and 600 µT, the concentration of H2O2 surpassed that of the control group significantly (p < 0.001), indicating an increase. Conversely, at 300 µT the concentration was lower than controls (p < 0.05), suggesting a decrease in H2O2 concentration.
Mitochondrial calcium levels (Figure 13C): The Rhod‐2 AM probe was used to determine mitochondrial Ca2+ level in HT‐1080 human fibrosarcoma cells. Exposure to static MFs lead to a notable increase in mitochondrial calcium concentration as MF intensity rose. Significant disparities were evident at 200 µT (p < 0.05) and across the range of 300–600 µT (p < 0.001), as compared with the control group.
Statistics: The total sample count is denoted as “n” (corresponding to the total number of samples across all replicates), while the number of independent replicates or trials is denoted as “N” (see Figure 13 legend). Statistics were conducted using Origin Pro 2017 Statistical Package (OriginLab, Northampton, MA). Differences in cell growth rates and fluorescence were deemed statistically significant when p < 0.05. For comparisons between two independent samples, Student's t-test was used, while one-way analysis of variance (ANOVA) with Tukey post-hoc was used to contrast the means of multiple independent samples. Each experimental group was normalized against its corresponding control group, maintained at 45 µT.
Xenopus hypomagnetic field experiments (Figure 14)
To investigate static MF effects on the development of vertebrate embryos, Xenopus laevis were exposed to either control (45 μT) or hypomagnetic field strengths (< ~10 μT) starting from the two-cell stage.
Two trials were performed as part of our initial runs. For both, “n” corresponds to the total number of animals across all replicates, while “N” is the number of independent replicates.
For trial 1 (n = 28 for control and n = 29 for experimental animals, N = 1 for each), the hypomagnetic field measured from 11 to 12 μT and embryo morphology was assessed at three days of development (Figure 14A). While all control animals displayed normal development, 2/29 experimental animals displayed stunted growth.
Figure 14. Preliminary developmental phenotypes from hypomagnetic field exposure of Xenopus larvae. Embryos exposed to either 45 µT (controls) or near-zero static MFs using the modified MagShield apparatus for hypomagnetic fields at 20 °C (room temperature). (A) Trial run 1: phenotypes at 3 days of development. (A1) Controls, phenotype n = 28/28. (A2-3) Near-zero affected phenotypes at 11–12 μT: Stunted growth with defects in anterior structures and anterior-posterior (AP) axis formation; n = 2/29. (A4) Representative phenotype of 11–12 μT animals with normal appearance. (B) Trial run 2: phenotypes at 5 days of development. (B1) Controls, phenotype n = 30/30. (B2-3) Near-zero affected phenotypes at 2–5 μT: Continued stunted growth with defects in anterior structures and AP axis formation; n = 3/30. (B4) Representative phenotype of 2–5 μT animals with normal appearance. Anterior is to the right and dorsal is up.
For trial 2 (n = 30, N = 1 for both controls and experimental animals), the hypomagnetic field measured from 2–5 μT and embryo morphology was assessed at 5 days of development (Figure 14B). Similar to our previous trial run, all control animals developed normally while 3/30 experimental animals had stunted growth.
These preliminary data are promising and suggest investigation of hypomagnetic fields on vertebrate development is an area for exploration.
General notes and troubleshooting
General notes
Magnetic field shielding
The mu-metal suggested in this protocol is not the only material that can be used to shield experiments from MFs. Any high magnetic permeability material (such as ferromagnetic metals) can be used to redirect surrounding MFs around and away from your experiments.
While there are companies other than The MuShield Company that have similar proprietary alloys for magnetic shielding, materials such as soft iron can be used if a less expensive option is desired.
The amount of shielding is directly linked to the thickness of the material. For instance, if using soft iron, the thickness of the enclosure would need to be substantially greater. See also Note 6.
Considering effects from radiofrequencies (RF)
Our recent data suggest that RF fields from wireless sources (such as cell phones and radio stations) can also cause biological effects [15,16] and could represent a confounding variable.
To protect experiments from RF interference, wrap an extra layer of shielding completely covering the mu-metal enclosure in the form of aluminum foil, which reduces exposure by 15–20 dB.
Designing a MagShield apparatus of different dimensions
Start the design process by defining the size of the coils needed. The size of your coils is closely tied to the dimensions of your sample plates, Petri dishes, or containers. This protocol is tailored for samples in 60 mm Petri dishes or 96-well plates.
Note: Reference [29] provides designs for many variations of Helmholtz coils.
There are limits to the number of samples that can be accommodated and maintain a uniform field. In general, only one 60 mm dish or welled plate should be placed in each partition, with one control plate in one partition and one experimental plate in the other.
Middle-positioned wells of 96-well plates are ideal for this setup but do not put samples in the outer wells as the field strengths will not be comparable (i.e., not uniform across the entire plate).
Note: If wanting to change the dimensions of the Helmholtz coils, please note that the design and wire winding process for Helmholtz coils can be quite complex and the specifics can vary greatly depending on your application requirements. It is essential to take the necessary safety precautions and, if needed, consult with experts in electromagnetics or coil design to ensure your Helmholtz coils function as intended.
Coil considerations and relationship to field uniformity and temperature
Changing Helmholtz coil size will affect the MF produced, as the MF is inversely related to radius. Coils with a larger radius would result in a smaller MF and vice versa, as long as the current remains the same.
If not 3D printing your base frames, ensure you have frames made from non-conductive material, such as PVC or wood. These frames should be shaped like circles or squares, depending on your design preference, and sized appropriately for the coils.
Determine the desired number of windings for each coil. This protocol used 50 windings tested with fields up to 900 µT. This depends on your specific application and the MF strength you want to generate. You can use mathematical formulas or MF simulation software for precise calculations.
The strength of the MF you intend to work with will affect the amount of heat that is produced. Higher field strengths generate more heat, which must be factored into your setup. In addition, any change to the power supplies used, as well as the thickness of the copper wire and the number of windings, will also affect the amount of heat produced.
Relationship of field strength to temperature
The power supplies recommended in this protocol can accommodate MF strengths ranging from near-zero to 1 mT without causing significant temperature fluctuations (see Supplemental File S5 for temperature fluctuation data at various MF strengths). In case higher field strengths are desired, potential modification options to mitigate any associated temperature increases include:
Increased number of windings: By increasing the number of windings in each row of a Helmholtz coil pair, the same MF strength can be achieved with a lower current, reducing heat generation.
Temperature-resistant coating: Applying a temperature-resistant coating to the copper wire/coils can help dissipate heat more effectively, minimizing potential temperature increases in the chamber.
Thicker copper wire: Using thicker wire for your coils can reduce the resistance and thus the heat generated. This can also help in maintaining a stable temperature within the experimental setup.
Note: Mu-metal barriers are typically custom-made for specific applications and must be designed according to the specific requirements of the magnetic shielding. If you are dealing with high-strength MFs or complex shielding requirements the barrier will need to be modified to be thicker or more layers may be necessary.
Relationship of Mu-metal thickness to shielding capabilities
The thickness of mu-metal plays a pivotal role in determining its shielding capabilities. Mu-metal barriers are typically custom-made to meet specific shielding requirements, especially for high-strength MFs or complex shielding needs. A stronger barrier is needed if you intend to work with a high-strength MF. In case a specific desired thickness is not available, layering is a potential option.
Accommodating other model systems
The specific model system you are using will have effects on the design of your MagShield apparatus. These mu-metal enclosures are suitable for placement inside incubators to maintain any required environmental conditions. This protocol specifically caters to room temperature aquatic model systems like Xenopus and planarians, as well as cell culture systems placed within a standard CO2 incubator with internal dimensions of 18.5 × 23.9 × 22.7 in (W × H × D). If a different-sized incubator is used, its interior dimensions must be taken into account when constructing the MagShield apparatus to ensure it will fit inside (see modifications for cell culture).
Acknowledgments
This work was funded by grants from: the National Science Foundation (NSF #2105474 to W.S.B., NSF #2217457 to I.N., and NSF #2244087 to A.F. and J.G.), Western Michigan University (WMU #FRACAA2624 to W.S.B.), National Aeronautics and Space Administration (NASA # 80NSSC20M0043 to K-AS.T.), the Defense Advanced Research Projects Agency (DARPA #HR00111810006 to F.S.B.), the National Institutes of Health (NIH #1R15GM150073-01 to W.S.B.), the University of Colorado (to F.S.B.), and the Milheim Foundation (to F.S.B.). Funding was also provided to J.V. by the Fulbright Foreign Student Program, which is sponsored by the U.S. Department of State. (The contents herein are solely the responsibility of the authors and do not necessarily represent the official views of the Fulbright Program or the Government of the United States.) The graphical abstract and the diagram in Figure 12A were created with BioRender.com. Thanks to Ashton Pearson (Machine Shop, UNLV Science and Engineering Building) for helping with 3D printing secondary containers for hypomagnetic exposures. We would like to acknowledge that the three original mu-metal boxes were obtained from Carl Blackman when his lab was closed; modifications to this design have been minor.
Competing interests
The authors confirm that there are no competing interests.
Ethical considerations
X. laevis were cultured through approved protocols and guidelines (UNLV Institutional Animal Care and Use Committee). Embryos were generated through in vitro fertilization and raised in 0.1× Marc's Modified Ringer (1 mM MgSO4, 2.0 mM KC1, 2 mM CaCl, 0.1 M NaC1, 5 mM HEPES, pH 7.8) medium. Embryos were grown to indicated stages/days.
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Supplementary information
The following supporting information can be downloaded here:
File S1: STL file for 3D printing coil base frames(stl)
File S2: Schematic for MagShield apparatus enclosure(pdf)
File S3: Calibration data for MF strength vs. power supply parameters(pdf)
File S4: Schematics for hypomagnetic field secondary enclosures(pdf)
File S5: Temperature fluctuations at different magnetic field strengths(pdf)
Article Information
Publication history
Received: May 22, 2024
Accepted: Jul 31, 2024
Available online: Sep 10, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Acutely Modifying Phosphatidylinositol Phosphates on Endolysosomes Using Chemically Inducible Dimerization Systems
WY Wei Sheng Yap
PK Peter K. Kim
MB Maxime Boutry
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5078 Views: 322
Reviewed by: Ralph Thomas BoettcherAlexandros C Kokotos Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Jun 2023
Abstract
Phosphoinositides are rare membrane lipids that mediate cell signaling and membrane dynamics. PI(4)P and PI(3)P are two major phosphoinositides crucial for endolysosomal functions and dynamics, making them the lipids of interest in many studies. The acute modulation of phosphoinositides at a given organelle membrane can reveal important insights into their cellular function. Indeed, the localized depletion of PI(4)P and PI(3)P is a viable tool to assess the importance of these phosphoinositides in various experimental conditions. Here, we describe a live imaging method to acutely deplete PI(4)P and PI(3)P on endolysosomes. The depletion assay utilizes the GAI-GID1 or the FRB-FKBP inducible dimerization system to target the catalytic domain of the PI(4)P phosphatase, Sac1, or the PI(3)P phosphatase domain of MTM1 to the endolysosome for localized depletion of these phosphoinositides. By using the fluorescently tagged biosensors, 2xP4M and PX, we can validate and monitor the depletion of PI(4)P and PI(3)P, respectively, on endolysosomes in real-time. We discuss a method for normalizing the fluorescence measurements to appropriate the relative amount of these phosphoinositides in the organellar membranes (endolysosomes), which is required for monitoring PI(4)P or PI(3)P levels during the acute depletion assay. Since the localization of the dimerization partners is specified by the membrane targeting signal, our protocol will be useful for studying the signaling and functions of phosphoinositides at any membrane.
Key features
• Acute depletion and real-time monitoring of PI(3)P and PI(4)P on the endolysosomal membrane using chemically inducible dimerization systems.
• Modifiable and adaptable to modulate other phosphoinositides on different organellar membranes.
Keywords: Phosphoinositides PI(4)P Chemically inducible dimerization system Endolysosomes Live-cell imaging High-resolution fluorescence microscopy
Graphical overview
Background
Phosphoinositides, such as PI(4)P and PI(3)P, are phosphorylated derivatives of phosphatidylinositol, a phospholipid found in the cytoplasmic leaflet of eukaryotic membranes. The inositol ring of the phosphatidylinositol headgroup can be phosphorylated by specific kinases, and dephosphorylated by phosphatases, in position 3, 4, and 5 for a total of seven different phosphoinositides [1]. Phosphoinositides are key signaling lipids that regulate organelle membrane identity and dynamics. They notably play major roles in orchestrating various endolysosomal functions and dynamic events such as tubulation and fission, which are required for the reformation of lysosomes at the end of the autophagic and endocytic processes [2–5].
One elegant way to investigate the role of phosphoinositides in organelle biology is to modulate their levels at a given membrane. This can be done by specifically targeting phosphoinositide kinases or phosphatases to an organelle of interest. Targeting can be achieved by fusing the catalytic domain of kinases or phosphatases to a membrane-targeting domain (typically a transmembrane domain) of a resident membrane protein of the desired organelle. This allows for the constitutive depletion of the phosphoinositides upon the expression of the protein. However, to allow for temporal and acute regulation of the phosphoinositides species, the catalytic domain can be targeted using a chemically induced dimerization system. Here, one part of the dimer is anchored at a membrane of interest by fusing it to a transmembrane protein/targeting sequence, and the other is coupled to a cytosolic version of a phosphatase or kinase. Upon the addition of chemical inducers of dimerization, the binding affinity between the two-part dimerization system increases drastically, promoting the recruitment of the cytosolic part to the membrane-anchored part of the dimer. As a result, the recruited kinases and phosphatases actively phosphorylate and dephosphorylate the phosphoinositide substrate, respectively, depleting them from the organellar membrane.
In addition, the level of specific phosphoinositides at membranes in cells can be evaluated by the ectopic expression of genetically encoded phosphoinositides biosensors coupled to a fluorescent protein, such as GFP or mCherry. Over the last 20 years, high-affinity binding domains for specific phosphoinositides have been identified and used as biosensors to detect different phosphoinositide species [6]. Here, we present a protocol utilizing chemically induced dimerization systems to deplete specific phosphoinositides at endolysosomes and biosensors to detect their depletion. We describe how to use two different dimerization systems in cells, the GAI-GID1 and the FRB-FKBP, to deplete PI(4)P and PI(3)P, respectively, at Rab7-positive endolysosomes (hereafter referred to as endolysosomes). However, any of the two dimerization systems can be used to deplete the phosphoinositides on any other membrane-bound organelle of interest. In this protocol, we discuss the experimental setup to express these systems in mammalian cells and how to perform image acquisition and analysis to validate the depletion of PI(4)P or PI(3)P.
The FRB-FKBP dimerization system was invented in the 90’s [7] and relies on the dimerization of FRB and FKBP in the presence of rapamycin, a cell-permeant chemical most famous for its inhibitory effect on mTORC1 (mammalian target of rapamycin complex 1). The FRB and FKBP domains are relatively small (~10 kDa) and thus can be efficiently transfected and expressed, reducing the likeliness of impacting the function of the protein of interest fused to these domains. However, using rapamycin in the FRB-FKBP dimerization system may cause undesirable changes in certain mammalian biological processes such as autophagy and nutrient-sensing pathways, notably due to its effect on mTORC1. To circumvent this problem, rapalogs (rapamycin analogs) can be used to alleviate this cross-reactivity [8]. Another way to mitigate the complications of rapamycin is to use the plant-derived GA3-AM-inducible GAI-GID1 dimerization system since there is no known mammalian target of GA3-AM [9]. However, unlike the FRB-FKBP system, both GAI and GID1 are medium-sized protein domains (~59 and ~40 kDa, respectively) which may impede the efficient introduction or expression of proteins. Control experiments can then be performed to validate that tagging with dimerizing proteins does not impair the localization or the function of the protein of interest. Here, we present a protocol to target PI(4)P or PI(3)P phosphatases to endolysosomes and to validate phosphoinositides depletion using genetically encoded PI(4)P or PI(3)P biosensors.
Materials and reagents
Cell line
HeLa cells from ATCC (CCL-2) [here, HeLa cells are used. However, this can be used in any cell lines. The only difference may be the system to insert the necessary plasmid into the cell (transfections vs. transduction vs. electroporation, etc.)]
Plasmids (Table 1)
Table 1. Plasmids used in this protocol
Plasmid Addgene number Reference
iRFP-GID1-Rab7 Levin-Konigsberg et al. [10]
CFP-GAI-Sac1 Boutry and Kim [11]
CFP-GAI-Sac1 C392S Boutry and Kim [11]
mCherry-2xP4M Levin-Konigsberg et al. [10]
Lamp1-mCherry 45147 Van Engelenburg and Palmer [12]
iRFP-FRB-Rab7 51613 Hammond et al. [13]
mCherry-FKBP-MTM1 51614 Hammond et al. [13]
mCherry-FKBP 67514 Varnai et al. [14]
PX-GFP 19010 Kanai et al. [15]
Lamp1-GFP 16290 Minin et al. [16]
Reagents
DMEM, 4.5 g/L glucose, supplemented with L-glutamine and sodium pyruvate (Wisent, catalog number: 319-005 CL)
Fetal bovine serum (FBS) (Wisent, catalog number: 098150)
Trypsin-EDTA (0.05% Trypsin, 0.53 mM EDTA) (Wisent, catalog number: 325542105)
D-PBS (Wisent, catalog number: 311-45 CL)
Neon electroporation kit 100 μL (Invitrogen, catalog number: MPK10096)
Resuspension buffer R
Electrolytic buffer E2
Neon electroporation tips (100 µL)
Electroporation tubes
Rapamycin (Sigma-Aldrich, catalog number: 553210)
GA3-AM (Sigma-Aldrich, catalog number: SML1959)
Laboratory supplies
T75 cm2 cell culture flasks (Sarstedt Inc, catalog number: 50-809-261)
4-well Lab-Tek II chambered cover glass (Nunc, catalog number: 155382)
15 mL conical tubes
1.5 mL Eppendorf tubes
Equipment
Micropipette (P1000, P200, and P10)
Centrifuge with adaptors for 15 mL conical tubes
Tissue culture CO2 incubator
Standard inverted light microscope
Hemocytometer with cover glass
Inverted microscope (Plan-Apochromat 63×/1.40 oil objective) (Zeiss, model: Airyscan 2 LSM 980) with temperature- and CO2-controlled imaging chamber
Neon NxT Electroporation System (electroporation device, electroporation pipette and pipette station, electroporation tube, and tube chamber)
Software and datasets
ImageJ [17] or Fiji software [18] (both open source)
Procedure
Transfection
We found that lipofection-based methods of transfection can affect the morphology of endolysosomes. Therefore, we recommend electroporation methods such as the Neon electroporation system (Invitrogen) for these assays. The use of antibiotics and antimycotics should be avoided as it affects cell viability after electroporation.
Culture HeLa cells in T75 cm2 flasks in DMEM containing 10% FBS (complete media) until cells reach 70%–80% confluency.
Remove media and wash with D-PBS. Add 2 mL of trypsin-EDTA per flask and incubate for 5 min at 37 °C or until cells are detached.
Add 8 mL of complete media to stop trypsin activity.
Place a clean cover glass on top of the hemocytometer (counting chamber facing up). Transfer 10 µL of the cell suspension into the counting chamber by capillary action and view under a standard inverted light microscope. The hemocytometer chamber is composed of nine large, gridded squares, each containing 0.1 mm3 space (= 0.1 µL). Count the cells in every gridded square and average them. Estimate the cell density by dividing the average number of cells per gridded square by 1 mm3 or 0.1 µL.
Transfer 0.5 × 106 cells to a new 15 mL conical tube per transfection.
Spin cells at 300× g for 5 min, remove supernatant, and resuspend with 10 mL of D-PBS.
Spin at 300× g for 5 min and remove the supernatant. Resuspend the cell pellet in 100 µL of Neon electroporation R buffer and transfer to a 1.5 mL Eppendorf tube.
Add DNA (dissolved in water or Tris-EDTA elution buffer) to cells. The manufacturer recommends adding 5 µg of total DNA per 100 µL of cells resuspended in R buffer. The DNA amount is plasmid-dependent and may require optimization.
For PI(4)P depletion using the GAI-GID1 system:
An equal amount of iRFP-GID1-Rab7 + CFP-GAI-Sac1 or CFP-GAI-Sac1 C392S (catalytically dead Sac1 used as a negative control). Co-transfection with an equal amount of PI(4)P biosensor mCherry-2xP4M to assess PI(4)P depletion or Lamp1-mCherry to assess the effect of PI(4)P depletion on endolysosomal morphology.
For PI(3)P depletion using the FKBP-FRB dimerization system:
An equal amount of iRFP-FRB-Rab7 + mCherry-FKBP-MTM1 or mCherry-FKBP (used as a negative control). Co-transfection with an equal amount of PI(3)P biosensor PX-GFP to monitor for PI(3)P depletion or Lamp1-GFP to assess the effect of PI(3)P depletion on endolysosomes.
Fill the electroporation tube with 3 mL of buffer E2. Insert the tube into the tube chamber.
Ensure that the electroporation parameters are customized according to the manufacturer’s protocol (https://www.thermofisher.com/ca/en/home/life-science/cell-culture/transfection/neon-transfection-system/neon-transfection-system-cell-line-data.html). For HeLa cells, set the pulse number to 2, voltage at 1,005 V, and width at 35 ms. Start the electroporation.
Dilute the electroporated cell suspension with 2.4 mL of warm complete media.
Seed 0.5 mL of the diluted cell suspension (equivalent to 0.1 × 106 cells) per cell chamber (Lab-tek II chambered cover glass, 1.7 cm2 per well).
Incubate the cell chamber in the tissue culture CO2 incubator for 12–36 h for plasmid expression.
Imaging and dimerization
Perform live microscopy imaging 24 h post-electroporation. On the day of imaging, prepare an aliquot of complete media supplemented with either 20 µM GA3-AM or 200 nM rapamycin (2× concentration, to a final concentration of 10 µM GA3-AM or 100 nM rapamycin). For each condition, prepare 300 µL of solutions (sufficient to cover a 1.7 cm2 cell chamber).
This protocol uses the Zeiss Airyscan 2 LSM 980 inverted microscope with Plan-Apochromat 63×/1.40 oil objective at 37 °C in a 5% CO2-controlled imaging chamber. Before imaging, ensure the appropriate lasers and filters setup according to the wavelengths (λ) of the excitation (Ex)/emission (Em) maxima of the fluorescent proteins listed in Table 2. Furthermore, ensure that the stage temperature is stable.
Table 2. Excitation/emission maxima of the fluorescent proteins used in this protocol
Fluorescent protein Ex maximum λ (nm) Em maximum λ (nm) Laser Ex λ (nm)* Em filter*
iRFP 690 713 640 LP660
CFP 456 480 488 BP420-500
GFP 488 507 488 BP495-555
mCherry 587 610 561 BP570-620
*Note: The laser Ex λ and Em filters setup is based on our Airyscan 2 LSM 980 system. Similar lasers and filters can be used on other systems.
Replace each cell chamber with 300 µL of fresh complete media.
Secure the cell chamber slide on the stage with the appropriate adaptor.
Using ZEN microscopy software, add “time series” and “Z-stacks” to the acquisition parameters to record stacks of time-lapse images. Dimerization should occur within 10 min after adding the inducing agents (GA3-AM or rapamycin). We suggest capturing a 10-min video with an interval of 1 min for kinetic assays. For end-point assays, capture images up to 1 h after induction.
For kinetic assays, move the stage to a field of monolayer cells and start the acquisition using the parameters as described in step 5. After acquiring the first stacks, wait for 30 s and carefully pipette an equal volume (300 µL) of the solution prepared from step B1 into the cell chamber well (the final volume is now 600 µL). The acquisition should continue automatically as preconfigured in the software.
For end-point assays, image the cells before induction. Then, add an equal volume of the solution and incubate for at least 10 min and up to 1 h followed by another round of post-induction imaging.
Data analysis
Here, we present analysis pipelines to assess (1) the recruitment of the phosphatases to endolysosomes (= dimerization efficiency) and (2) the subsequent specific phosphoinositides level changes caused by this recruitment. The dimerization efficiency is monitored by quantifying the colocalization between the phosphatase and the Rab7 constructs used to recruit phosphatases to endolysosomes. We find that the GAI-GID1 and FKBP-FRB systems display very similar recruitment kinetics, leading to an efficient dimerization within minutes. The depletion of phosphoinositides, here PI(4)P and PI(3)P, is monitored by measuring the levels of specific biosensors [2xP4M for PI(4)P and PX for PI(3)P] at endolysosomes normalized to either plasma membrane (PM) levels of 2xP4M and cytosolic background for PX. We use Fiji, an open-source software for this analysis. Other software such as Volocity and CellProfiler can be used. When opening the raw ZEN files using Fiji, drag the file onto the Fiji main control panel. When the Bio-Formats Import Options window pops up, ensure that the Color mode option is set to Default, which opens the images with the channels split (Figure S1). By default, the ZEN files are in 16-bit depths.
(1) Validation of GAI or FKBP recruitment to endolysosomes
Open the raw ZEN files using Fiji. In this example, for simplicity, the endolysosomal marker channel is designated as Channel 1 (Ch1) and the GAI- or FKBP-fused fluorescence channel is designated as Channel 2 (Ch2).
Specify (Edit > Selection > Specify…) a subregion for analysis. Here, a Z-slice of a ROI of 15 × 15 µm square region (highlighted in yellow) containing resolvable endolysosomes was selected (Figure S2, box A).
Crop (Image > Crop) or duplicate (Image > Duplicate…) this subregion for subsequent analysis.
Split the channels (Image > Color > Split Channels).
Select Ch1 window (Figure S2, box B). Threshold (Image > Adjust > Threshold…) the image to accurately mark the endolysosomal regions. Click Apply and this will automatically binarize the image and convert it to 8-bit (Figure S2, box C).
Create a mask using the Create Selection (Edit > Selection > Create Selection) and the Add to Manager (Edit > Selection > Add to Manager) commands. Here, the mask is outlined in yellow (Figure S3, box A) and the selected mask is registered at the ROI manager (Figure S3, box B).
Switch to Ch2 window (Figure S3, box C).
Select the registered ROI (from step 6) and use the Measure command (Analyze > Measure) to acquire the mean intensity of fluorescent protein-fused GAI or FKBP in the specified endolysosomal regions (Figure S3, box D). If there is no “Mean” column displayed in the Results window, execute the Set Measurements… command (Analyze > Set Measurements…). Tick the Mean gray value checkbox and rerun the Measure command.
Repeat the measurement (steps 3–8) for all the time-lapse slices, if any.
The measured fluorescence intensity after the induction (F) is normalized to the first slice (F0, before the addition of GA3-AM or rapamycin). Use the equation below to calculate the normalized endolysosomal GAI or FKBP fluorescence intensity (Ch2norm) of each slice:
C
h
2
F
C
h
2
F
0
=
C
h
2
n
o
r
m
The change in the GAI or FKBP fluorescence intensity on the endolysosomal mask over time indicates the recruitment efficiency. Plot the Ch2norm value against time to obtain the kinetics curve as shown in Figure 1.
Figure 1. Validation of GAI or FKBP recruitment to endolysosomes and its efficiency. (A) Representative live images of HeLa cells co-expressing RFP-FKBP (purple) and iRFP-RFB-Rab7 (green) before and after dimerization induction with 100 nM rapamycin. Scale bar, 15 µm. The right panel shows the dimerization kinetics of the endo-lysosomal targeting FRB-FKBP system in this assay. Error bar = SEM. Cells from three independent experiments, n = 11. (B) Representative live images of HeLa cells co-expressing CFP-GAI (purple) and iRFP-GID1-Rab7 (green) before and after dimerization induction with 10 µM GA3-AM. Scale bar, 15 µm. The right panel shows the dimerization kinetics of the endolysosomal targeting GAI-GID1 system in this assay. Error bar = SEM. Cells from three independent experiments, n = 10.
(2) Validating phosphoinositides depletion at endolysosomes:
Validation of PI(4)P depletion at endolysosomes
Open the raw ZEN files using Fiji. In this example, for simplicity, the mCherry-2xP4M channel is designated as Ch1, and the Rab7 endolysosomal marker channel is designated as Ch2.
The PM contains a great amount of PI(4)P (and therefore high 2xP4M fluorescence intensity) and is not expected to change upon the induced recruitment of Sac1 to the endolysosomes, making the PM 2xP4M signal a great reference for normalization. Measure PM mCherry-2xP4M signal intensity by performing line scans perpendicularly across three PM regions in Ch1. Record the maximum intensity (do not record the mean intensity as the signal is expected to be enriched only at the PM but not in the cytosol and extracellular region) using the Plot Profile command (Analyze > Plot Profile). Average these three values.
Switch to Ch2, specify a 15 × 15 µm square region, and threshold the image. Create a mask of Rab7-positive endolysosomes and add it to the ROI manager.
To measure the mean intensity of endolysosomal mCherry-2xP4M, switch back to Ch1 and select the ROI registered from step 3. Record the mean intensity.
Divide the mean lysosomal mCherry-2xP4M intensity obtained from step 4 by the mean PM mCherry-2xP4M intensity from step 2 to calculate the normalized lysosomal PI(4)P level:
C
h
1
R
a
b
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=
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m
a
l
i
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4
)
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An example of such an analysis is shown in Figure 2A.
(3) Validation of PI(3)P deletion at endolysosomes
Open the raw ZEN files using Fiji. In this example, for simplicity, the PX-GFP channel is designated as Ch1, and the Rab7 endolysosomal marker channel is designated as Ch2.
Specify a 15 × 15 µm square region for analysis. Avoid any area outside of the cell as it will affect normalization.
In Ch1, measure the mean intensity. This value will be used as background normalization.
Switch to Ch2 and threshold the image. Make a mask of the Rab7-positive endolysosome and add it to the ROI manager.
Switch back to Ch1 and specify the ROI registered from step 4. Measure the mean intensity.
Divide the mean lysosomal PX-GFP intensity obtained from step 5 by the overall mean PX-GFP intensity to calculate the normalized lysosomal PI(3)P level:
C
h
1
R
a
b
7
C
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m
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3
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An example of such an analysis is shown in Figure 2B.
Figure 2. Validation of endolysosomal PI(4)P or PI(3)P depletion. (A) Representative live images of HeLa cells co-expressing mCherry-2XP4M (purple) and iRFP-GID1-Rab7 (green) in addition to GAI-Sac1 or GAI-Sac1(C392S) expression. Images were acquired after dimerization induction with 10 µM GA3-AM. Scale bar, 15 µm. Right panel shows the quantification of the lysosomal 2XP4M signal normalized to the PM 2XP4M signal. Statistical significance was calculated from the Student’s t-test. Error bar = SEM. n = 31 cells (GAI-Sac1) and n = 30 cells (GAI-Sac1 C392S) from three independent experiments. (B) Representative live images of HeLa cells co-expressing PX-GFP (purple) and iRFP-FRB-Rab7 (green) in addition to FKBP-MTM1 or FKBP expression. Images were acquired after dimerization induction with 100 nM rapamycin. Scale bar, 15 µm. Right panel shows the quantification of the lysosomal PX signal normalized to the cytosolic PX signal. Statistical significance was calculated from the Student’s t-test. Error bar = SEM. n = 31cells (FKBP-MTM1) and n = 31 cells (FKBP) from three independent experiments.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Boutry et al. [5]. Arf1-PI4KIIIβ positive vesicles regulate PI(3)P signaling to facilitate lysosomal tubule fission. J Cell Biol (Figure 6, panels D–J).]
Boutry and Kim. [11]. ORP1L mediated PI(4)P signaling at ER-lysosome-mitochondrion three-way contact contributes to mitochondrial division. Nature Communications (Figure 3, panels I–N; Figure 4, panel M; Figure 5, panels I–L; Figure 7, panels A–C and I–K; Supplementary Figure 8, panels A–I).
Acknowledgments
Infrastructure for the Kim Laboratory was provided by a John Evans Leadership Fund grant from the Canadian Foundation for Innovation and the Ontario Innovation Trust. This work was supported by operating grants from the Canadian Institutes of Health Research to P.K. Kim (PJT#180476); M. Boutry was supported by a Restracomp Fellowship from the Hospital for Sick Children and a Canadian Institutes of Health Research Postdoctoral fellowship. This protocol was used in: Boutry et al. J Cell Biol (2023), DOI: 10.1083/jcb.202205128 and Boutry and Kim. Nat Commun (2021), DOI: 10.1038/s41467-021-25621-4 and was adapted from Miyamoto et al. Nat Chem Biol (2012), DOI: 10.1038/nchembio.922 and Spencer et al. Science (1993), DOI: 10.1126/science.7694365.
Competing interests
The authors declare that they have no competing interests.
References
Balla, T. (2013). Phosphoinositides: Tiny Lipids With Giant Impact on Cell Regulation. Physiol Rev. 93(3): 1019–1137.
Sridhar, S., Patel, B., Aphkhazava, D., Macian, F., Santambrogio, L., Shields, D. and Cuervo, A. M. (2012). The lipid kinase PI4KIIIβ preserves lysosomal identity. EMBO J. 32(3): 324–339.
Rong, Y., Liu, M., Ma, L., Du, W., Zhang, H., Tian, Y., Cao, Z., Li, Y., Ren, H., Zhang, C., et al. (2012). Clathrin and phosphatidylinositol-4,5-bisphosphate regulate autophagic lysosome reformation. Nat Cell Biol. 14(9): 924–934.
Jani, R. A., Di Cicco, A., Keren-Kaplan, T., Vale-Costa, S., Hamaoui, D., Hurbain, I., Tsai, F. C., Di Marco, M., Macé, A. S., Zhu, Y., et al. (2022). PI4P and BLOC-1 remodel endosomal membranes into tubules. J Cell Biol. 221(11): e202110132.
Boutry, M., DiGiovanni, L. F., Demers, N., Fountain, A., Mamand, S., Botelho, R. J. and Kim, P. K. (2023). Arf1-PI4KIIIβ positive vesicles regulate PI(3)P signaling to facilitate lysosomal tubule fission. J Cell Biol. 222(9): e202205128.
Hammond, G. R. and Balla, T. (2015). Polyphosphoinositide binding domains: Key to inositol lipid biology. Biochim Biophys Acta Mol Cell Biol Lipids. 1851(6): 746–758.
Spencer, D. M., Wandless, T. J., Schreiber, S. L. and Crabtree, G. R. (1993). Controlling Signal Transduction with Synthetic Ligands. Science. 262(5136): 1019–1024.
Bayle, J. H., Grimley, J. S., Stankunas, K., Gestwicki, J. E., Wandless, T. J. and Crabtree, G. R. (2006). Rapamycin Analogs with Differential Binding Specificity Permit Orthogonal Control of Protein Activity. Chem Biol 13(1): 99–107.
Miyamoto, T., DeRose, R., Suarez, A., Ueno, T., Chen, M., Sun, T. p., Wolfgang, M. J., Mukherjee, C., Meyers, D. J., Inoue, T., et al. (2012). Rapid and orthogonal logic gating with a gibberellin-induced dimerization system. Nat Chem Biol. 8(5): 465–470.
Levin-Konigsberg, R., Montaño-Rendón, F., Keren-Kaplan, T., Li, R., Ego, B., Mylvaganam, S., DiCiccio, J. E., Trimble, W. S., Bassik, M. C., Bonifacino, J. S., et al. (2019). Phagolysosome resolution requires contacts with the endoplasmic reticulum and phosphatidylinositol-4-phosphate signalling. Nat Cell Biol. 21(10): 1234–1247.
Boutry, M. and Kim, P. K. (2021). ORP1L mediated PI(4)P signaling at ER-lysosome-mitochondrion three-way contact contributes to mitochondrial division. Nat Commun. 12(1): 5354.
Van Engelenburg, S. B. and Palmer, A. E. (2010). Imaging type-III secretion reveals dynamics and spatial segregation of Salmonella effectors. Nat Methods. 7(4): 325–330.
Hammond, G. R., Machner, M. P. and Balla, T. (2014). A novel probe for phosphatidylinositol 4-phosphate reveals multiple pools beyond the Golgi. J Cell Biol. 205(1): 113–126.
Varnai, P., Thyagarajan, B., Rohacs, T. and Balla, T. (2006). Rapidly inducible changes in phosphatidylinositol 4,5-bisphosphate levels influence multiple regulatory functions of the lipid in intact living cells. J Cell Biol. 175(3): 377–382.
Kanai, F., Liu, H., Field, S. J., Akbary, H., Matsuo, T., Brown, G. E., Cantley, L. C. and Yaffe, M. B. (2001). The PX domains of p47phox and p40phox bind to lipid products of PI(3)K. Nat Cell Biol. 3(7): 675–678.
Minin, A. A., Kulik, A. V., Gyoeva, F. K., Li, Y., Goshima, G. and Gelfand, V. I. (2006). Regulation of mitochondria distribution by RhoA and formins. J Cell Sci. 119(4): 659–670.
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Supplementary information
The following supporting information can be downloaded here:
Figure S1. Import options in the Bio-Formats Import Options used in this protocol.
Figure S2. Thresholding pipeline to make a mask of endolysosomal regions.
Figure S3. Measuring the fluorescence intensity in the GAI- or FKBP-fused fluorescence channel within the mask of endolysosomal regions.
Article Information
Publication history
Received: May 25, 2024
Accepted: Aug 7, 2024
Available online: Sep 13, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cell Biology > Cell imaging > Live-cell imaging
Cell Biology > Cell imaging > Confocal microscopy
Cell Biology > Cell structure > Cell organelle
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Measuring Piezo1 and Actin Polarity in Chemokine-Stimulated Jurkat Cells During Live-Cell Imaging
CL Chinky Shiu Chen Liu
PB Parijat Biswas
Dipyaman Ganguly
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5079 Views: 258
Reviewed by: Ivonne SehringEVANGELOS THEODOROUSamantha Haller
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Original Research Article:
The authors used this protocol in eLIFE Feb 2024
Abstract
The process of T-lymphocyte migration involves a complex interplay of chemical and mechanical signals. Mechanotransduction mechanisms in T lymphocytes enable them to efficiently navigate through diverse architectural and topographical features of the dynamic tissue macro- and micro-niches encountered during immune responses. Piezo1 mechanosensors are crucial for driving optimal T-cell migration by driving actin-cytoskeletal remodeling. Chemokine-stimulated T lymphocytes demonstrate significant asymmetry or polarity of Piezo1 and actin along the cell axis. The establishment and maintenance of polarity in migrating cells are paramount for facilitating coordinated and directional movements along gradients of chemokine signals. Live-cell imaging techniques are widely employed to study the trajectories of migrating cells. Our approach expands upon current methodologies by not only tracking migrating cells but also imaging fluorescently labeled cellular components. Specifically, our method enables measurement of protein enrichment in the front and rear halves of the moving cell by analyzing the temporal direction of cell trajectories, subsequently bisecting the cell into front-back halves, and measuring the intensities of the fluorescent signals in each cell half at each time frame. Our protocol also facilitates the quantification of the angular distribution of fluorescent signals, enabling visualization of the spatial distribution of signals relative to the direction of cell migration. The protocol describes the examination of polarity in chemokine-treated Jurkat cells transfected with Piezo1-mCherry and actin-GFP constructs. This approach can be extended to live-cell imaging and polarity assessment of other fluorescently labeled proteins.
Key features
• This experimental protocol allows real-time imaging of Jurkat cells expressing two fluorescent proteins (Piezo1 mCherry and actin-GFP).
• Measures cell polarity by examining spatial enrichment of Piezo1 and actin proteins within the front and rear halves of a moving Jurkat cell.
• The protocol enables analysis of cell polarity in 2D tracks of moving cells.
• Polarity analysis includes measuring fluorescent signal intensities in front-rear halves of a moving cell and calculation of signal polarization angles relative to the cell trajectory.
Keywords: Time-lapse confocal imaging Jurkat cell line Piezo1 mCherry Actin-GFP imaging Cell polarity Angular distribution
Graphical overview
Schematic illustration of the experimental and analysis workflow. A. Jurkat cells were nucleofected with plasmids expressing Piezo1 mCherry and actin-GFP. The pulsed cells were cultured in complete RPMI medium for 36 h at 37 °C/5% CO2 to allow protein expression. B. Transfected Jurkat cells were collected, washed, and resuspended in imaging medium. Cells were seeded in ICAM-1-coated glass bottom confocal dishes and incubated for 1 h at 37 °C/5% CO2 to facilitate cell attachment. C. Live-cell tracking of transfected cells was performed in a confocal incubation setup prewarmed to 37 °C. Chemokine SDF1α was added 10–15 min prior to image acquisition. Images were acquired for a total duration of 5 min at intervals of 30 s. Z-stacks of 1 µm were captured at each frame. Fiji/ImageJ was used for cell segmentation by thresholding, ROI detection, trajectory, and polarity analysis.
Background
Migration of immune cells is finely orchestrated by an intricate interplay of molecular and physical signals [1–3]. These signals efficiently navigate the cells toward inflamed tissues and secondary lymphoid organs, where they mount the onset of the vital adaptive immune response. Migration of immune cells including T lymphocytes is mechanically intensive [4–6]. Motile T lymphocytes actively engage in continuous sensing and adaptation to mechanical cues within their surroundings. These signals encompass various parameters such as local tissue structure and topography, mechanical rigidity, confinement, hydrostatic pressure, and shear stress from interstitial fluid and circulating blood [5–7]. In addition, T lymphocytes also sense cell-intrinsic mechanical forces that stem from numerous cellular processes including tethering interactions with the extracellular matrix (ECM), actomyosin cytoskeletal remodeling, generation of traction forces, consequent exertion of forces on the tethered membrane, and changes in membrane tension [4–7]. Forces produced by cells facilitate unimpeded movement by overcoming resistance stress generated from local tissue deformation during migration [8–10].
The mechanosensitive ion channel Piezo1 has been widely implicated in numerous physiological processes that are subject to intricate mechanical regulation [11–13]. These processes include vascular remodeling, stem cell differentiation, epithelial homeostasis, and regulation of red blood cell volume [14–19]. Key research has also delved into the function of these mechanosensors in regulating the activation and function of T and B lymphocytes, dendritic cells, and macrophages [20–27]. The homo-trimeric subunits arranged in a propeller blade-like fashion facilitate Piezo1 to sense changes in membrane tension caused by applied force. Piezo1 subsequently undergoes a change in conformation allowing the opening of the central ion pore and passage of extracellular cations into the cell [28,29].
A pivotal investigation conducted by our team discovered the crucial role of Piezo1 in driving optimal human T-lymphocyte activation in response to T-cell receptor (TCR) triggering [20,30]. Further examination revealed that Piezo1-driven calcium response activates downstream calpain-dependent polymerization of the actin cytoskeleton, resulting in the formation of stable immunological synapses [20]. Subsequent deeper examination enabled us to identify a similar mechanism operating during chemokine-driven, ICAM-1-dependent human CD4+ T-lymphocyte migration [31]. Utilizing interference reflection microscopy (IRM) and confocal imaging, the study observed an increase in membrane tension in chemokine-stimulated T lymphocytes at focal adhesion sites. Polarized Piezo1 recruitment to these high-tension regions at the leading edge was accompanied by localized calcium signaling and calpain-driven F-actin polymerization. Local cytoskeletal rearrangement facilitated the recruitment of CD11a/LFA-1, triggering downstream PI3-kinase/Akt signaling that promoted further actin polymerization and flow, thus driving cell movement [31]. A notable finding of this study was the enrichment of Piezo1 at the leading edges of the cell, accompanied by a dynamic distribution of actin. In order to examine the spatial polarity of Piezo1 and actin in moving cells, we used the Jurkat cell line expressing Piezo1 mCherry and actin-GFP. Time-lapse imaging was conducted. Using a custom-designed code, the trajectories of the transfected cells were analyzed to determine the front-back polarity of the fluorescent signals, along with the distribution of their polarization angles.
Migrating cells exhibit distinct polarity characterized by an asymmetric distribution of cellular components along their migratory axis [32–34]. Generation and maintenance of cell polarity are essential for driving coordinated and directed movement of cells, in response to chemical (such as chemotaxis) and physical cues (such as haptotaxis) [35,36]. The protocol outlined below offers a comprehensive, step-by-step account of the experimental methodology and analytical approach employed to assess the polarity of Piezo1 mCherry and actin-GFP in chemokine-stimulated Jurkat cells. The methodologies described herein can be readily adapted for the evaluation of other proteins and in various experimental contexts.
Materials and reagents
Biological materials
Jurkat cell line [from Dr. Santusabuj Das at National Institute of Cholera and Enteric Diseases (NICED), Kolkata, India] (ATCC, catalog number: TIB-152)
Piezo1 mCherry plasmid (from Dr. Charles Cox at Victor Chang Cardiac Research Laboratory, Darlinghurst, Australia)
pCAG-mGFP-Actin plasmid (Addgene, catalog number: 21948)
Reagents
Recombinant human SDF1α/CXCL12 (PeproTech, catalog number: 300-28A)
Recombinant human ICAM-1 (PeproTech, catalog number: 150-05)
SE Cell Line 4D-NucleofectorTM X Kit L (Lonza, catalog number: V4XC-1024)
RPMI 1640 medium (Gibco, catalog number: 11875093)
RPMI 1640 medium, no phenol red (Gibco, catalog number: 11835030)
Fetal bovine serum (FBS) (Gibco, catalog number: 16000044)
Sodium pyruvate, 100 mM (Gibco, catalog number: 11360070)
MEM non-essential amino acids solution, 100× (Gibco, catalog number: 11140050)
Penicillin-streptomycin solution, 100× (Gibco, catalog number: 15140122)
Antibiotic-antimycotic, 100× (Gibco, catalog number: 15240062)
HEPES (Sigma-Aldrich, catalog number: H3375)
Phosphate-buffered saline, pH 7.4 (Himedia, catalog number: M1866)
Sodium hydroxide (Himedia, catalog number: PCT1325), 10 N solution in double-distilled water
Endotoxin-free water (InvivoGen, catalog number: h2olal-1.5)
UltraPureTM DNase/RNase-free distilled water (Invitrogen, catalog number: 10977015)
Solutions
Complete RPMI medium (see Recipes)
Post-nucleofection RPMI medium (see Recipes)
1 M HEPES solution, pH 7.4 (see Recipes)
RPMI medium without phenol red for imaging (see Recipes)
Phosphate-buffered saline (1×) (see Recipes)
Nucleofection solution (see Recipes)
Recombinant human ICAM-1 stock solution (see Recipes)
Recombinant human SDF1α stock solution (see Recipes)
Recipes
Complete RPMI medium
Reagent Final concentration Quantity or Volume
RPMI 1640 500 mL
Fetal bovine serum 10% 50 mL
Sodium pyruvate (100×) 1× 6 mL
MEM Non-essential amino acid (100×) 1× 6 mL
Penicillin-streptomycin (100×) 1× 6 mL
Antibiotic-antimycotic (100×) 1× 6 mL
* Filter final media using a 0.22 µm bottle-top vacuum filter system and store at 4 °C.
Post-nucleofection RPMI medium
Reagent Final concentration Quantity or Volume
RPMI 1640 40 mL
Fetal bovine serum 20% 10 mL
Sodium pyruvate (100×) 1× 500 µL
MEM Non-essential amino acid (100×) 1× 500 µL
* Filter final media using a 0.22 µm bottle-top vacuum filter system and store at 4 °C.
1 M HEPES solution, pH 7.4
Reagent Final concentration Quantity or Volume
HEPES 1 M 11.9 g
10 N sodium hydroxide as needed for pH = 7.5
Ultrapure distilled water 50 mL
* Adjust the pH to 7.5 using sodium hydroxide and make up the volume with ultrapure distilled water to 50 mL. Filter the solution using a 0.22 µm syringe filter and store at 4 °C.
RPMI medium without phenol red
Reagent Final concentration Quantity or volume
RPMI 1640 medium, no phenol red 500 mL
Fetal bovine serum 10% 50 mL
Sodium pyruvate (100×) 1× 6 mL
MEM Non-essential amino acid (100×) 1× 6 mL
Penicillin-streptomycin (100×) 1× 6 mL
Antibiotic-antimycotic (100×) 1× 6 mL
HEPES (100 mM), pH 7.5 25 mM 1.25 mL
* Filter the final media using a 0.22 µm bottle-top vacuum filter system and store at 4 °C.
Phosphate-buffered saline (PBS), 1×
Reagent Final concentration Quantity or volume
Phosphate-buffered saline 1× 9.9 g
Milli-Q water 1 L
* Autoclave the buffer and filter using a 0.22µm vacuum filter system. Store at 4 °C.
Nucleofection solution
Reagent Final concentration Quantity or volume
SE Cell Line NucleofectorTM solution (Lonza kit) 2.25 mL
Supplement buffer (Lonza kit) 0.5 mL
* Store the solution at 4 °C.
ICAM-1 stock solution
Reagent Final concentration Quantity or volume
Recombinant human ICAM-1 0.5 mg/mL 50 µg
LAL endotoxin-free water 100 µL
* Make small aliquots of the reconstituted stock solution and store them at -80 °C.
SDF1α stock solution
Reagent Final concentration Quantity or volume
Recombinant human SDF1α/CXCL12 0.1 mg/mL 10 µg
Endotoxin-free water 100 µL
* Make small aliquots of the reconstituted stock solution and store them at -80 °C.
Laboratory supplies
NuncTM glass-bottom confocal dishes (Thermo Scientific, catalog number: 150680)
T25 cell culture flasks (Thermo Scientific, catalog number: 156340)
1.5 mL microcentrifuge tubes (Tarsons, catalog number: 500010)
15 mL centrifuge tubes (Tarsons, catalog number: 546021)
Corning® Costar® tissue culture-treated 6-well plates (Merck, catalog number: CLS3516)
Corning® bottle-top vacuum filter system (Merck, catalog number: CLS431097)
Millex® PVDF syringe filter, pore size: 0.22 µm, diameter: 33 mm (Merck, catalog number: SLGVR33RS)
Nitrile gloves (Kimtech, catalog number: 55081)
Neubauer chamber
Equipment
AmaxaTM 4D-Nucleofector (Lonza)
Zeiss LSM 980 confocal microscope with 37 °C heating equipment and chamber
Laminar flow hood
37 °C, 5% CO2 incubator
pH meter
Refrigerated centrifuge
Software and datasets
Zen blue v3.3 (for imaging)
Fiji (for image analysis, https://fiji.sc/)
GraphPad Prism 8.0
Origin 2019b
Procedure
Preparation of Jurkat cell line for plasmid nucleofection
Passage approximately 0.1 × 106/mL Jurkat cells in T-25 flasks containing 5 mL of complete RPMI medium (see Recipe 1) between 36 and 48 h before transfection.
Do not allow confluency to reach more than 75% on the day of nucleofection.
Avoid using cells that have undergone more than 10 passages since revival.
Nucleofection of Piezo1-mCherry and actin-GFP plasmids
Pre-equilibrate post-nucleofection RPMI media (see Recipe 2) at 37 °C/5% CO2 in tissue-culture-treated 6-well plates. Add 2 mL of media per well.
Prewarm an aliquot of post-nucleofection RPMI media at 37 °C.
Switch on the Amaxa 4D-Nucleofector system at least 30 min before nucleofection.
Incubate the nucleofection solution at room temperature prior to nucleofection.
Count Jurkat cells using a Neubauer chamber.
Centrifuge Jurkat cells at 165× g for 3 min and remove media completely. Resuspend 2 × 106 cells in 100 µL of nucleofection solution.
Caution: Avoid leaving the cells in nucleofection solution for more than 5 min. Prolonged incubation of cells in the nucleofection solution causes a drastic reduction in cell viability.
Add Piezo1 mCherry-expressing plasmid and actin-GFP plasmid at a concentration of 2–3 µg/mL each into the Jurkat cell suspension. Mix gently. Ensure that the volume of plasmids does not exceed 10% of the final sample volume.
Transfer the plasmid–cell mix into nucleofection cuvettes and nucleofect using program CL-120, specific for the Jurkat cell line, in the Amaxa 4D-Nucleofector system.
Add 1 mL of prewarmed post-nucleofection RPMI medium to the cuvettes containing nucleofected cells and let it stand for 2 min.
Mix the cells by gentle pipetting and transfer the cells to culture plates containing equilibrated post-nucleofection RPMI medium. Seed the transfected cells at a density of 1 × 106 cells per well.
Incubate the cells at 37 °C/5% CO2 for 12 h. Wash the cells with 1× PBS at 165× g for 3 min and add fresh post-nucleofection RPMI medium for approximately an additional 24 h before imaging.
Preparation of cell chambers for live-cell imaging
Prepare 4 µg/mL of ICAM-1 solution in sterile and filtered 1× PBS from stock solution (see Recipes).
Coat glass-bottom confocal dishes (12 mm diameter) with 100 µL of ICAM-1 solution overnight at 4 °C.
Remove ICAM-1 solution. Wash the dish five times with 1× PBS (200 µL, 5 min each).
Remove PBS completely and allow the dishes to dry at room temperature for 2–3 h. The coated dishes can be stored at 4 °C in sealed conditions for approximately four weeks.
Preparation of Piezo1 mCherry/actin-GFP nucleofected Jurkat cells for live-cell imaging
Prewarm RPMI medium without phenol red at 37 °C (see Recipes).
Wash transfected Jurkat cells twice in 1× PBS at 165× g for 3 min. Resuspend cells in RPMI medium without phenol red for imaging. Measure transfection efficiency using flow cytometry (cells showing at least 30% transfection efficiency were used for downstream imaging).
Count cells using a Neubauer chamber.
Seed approximately 50,000 transfected Jurkat cells per 12 mm ICAM-1-coated confocal dish in 100 µL of imaging medium.
Allow the cells to adhere to the coated dishes for 1 h at 37 °C/5% CO2.
Note: The imaging medium contains 25 mM HEPES (pH 7.5). The buffering provided by HEPES operates independently of CO2/bicarbonate buffering system. Since our imaging system lacked CO2 supply, the addition of HEPES ensured the stability of medium pH during imaging.
Live-cell tracking of Piezo1 mCherry/actin-GFP expressing Jurkat cells
Prewarm the incubation chamber of the confocal microscope to 37 °C at least 30 min prior to imaging.
Transfer the confocal dish into the 37 °C incubation chamber of the confocal imaging setup. Incubate for an additional 15 min before tracking.
Note: Adequate prewarming and pre-incubation of cells within the microscope’s incubation chamber are critical to ensure the stability of temperatures and prevent thermal drift, which could otherwise cause gradual loss of focus or focal drift during imaging.
Add 0.1 µg/mL of recombinant SDF1α (diluted in imaging medium, see Recipe 8) to the dish 10–15 min prior to acquisition during incubation.
Duration and frequency of imaging for a time-lapse video depends on the nature of the events under examination. The time-lapse imaging of the representative cell was performed at intervals of 30 s for a total duration of 5 min. Z-stacks of 1 µm were acquired (total 12 stacks) at each frame. Images were acquired at 63×/1.4 magnification. For higher-speed imaging, 512 × 512 frames at a bi-directional scanning speed of 8–9 (maximum) was used.
Live-cell imaging without CO2 supplementation and adequate humidification should be limited to shorter durations of no more than 30–40 min.
Imaging sessions should be kept brief to prevent photo-bleaching of fluorescent proteins. In our study, we have restricted the maximum acquisition duration to 15–20 min, with intervals of 30 s between captures, and limited the total number of Z-stacks to 12–14. This approach helps minimize prolonged illumination and consequent photo-bleaching.
Note: The confocal dishes were sealed with parafilm to avoid the effect of air currents generated as a result of the heating mechanism and consequent drift.
Data analysis
Measuring Piezo1 and actin polarity in moving cells
Import the exported imaging files into Fiji (https://fiji.sc/).
In the Bio-Formats Import Options window, view the image as Hyperstack and in Composite color mode.
Duplicate the hyperstack. Specify the channels (fluorescence signals for Piezo1 mCherry and actin-GFP) and delete the stacks (or slices) and time frames that are not essential for the analysis from either/both ends of the range. Use this duplicated hyperstack for further analysis.
Select Image > Stacks > Z Projection > Max intensity (or other suitable method of Z projection). This will produce a time-lapsed 2D projection of your hyperstack. Use this resulting 2D video for further tracking and polarity analyses. The hyperstack is split into its constituent channels—Piezo1-mCherry (red) and actin-GFP (green)—for subsequent analysis (Image > Color > Split Channels) (Figure 1).
Figure 1. Generation of 2D time lapse videos by performing maximum intensity Z-projection of the multi-channels/multi-time-points hyperstacks. The multichannel 2D time hyperstack was split into two time-lapse videos corresponding to actin-GFP (green) and Piezo1 mCherry (red).
The analysis described below employs a custom code designed for the study by Liu et al. [31] to facilitate automated image processing of multiple regions of interest (ROIs) across multiple time frames of a single time-lapse video.
Optional: Gamma adjustment can be performed to control brightness and contrast of the images to aid the intensity thresholding. It employs a nonlinear mapping of the pixel intensity values. Gamma < 1 amplifies the faint pixels while reducing the intensities of the bright pixels, thereby making the faint pixels more visible. Conversely, Gamma >1 amplifies the bright pixels while further lowering the intensities of the faint pixels. Linear mapping is performed for gamma values equal to 1 (Figure 2A).
Select an ROI to measure background intensity and execute subtraction of background noise (Figure 2B).
Figure 2. Preprocessing of 2D time-lapse stack before ROI detection. A. Gamma adjustment enhances the contrast between the object of interest and background signal. B. Background signal correction involves selecting an ROI in the background region, measuring its average intensity, and subtracting the signal from the original image. C. The maximum intensity projection along the time axis generates a comprehensive view of the cell’s overall trajectory over the course of imaging. Defining this ROI facilitates easier detection and analysis of the cell within it across all time points.
Merge the two fluorescence channels. Perform a maximum intensity Z projection along the time axis to obtain a 2D image depicting the total extent of all the cell trajectories within the x-y frame. Use the freehand selection tool to manually draw ROIs encompassing the cell trajectories suitable for downstream analysis. Each ROI should contain a single-cell trajectory exhibiting signals in both fluorescence channels. Save these ROIs, as cells contained within will be used for tracking and polarity analyses across all time frames in the 2D time-lapse video (Figure 2C).
Create binary masks for each cell across all time frames within each delineated trajectory ROI, as specified above. Smooth the time stacks to blur the time-lapsed images and diminish speckled background noise. This facilitates image thresholding, ensuring accurate delineation of the specific cell ROI. Execute auto threshold by appending dark stack to the thresholding method, indicating that thresholding is performed on an image stack where the object of interest appears darker than the background. Utilize Covert to Mask and background = dark to generate binary masks of the cells, where darker regions (object of interest/cells) are interpreted as foreground while brighter regions are designated as image background (pixel values: 0 for background, 255 for foreground, for an 8-bit image) (Figure 3). These settings ensure precise segmentation of cells across all time frames. Utilize Fill Holes and Analyze Particles while specifying the maximum and minimum cell area to generate outlines of ROIs corresponding to each cell mask. Save these ROIs (Figure 3).
Figure 3. Mapping out cell ROIs. A. A generalized analysis scheme for ROI detection using thresholding and creation of binary masks (0–255). B. Resulting binary masks and outline of analyzed cell ROI at each time frame. Both actin-GFP and Piezo1-mCherry channels have been shown for the detected ROI.
Using Set Measurements, analyze the following parameters of each cell ROI:
Area
Center of Mass (intensity-weighted average of x and y of all pixels) = XM and YM
Centroid (average of x and y coordinates of all pixels) = X and Y
Shape descriptors = Circularity, Aspect Ratio, Round, and Solidity
Fit ellipse = lengths of Major (primary) and Minor (secondary) axis, and Angle between the primary axis of the cell (the major axis of the fitted ellipse) and a line parallel to the x-axis of the image.
Store these values corresponding to each cell in every time frame as separate arrays.
Convert image stacks to 32-bit. Perform Fit Ellipse on the cell ROI to extract the shape parameters, then bisect each cell ROI into equal parts along the minor axis using the following calculations:
From the Angle value given by ImageJ, calculate the angle subtended by the minor axis of the fitted ellipse on the x-axis of the image (Figure 4A).
φ
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θ = angle perpendicular to the direction of the major axis of ellipse, i.e., the minor axis
Draw the minor axis from the ellipse’s centroid to one edge of the ROI by specifying coordinates of the edge (xbl1, ybl1) (Figure 4A).
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Draw minor axis from the centroid to the other edge of the ROI by specifying coordinates of the other edge (xbl2, ybl2) (Figure 4A).
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widthPxl and heightPxl = width (along x-axis) and height (along y-axis) of the ROI.
Xc[j] and Yc[j] = centroids of the ROIs (no. of ROIs = j).
Major = length of the major axis of the ellipse.
Draw a line connecting xbl1, ybl1 and xbl2, ybl2, which will bisect the ellipse along the minor axis of the ROI ellipse. The length of the bisecting line along the minor axis is equal to the length of the major axis so that the straight line completely passes over the ellipse’s edges. Without the overlap, gaps remain between the endpoints of the line and the ellipse’s edges, which does not allow selecting either half of the ellipse using the Wand tool later. Do this for all ROIs corresponding to cell masks in each time frame (Figure 4A, B).
Figure 4. Bisecting cell ROI into front and back halves. A. A cell ROI is fitted into an ellipse and its shape parameters are measured. The centroids (XC, YC) of the cell ROI at subsequent time frames (Tj-1 and Tj) are used to calculate the cell displacement. The ROI is split into two halves along the minor axis (perpendicular to the major axis). θ= angle perpendicular to the major axis, ϕ= angle (radians) subtended by the minor axis. To determine the back and front half of the bisected ROI, dist1 and dist2 are calculated. dist1 < dist2. dist1 corresponds to the back half and dist2 corresponds to the front half of the bisected ROI. (xh1, yh1): half-centers of front half; (xh2, yh2): half-centers of back half. B. The cell ROIs across time frames (F0–F10) were fitted into an ellipse. The ellipse was bisected along the minor axis (perpendicular to the major axis) to generate two halves of the moving cell.
To define the front and back halves of the bisected moving cell
Measure the coordinates of the half-centers of each half of the cell. The half-center of each cell-half lies at the mid-point between the centroid of the whole cell and the centers of each half-cell (Figure 4A):
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xh1, yh1 and xh2, yh2 are half-center coordinates of each half of the ROI.
Note: Ideally, any point within each bisected ROI would enable Fiji to use the Wand tool and detect each bisected ROI. To avoid randomness, we chose the “half-center” that lies closer to the bisection line, thereby increasing the probability of finding positive fluorescence pixels that will facilitate detection (Figure 4A).
Calculate the distance of the half-center points of each of the above-calculated half-ellipses from the ellipse’s centroid in the preceding frame in the trajectory. The closest half is the back half of the cell while the farthest half is the front half of the cell (Figure 4A).
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Using the whole cell masks or ROIs stored earlier (step 8) and the calculated half-center coordinates of each half of the ellipse (xh1, yh1 and xh2, yh2), measure the ROIs corresponding to each half of the ellipse using the Wand tool. Measure the raw integrated density (RawIntDen) of both front and back halves of the ellipse.
Measure the polarity index by determining the ratio of the raw integrated density of the front and back halves of the cell (F/B). File S1 contains the results of the analysis. A polarity index greater than 1 signifies skewed frontal polarization and distribution of the fluorescent signals (Figure 5A). A higher Piezo1 F/B polarity compared to actin aligns with the leading-edge distribution of Piezo1, as observed in Liu et al. [31].
Note: Perform the above analysis steps for both fluorescent channels to calculate F/B polarity of both Piezo1 mCherry and actin-GFP.
Figure 5. Measuring live-cell polarity of Piezo1 mCherry and actin-GFP in chemokine-stimulated Jurkat cells. A. Front/back (F/B) polarity of Piezo1 and actin of a single cell. Each point corresponds to F/B value at a specific time frame of a single moving cell. B. Polar plots depicting relative angles of Piezo1 (upper panel) and actin (lower panel). The relative angles correspond to a single cell against different time frames. C. Vector plot of a single cell depicting the direction of cell trajectory (blue), Piezo1 polarization (red), and actin polarization (green) at each time frame.
The relative angle of polarity or angular distribution of fluorescent signals with respect to displacement of cell can also be measured. Perform the calculation below for all the ROIs (no. of ROIs = j), corresponding to varying cell positions across different time frames.
Using the atan2 function as below, the polarization angle of fluorescent signals can be calculated with respect to the cell center. Convert the angles to degrees (
t
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XcMA, YcMA = center of mass (intensity-weighted spatial coordinates)
Xc, Yc = centroid (spatial coordinates of center of cell area)
Similarly, the displacement magnitude and the displacement angle (direction of trajectory) can also be calculated. Convert the angles to degrees.
D
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j-1, j = two subsequent time frames.
Calculate the relative angle of polarization of fluorescence intensity with respect to the trajectory of displacement by subtracting the displacement angle from the polarization angle (Figure 5B).
Note: Perform the above analysis steps for both fluorescent channels to obtain polarization angles of both Piezo1 mCherry and actin-GFP across all time frames.
Utilize the Arrow tool alongside a suitable scaling method (bilinear interpolation) to depict vectors representing the polarization angles for all the ROIs across time frames (Figure 5C).
Visualization software such as GraphPad Prism 8 and Origin 2019b can be used to plot the F/B ratios along with polarization angles of both Piezo1 mCherry and actin-GFP signals. Polar plots can be generated to examine the distribution of Piezo1 and actin relative angles. The relative angle values calculated for all cells at each time frame can be binned into regular intervals ranging from -180° to +180°. The frequency distribution can then be subsequently plotted.
Validation of protocol
The protocol was used and published in the following research article:
Liu et al. [31]. Piezo1 mechanosensing regulates integrin-dependent chemotactic migration in human T cells. Experimental setup and analysis were used for figures 6D, E, and figure 6 – figure supplement 1B and C.
General notes and troubleshooting
General notes
Nucleofection often leads to significant cell death. To enhance transfection efficiencies, ensure that the cell line is healthy and has undergone no more than 10 passages. Increase cell viability by using prewarmed and CO2-equilibrated medium. Avoid using confluent cells for transfection, as this can lead to decreased viability and transfection efficiencies. Additionally, minimizing cell death can be achieved by increasing FBS content and removing antibiotics in the post-nucleofection media.
All media and buffers used for cell preparation and imaging should be filtered using 0.22 µm filters. This step helps reduce debris contaminants, which can otherwise compromise imaging quality by contributing to elevated background noise and signal artifacts.
It is crucial to maintain stable temperatures throughout the imaging process. Temperature fluctuations and other mechanical disturbances can cause substantial focal drift during imaging, rendering the resulting images unsuitable for subsequent analysis. While there are tools and algorithms available to compensate for manual focal drift, it is advisable to minimize disturbances to capture high-quality time-lapse images. Prior heating of the enclosed imaging chamber minimizes such temperature- and mechanical-induced variations.
Images should be carefully monitored for any evidence of photobleaching and consequent loss of signal during the course of live-cell imaging. Minimizing the total duration of acquisition, along with the number of frames and/or Z-stacks, reduces the frequency of illumination of the sample and mitigates resulting photobleaching.
Additionally, given the dynamic nature of live-cell imaging, it is crucial to capture each frame as quickly as possible to minimize significant cell movement within a single frame. This can be achieved by employing bi-directional high-speed scanning at a moderate resolution of 512 × 512 pixels and enhancing magnification for faster scanning of smaller areas.
Efficient segmentation of cells to determine ROIs for subsequent analysis relies on robust pre-processing of the images, including enhancing image contrast, smoothing for background noise, and correcting background intensities. There is no singular method that can be considered ideal for achieving optimal results in these steps. The choice of method depends on various parameters, including image quality, signal strength, and background noise level. Numerous tools and algorithms are available for the purposes of reducing background and thresholding (delineation of cell ROIs), which should be carefully tried and implemented to obtain accurate results.
Acknowledgments
We express our sincere gratitude towards all co-authors of the corresponding study [31] for their significant contributions and support. The study was funded by the Council of Scientific and Industrial Research, India (Grant no. FBR MLP-140). D.G. was additionally supported by the Swarnajayanti Fellowship award granted by the Department of Science and Technology, Government of India. Special thanks are extended to Dr. Charles Cox at the Victor Chang Institute of Cardiac Research, Australia, for generously providing the Piezo1 constructs.
Competing interests
The authors declare no competing financial interests.
Ethical considerations
There are no ethical considerations associated with this protocol.
References
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Supplementary information
The following supporting information can be downloaded here:
File S1
File S1.xlsx contains source data for Figures 6A–B. It contains the Fiji analysis outputs: ROI parameters(area, circularity), trajectory analysis(displacement magnitude, displacement angle), polarization angles and relative polarization angles for Piezo1(P) and actin(A), raw integrated densities of front and back-halves of the ROI, along with front/back(F/B) ratios of Piezo1 and actin. Each data point corresponds to a single cell at different time frames. Calculation of polar angular distribution has also been added to the file.
Article Information
Publication history
Received: May 22, 2024
Accepted: Aug 12, 2024
Available online: Sep 17, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Fluorescence Lifetime-Assisted Probing of Protein Aggregation with sub-Organellar Resolution
KG Karnika Gupta
DM Daniel C. Maddison
EM Eduardo P. Melo
AC Ana Rosa M. da Costa
EA Edward Avezov
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5080 Views: 464
Reviewed by: Elena A. OstrakhovitchJose Martinez Hernandez Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications May 2022
Abstract
Protein misfolding fuels multiple neurodegenerative diseases, but existing techniques lack the resolution to pinpoint the location and physical properties of aggregates within living cells. Our protocol describes high-resolution confocal and fluorescent lifetime microscopy (Fast 3D FLIM) of an aggregation probing system. This system involves a metastable HaloTag protein (HT-aggr) labeled with P1 solvatochromic fluorophore, which can be targeted to subcellular compartments. This strategy allows to distinguish between aggregated and folded probe species, since P1 fluorophore changes its lifetime depending on the hydrophobicity of its microenvironment. The probe is not fluorescence intensity-dependent, overcoming issues related to intensity-based measurements of labeled proteins, such as control of probe quantity due to differences in expression or photobleaching of a proportion of the fluorophore population. Our approach reports on the performance of the machinery dealing with aggregation-prone substrates and thus opens doors to studying proteostasis and its role in neurodegenerative diseases.
Key features
• Aggregation state: Tracks aggregate formation and disaggregation with pulse-chase experiments
• Sub-organellar resolution: Pinpoints and allows control of aggregate location within the cell, exceeding traditional techniques
• Quantitative analysis: Measures aggregate load through image analysis
• Methodology:
Metastable HaloTag variant labeling with a solvatochromic small-molecule reporter ligand
High-resolution confocal microscopy coupled with FLIM for aggregate identification and localization
Image analysis for aggregate quantification and distribution within the ER
Pulse-chase experiments to track aggregates
Keywords: HT-aggrER ER FLIM 3D-FLIM Disaggregation
Graphical overview
Background
Protein misfolding and aggregation are hallmarks of numerous neurodegenerative diseases, including Alzheimer’s, Parkinson’s, and amyotrophic lateral sclerosis (ALS) [1]. Unraveling how the protein folding and quality control system handles aggregation-prone substrates is key to understanding why protein folding mechanisms fail, leading to age-associated aggregate accumulation in disease [2].
Proteostasis, the intricate network governing protein homeostasis within cells, relies on the coordinated efforts of various organelles and molecular machinery. Each cellular compartment contributes uniquely to the folding, trafficking, and degradation of proteins, with specific proteins and mechanisms orchestrating these processes. The endoplasmic reticulum (ER) stands as a primary site for protein folding and quality control. Within its lumen, chaperone proteins such as BiP/GRP78 and calnexin/calreticulin assist in guiding nascent polypeptides into their native conformations [3]. The ER also houses the unfolded protein response (UPR), a signaling pathway activated in response to ER stress, which coordinates adaptive measures to restore proteostasis. In the cytosol, molecular chaperones such as Hsp70 and Hsp90 safeguard protein folding, preventing misfolding and aggregation [4]. The ubiquitin-proteasome system (UPS) plays a vital role in protein degradation, tagging misfolded or damaged proteins with ubiquitin for proteasomal degradation. Mitochondria harbor molecular chaperones like Hsp60 and Hsp70, which facilitate the folding of mitochondrial proteins [5].
Understanding the specific proteins and mechanisms operating within each cellular compartment is essential for deciphering the molecular underpinnings of proteostasis and its dysregulation in disease. By elucidating these intricate pathways, researchers can identify novel therapeutic targets and develop interventions aimed at restoring protein homeostasis in various pathological conditions. Thus, the ability to probe the performance of the proteostasis machinery through detecting aggregation with organelle-level resolution is essential.
Existing tools for studying protein aggregation offer valuable insights, but they come with limitations. While immunoblots can detect total protein levels, they lack spatial information, preventing precise localization of aggregates within the cell beyond crude fractions. Electron microscopy provides high resolution but requires harsh fixation procedures, hindering live-cell studies and potentially altering the delicate ultrastructure of subcellular organelles. Fluorescent microscopy offers the advantage of live-cell imaging but often struggles to distinguish between misfolded aggregates and native protein structures, particularly within crowded confines of organelles such as the ER.
Our protocol addresses these limitations by introducing a subcellular resolution system combining confocal super-resolution microscopy with fast fluorescence lifetime imaging microscopy (FLIM) [6] by taking advantage of a metastable probe labeled with a solvatochromic fluorophore [7] that changes its lifetime depending on aggregation state [8]. Solvatochromic dyes change fluorescent properties based on their microenvironment, namely by polarity. P1 solvatochromic fluorophore is a modified BODIPY with a chloro-alkane chain ligand designed to covalently bind to Halotag, a modified bacterial haloalkane dehalogenase. HT-aggrER is Halotag further modified by the addition of signal peptide and KDEL motif for ER localization and point mutations that make the otherwise metastable protein more aggregate-prone [7]. When P1 binds to and labels HT-aggrER in living cells, its fluorescence lifetime is influenced by protein aggregation status, with a longer lifetime observed in aggregated proteins, which can be promoted by heat shock [8]. Monomeric protein can also be visualized by P1 labeling, but with a shorter lifetime, allowing for optical separation of protein species and thus real-time monitoring of protein aggregation status. HT-aggrER can also be labeled by fluorescent ligands such as TMR and Janelia Fluor 646, which label non-aggregated HT-aggr more brightly than P1 but are quenched and not fluorescent inside hydrophobic aggregates. This provides another degree of flexibility in the labeling of aggregated and soluble protein species. Resolving protein aggregates with this level of detail in live cells is affordable through the critical improvement in speed of time-correlated single-photon counting (TCSPC) fast FLIM on the Leica SP8 FALCON [9,10], which detects photons at ~80 mega counts per second and plots average arrival time for each pixel in near-real time without exponential fitting [11]. LIGHTNING deconvolution applies the Richardson-Lucy algorithm and the underlying principles of point spread function to remove background and out-of-focus signal from a 3D confocal image voxel by voxel, based on an adaptative decision mask. Implementing fast FLIM alongside LIGHTNING deconvolution of correlated confocal images allows for the localization of the FLIM data to sub-diffraction limit resolution counter images. This combination unlocks new avenues for studying protein aggregation state with subcellular resolution, offering several key advantages over existing methods. Beyond providing unparalleled insights into ER protein aggregation, our protocol is applicable for exploring other areas of cellular proteostasis, using the aggregation probe variants targeted to other cellular compartments. Additionally, this system could be used for live-cell drug screening, allowing researchers to rapidly assess the effectiveness of potential therapeutics in preventing or disassembling protein aggregates. The versatile protocol described here thus fills a gap in existing methodologies by offering high-resolution and quantitative insights into the complex protein aggregation process. Its implementation can help to gain a deeper understanding of protein misfolding-related diseases and the development of therapeutic strategies against them.
Materials and reagents
Biological materials
Chinese hamster ovary cells (CHO-K1, ATCC CCL-61) with stable expression of HT-aggrER (HaloTag with M21K F86L mutations)
Reagents
Nutrient mixture F12 Ham [Sodium bicarbonate (+), L-Glutamine (-)] (Sigma Merck, catalog number: N4888-500ML)
Fetal bovine serum (FBS) (Sigma, catalog number: F9665-500ML)
Penicillin-streptomycin (Pen/strep) (Thermo Scientific, catalog number: 15140122)
L-Glutamine (Thermo Scientific, catalog number: 25030024)
P1 solvatochromic fluorophore (prepared in-house—protocol described in General Notes and Supplementary File 1)
1× Dulbecco’s phosphate-buffered saline (DPBS) (Thermo Fisher Scientific, catalog number: 14190169)
0.05% Trypsin-EDTA (1×) (Thermo Fisher, catalog number: 25200056)
Janelia Fluor 646 HaloTag ligand (Promega, catalog number: GA1120) (see General Notes for further discussion of the use of commercially available HaloTag ligands to label HT-aggr)
Solutions
Complete nutrient mixture F12 Ham (see Recipes)
P1 staining solution (see Recipes)
Janelia Fluor 646 HaloTag staining solution (see Recipes)
Recipes
Complete nutrient mixture F12 Ham
Reagent Stock concentration Final concentration Amount
Nutrient mixture F12 Ham 100× 90% 500 mL
FBS 100× 10% 50 mL
Pen/strep 10,000 U/mL 1% 5.5 mL
L-Glutamine 200 mM 1% 5.5 mL
P1 staining solution
Reagent Final concentration Amount
Complete nutrient mixture F12 Ham NA 500 μL
P1 (2 mM) 4 μM 1 μL
Janelia Fluor 646 HaloTag staining solution
Reagent Final concentration Amount
Complete nutrient mixture F12 Ham NA 500 μL
Janelia Fluor 646 HaloTag (200 mM) 200 μM 0.5 μL
Laboratory supplies
Coverslip bottomed dishes, IBIDI (IBL Labor GmbH, catalog number: D35C4-20-1-N)
10 cm dish (Falcon, catalog number: 353003)
Centrifuge tube, 15 mL (Appleton Woods Ltd, catalog number: AB031)
Equipment
Leica Stellaris 8 FALCON FLIM Microscope (Leica, Wetzlar, Germany, model: STELLARIS8)
Benchtop centrifuge (Eppendorf, model: 5810 R, product code: 12813252)
Software and datasets
Leica LAS X (v5.2.2, 01.02.2024)
Icy (v2.5.2.0, 08.02.2024)
Procedure
Preparation of CHO HT-aggrER cells for 3D-FLIM imaging
Grow cells in complete nutrient mixture F12 media (see Recipe 1) in a cell culture incubator at 37 °C and 5% CO2 until cells reach 70%–80% confluency in a 10 cm dish.
Aspirate complete nutrient mixture F12 from cells, wash once with 1× DPBS, add 1 mL of Trypsin-EDTA, and return cells to incubator for ~3 min.
Once detached, add 3 mL of complete nutrient mixture F12 media to cells and pipette up and down 10 times to make a single-cell suspension.
Count cells using a hemacytometer or automated cell counter.
Transfer cells to a 15 mL falcon tube and centrifuge at 200× g for 5 min.
Aspirate supernatant and resuspend cells with complete media at 1×106 cells/mL.
Seed cells at 3,000 cells/cm2 in a coverslip glass-bottomed cell culture vessel.
Return to incubator and grow to ~70% confluency.
Pulse labeling of HT-aggrER with P1 solvatochromic fluorophore
Dilute P1 stock to 4 μM final concentration in prewarmed complete nutrient mixture F12 media (see Recipe 2).
Take cells grown from Section A and replace media with P1 staining solution.
Return cells to the incubator for 45 min.
Remove P1 staining solution and wash with prewarmed complete media.
Replace with prewarmed complete nutrient mixture F12 media.
Now that HT-aggER has been pulse-labeled, the cells can be subjected to a treatment of choice prior to imaging to observe the effect of the treatment on protein aggregates over time. For example, we have observed that mild ER stress induced by tunicamycin or thapsigargin results in a decrease in long-lifetime aggregates [8].
Counter-labeling with 646 HaloTag ligand to visualize ER
Directly before imaging, newly synthesized HT-aggrER can be pulse-labeled with Janelia Fluor 646 HaloTag ligand (see Recipe 3) (this only labels non-aggregated HT and thus serves as a counterstain for the endoplasmic reticulum).
Stain cells with Janelia Fluor 646 HaloTag staining solution for 45 min prior to imaging.
Remove Janelia Fluor 646 HaloTag staining solution and wash twice with prewarmed 1× DPBS.
Add 500 μL of fresh complete nutrient mixture F12 media to the well and subject to imaging. We have not observed any difference in fluorescence lifetime of our probe in media with or without phenol red or serum.
FLIM live cell imaging of P1-label
Warm the microscope stage to 37 °C and set CO2 level to 5%.
Place the cell sample on the microscope stage.
Locate and focus on cells through eyepieces using brightfield illumination to minimize bleaching of P1 fluorophore.
Set microscope parameters in LAS X software as follows:
FLIM mode
Laser 1: Excitation/emission 440/450–500 nm, pulsed laser (see notes; for P1 imaging, faster laser repetition rate may result in apparent shortening of fluorescence lifetime)
Laser 2: Excitation/emission 646/655–750 nm (for 646 HaloTag ligand imaging, follow steps C1–4)
HyDx detector in counting mode
Frame size 512 × 512 pixels
Scan speed 400 Hz
Count frame average of 15–20
Set Z-stack for the cell (begin and end)
Start image acquisition (Start Experiment button).
Confocal live imaging in the LIGHTNING mode of 646 HaloTag ligand
Immediately after FLIM imaging of cells, deselect FLIM mode and select LIGHTNING mode of LAS X software.
Do not change the Z-stack or frame-size settings acquired in FLIM mode.
Start image acquisition (Start experiment button).
Image rendering
Raw FLIM image rendering (Figure 1) produces a map of mean arrival time for each pixel of the acquired Fast FLIM image. The values are color-coded to distinguish protein aggregates with longer lifetime.
Select FLIM mode.
Set the lifetime range (2.5–6 ns). In this range, pixels with a lifetime of >4.5 ns will appear in yellow/red, allowing aggregates to be distinguished as red puncta from green-folded probe species. We also observe concentrated areas of shorter lifetime (blue) signal. Further work is required to understand the biological relevance of these regions. Since their lifetime does not correspond to that of aggregated HT-aggr such as that induced by heat shock [8], we do not consider these as aggregates when we perform image analysis.
Export image in TIFF or PNG format (ensure “Save Palette Image as RGB” is selected to maintain the defined FLIM lookup table in the saved image).
Export in the format of 3D-movie.
Figure 1. Illustration of Raw 3D FLIM image. A. Settings for acquiring FLIM image using Leica Stellaris 8 confocal. B. Micrograph of 2D plane of CHO M21K F86L cells acquired using settings as in A. C. Micrograph of 3D FLIM image of cell as in B. White arrows point toward aggregates. Images are at 63× magnification and were acquired using Leica Stellaris 8 FALCON. Scale bar: 20 µm. Lifetime range: 2.5–6.0.
Aggregate size analysis of FLIM data in Icy Software (Figure 2)
Open the exported TIFF or PNG FLIM image files in Icy Software (v2.5.2) [12].
Select Build RGB Image.
Deselect Ch1 and Ch2 (green and blue) channels to leave only red pixels, reflecting regions of fluorescence lifetime >4.5 ns.
Open the Aggregates Detector protocol.
Adjust the Threshold and Min volume settings to identify aggregates of the desired intensity and size. We typically use threshold = 125 and min volume = 5 to avoid quantification of diffuse signal.
Run Aggregates Detector and then navigate to the ROI tab on the right-hand side of the viewer. There will be a list of ROIs corresponding to aggregates with corresponding shape descriptors such as area. This table can be exported to Excel for subsequent analysis of shape descriptor metrics (Figure 2D).
Figure 2. Schematic representation of calculating frequency distribution of aggregates’ sizes extracted from FLIM using Icy software. A. 2D FLIM image is imported into Icy and converted to RGB image. B. Green and blue channels are deselected to leave only red pixels corresponding to longer lifetime areas. C. Aggregates Detector plugin is utilized to identify and extract shape descriptors from long-lifetime aggregates. D. Representative graph of aggregate size in correspondence to its relative frequency.
3D FLIM image rendering (Figure 3). This produces a 3D image of the FLIM map overlayed with the high-resolution ER counter image. This should be used for visual purposes only and not for quantitative analysis of aggregates.
Select 3D tool on LASX software.
Open 3D FLIM and corresponding LIGHTNING confocal images.
Under the Processing tab, select the concatenate function.
Select both 3D FLIM and LIGHTNING images to concatenate by channel.
Apply concatenation.
Deselect blue and green channels.
Set minimum intensity threshold to 40.
Using movie editor, export image in movie mode (Video 1).
Figure 3. Processed high-resolution 3D FLIM image. A. Micrographs of FLIM and ER acquired using FLIM mode and LIGHTNING confocal mode, respectively. B. 3D overlay of micrograph of FLIM (red/green) and ER (grey) image. Red spots represent aggregated probes. Images are at 63× magnification and were acquired using Leica Stellaris 8 FALCON. Scale bar: 20 μm. Lifetime range: 2.5–6.0 ns.
Video 1. 3D fast-FLIM image of P1-labeled HT-aggrER aggregates acquired by FLIM (red), overlayed with HaloTag ligand 646-labeled newly synthesized HT-aggrER acquired by confocal (grey). Red spots represent longer lifetime aggregates.
Validation of protocol
This protocol has been used and validated in the following research article [8]:
Melo et al. (2022). Stress-induced protein disaggregation in the endoplasmic reticulum catalysed by BiP. Nat Commun (Figure 2, panel e).
General notes and troubleshooting
Here, the protocol has been demonstrated on the CHO-K1 cell line harboring stable integration of the HT-aggrER construct with M21K F86L mutations. This cell line and other mutant cell lines used in Melo et al. [8] are available upon request.
Alternatively, the plasmids used in Melo et al. [8] and this protocol are available via Addgene. We recommend using stable cell lines harboring the plasmids, as the formation of protein aggregates occurs several days after expression and thus cannot be visualized transiently.
P1 solvatochromic HaloTag ligand was synthesized by Dr. Ana Costa at the University of Algarve, Portugal, and is available upon request to Dr. Edward Avezov. A full methodology for P1 synthesis can be found in Supplementary File 1.
Other HaloTag ligands can be used to label HT-aggrER but behave differently from P1 in terms of binding propensity and fluorogenicity to aggregates. For example, Janelia Fluor HaloTag TMR and 646 do not label the aggregates we observe with long fluorescence lifetime as measured by P1 labeling. However, diffuse HT-aggr and other short-lifetime puncta are labeled by these ligands.
Overnight P1 labeling can also be performed.
Before staining cells with HaloTag ligand 646, wash away P1 stain and allow cells to grow in fresh nutrient mixture F12 medium for a minimum of 4 h. This is to allow sufficient time for new HT-aggrER to be synthesized after washing excess P1 label away, which will be available to covalently bind to HaloTag ligand 646 and thus act as a counterstain for the ER.
Fluorescence lifetime is dependent on the frequency of the laser used. For the original research article in which the protocol was validated, Zeiss LSM710 with 20 MHz laser was used for 2D FLIM. For this Bio-protocol, Leica Stellaris 8 FALCON FLIM microscope with 80 MHz laser was used to acquire 3D FLIM images. The differing laser frequencies have an impact on the observed fluorescence lifetime of the P1-labeled probe.
3D FLIM is enabled by improved imaging speeds of Fast-FLIM, made possible by reduced detector dead time, which allows for ~80 mega counts per second photon detection speeds. Other microscope systems capable of TCSPC FLIM with comparable detection speed, such as Becker & Hickl FASTAC FLIM or picoQuant MultiHarp 150, should thus be able to reproduce our approach when combined with a sub-diffraction limit resolution confocal system such as Airyscan or stimulated emission depletion (STED).
Acknowledgments
E.A. in this work is supported by the UK Dementia Research Institute through UK DRI Ltd, principally funded by the Medical Research Council, The Evelyn Trust and Alzheimer’s Society. K.G. is supported by Cambridge Indian Ramanujan Scholarship. E.P.M. received Portuguese national funds from FCT - Foundation for Science and Technology through projects UIDB/04326/2020 (DOI:10.54499/UIDB/04326/2020), UIDP/04326/2020 (DOI:10.54499/UIDP/04326/2020) and LA/P/0101/2020 (DOI:10.54499/LA/P/0101/2020). This protocol is adapted from Melo et. al [8].
Competing interests
The authors declare no competing interests.
References
Scheres, S. H. W., Ryskeldi-Falcon, B. and Goedert, M. (2023). Molecular pathology of neurodegenerative diseases by cryo-EM of amyloids. Nature. 621(7980): 701–710. https://doi.org/10.1038/s41586-023-06437-2
Hipp, M. S., Kasturi, P. and Hartl, F. U. (2019). The proteostasis network and its decline in ageing. Nat Rev Mol Cell Biol. 20(7): 421–435. https://doi.org/10.1038/s41580-019-0101-y
Wang, M. and Kaufman, R. J. (2016). Protein misfolding in the endoplasmic reticulum as a conduit to human disease. Nature. 529(7586): 326–335. https://doi.org/10.1038/nature17041
Rutledge, B. S., Choy, W. Y. and Duennwald, M. L. (2022). Folding or holding?—Hsp70 and Hsp90 chaperoning of misfolded proteins in neurodegenerative disease. J Biol Chem. 298(5): 101905. https://doi.org/10.1016/j.jbc.2022.101905
Jebara, F., Weiss, C. and Azem, A. (2017). Hsp60andHsp70Chaperones: Guardians of Mitochondrial Proteostasis. In Wiley, Encyclopedia of Life Sciences (1st ed., pp. 1–9). https://doi.org/10.1002/9780470015902.a0027152
Holcman, D., Parutto, P., Chambers, J. E., Fantham, M., Young, L. J., Marciniak, S. J., Kaminski, C. F., Ron, D. and Avezov, E. (2018). Single particle trajectories reveal active endoplasmic reticulum luminal flow. Nat Cell Biol. 20(10): 1118–1125. https://doi.org/10.1038/s41556-018-0192-2
Liu, Y., Fares, M., Dunham, N. P., Gao, Z., Miao, K., Jiang, X., Bollinger, S. S., Boal, A. K. and Zhang, X. (2017). AgHalo: A Facile Fluorogenic Sensor to Detect Drug‐Induced Proteome Stress. Angew Chem Int Ed. 56(30): 8672–8676. https://doi.org/10.1002/anie.201702417
Melo, E. P., Konno, T., Farace, I., Awadelkareem, M. A., Skov, L. R., Teodoro, F., Sancho, T. P., Paton, A. W., Paton, J. C., Fares, M., et al. (2022). Stress-induced protein disaggregation in the endoplasmic reticulum catalysed by BiP. Nat Commun. 13(1): 2501. https://doi.org/10.1038/s41467-022-30238-2
Konno, T., Parutto, P., Crapart, C. C., Davì, V., Bailey, D. M. D., Awadelkareem, M. A., Hockings, C., Brown, A. I., Xiang, K. M., Agrawal, A., et al. (2024). Endoplasmic reticulum morphology regulation by RTN4 modulates neuronal regeneration by curbing luminal transport. Cell Rep. 43(7): 114357. https://doi.org/10.1016/j.celrep.2024.114357
Crapart, C. C., Scott, Z. C., Konno, T., Sharma, A., Parutto, P., Bailey, D. M. D., Westrate, L. M., Avezov, E. and Koslover, E. F. (2024). Luminal transport through intact endoplasmic reticulum limits the magnitude of localized Ca2+ signals. Proc Natl Acad Sci U S A. 121(13): e2312172121. https://doi.org/10.1073/pnas.2312172121
Alvarez, L. A., Widzgowski, B., Ossato, G., van den Broek, B., Jalink, K., Kuschel, L., Roberti, M. J., Hecht, F. (2019). Application note : SP8 FALCON : A novel concept in fluorescence lifetime imaging enabling video-rate confocal FLIM. Nat Methods. Available at: https://www.nature.com/articles/d42473-019-00261-x
de Chaumont, F., Dallongeville, S., Chenouard, N., Hervé, N., Pop, S., Provoost, T., Meas-Yedid, V., Pankajakshan, P., Lecomte, T., Le Montagner, Y., et al. (2012). Icy: an open bioimage informatics platform for extended reproducible research. Nat Methods. 9(7): 690–696. https://doi.org/10.1038/nmeth.2075
Supplementary information
The following supporting information can be downloaded here:
Supplementary File 1
Article Information
Publication history
Received: May 10, 2024
Accepted: Aug 8, 2024
Available online: Sep 29, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Biophysics > Microscopy
Neuroscience > Nervous system disorders > Neurodegeneration
Cell Biology > Cell imaging > Live-cell imaging
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Alternative Method for Obtaining Human-Induced Pluripotent Stem Cell Lines and Three-Dimensional Growth: A Simplified, Passage-Free Approach that Minimizes Labor
MT Masaya Tsukamoto
TK Tomoyuki Kawasaki
AU Akihiro Umezawa
Hidenori Akutsu
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5081 Views: 480
Reviewed by: Alessandro DidonnaMasashi ToyodaGiovanna Piovani
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Original Research Article:
The authors used this protocol in Regenerative Therapy Dec 2024
Abstract
Induced pluripotent stem cells (iPSCs) hold significant promise for numerous applications in regenerative medicine, disease modeling, and drug discovery. However, the conventional workflow for iPSC generation, with cells grown under two-dimensional conditions, presents several challenges, including the need for specialized scientific skills such as morphologically assessing and picking colonies and removing differentiated cells during the establishment phase. Furthermore, maintaining established iPSCs in three-dimensional culture systems, while offering scalability, necessitates an enzymatic dissociation step for their further growth in a complex and time-consuming protocol. In this study, we introduce a novel approach to address these challenges by reprogramming somatic cells grown under three-dimensional conditions as spheres using a bioreactor, thereby eliminating the need for two-dimensional culture and colony picking. The iPSCs generated in this study were maintained under three-dimensional conditions simply by transferring spheres to the next bioreactor, without the need for an enzymatic dissociation step. This streamlined method simplifies the workflow, reduces technical variability and labor, and paves the way for future advancements in iPSC research and its wider applications.
Key features
• Establishment of induced pluripotent stem cells in a three-dimensional environment.
• Maintenance and cryopreservation of iPSCs without the need for a dissociation step.
Keywords: Bioreactor Enzyme-free passaging iPSCs Reprogramming 3D culture
Graphical overview
Background
Induced pluripotent stem cells (iPSCs), derived from somatic cells, exhibit characteristics similar to those of embryonic stem cells. This makes them valuable tools in regenerative medicine and disease modeling [1]. However, the process of generating iPSCs is fraught with challenges, especially during the establishment phase, which requires specialized technical skills [2,3]. During the reprogramming phase, partially reprogrammed cells, re-differentiated cells, and non-reprogrammed cells all typically proliferate. However, a common issue is trying to attain spontaneous differentiation until cells achieve stable pluripotent states. It is therefore necessary to select and isolate iPSC colonies with optimal morphology during the initial passages. Such complexities contribute to the difficulties and inherent variability of iPSC cultures, which often depend on the technician's skills.
Traditionally, iPSCs are maintained under two-dimensional conditions using a specialized medium and feeder cells or an extracellular matrix reagent. This approach hampers scalability, which is crucial for iPSC downstream applications. Recently, the maintenance of iPSC lines under three-dimensional (3D) conditions has been shown to offer scalability. This has been achieved using agitation rotor machines or bioreactors [4,5]. However, the enzymatic dissociation step required for single-cell cultures under 3D conditions is both time-consuming and complex [4,6].
In particular, the generation of personalized iPSCs for use in regenerative medicine necessitates a simplified and streamlined workflow for iPSC studies. This workflow should minimize human handling during cell reprogramming, the maintenance of iPSCs, and their differentiation into specific cell types. Furthermore, it is also important that this simplified method can be scaled up to produce adequate cell mass.
To address these issues and reduce variability and workload for technicians, a more streamlined and convenient iPSC workflow is needed. We have developed the protocol presented here to enable iPSC generation under 3D culture conditions, thereby eliminating the need for a colony-picking step [7]. Moreover, our 3D culture system allows for the maintenance and cryopreservation of iPSCs without the need for enzymatic cell dissociation steps, resulting in a significantly simplified and easy-to-follow workflow. Given the compact size of the bioreactor used in this protocol (approximately 148 mm wide × 267 mm deep × 33 mm high with a maximum capacity of six samples at the same time), our protocol holds promise for the full automation of iPSC generation and maintenance in the future.
Materials and reagents
Biological materials
Stealth RNA vector (SRV)TM iPSC vector; SRV iPS-2 vector (encoding OCT4, KLF4, SOX2, and C-MYC; TOKIWA-Bio Inc., Ibaraki, Japan, catalog number: S1011694A) based on the Sendai virus
SRVTM iPSC vector; SRV iPS-4 vector (carrying OCT4, KLF4, SOX2, C-MYC, NANOG, and LIN28; TOKIWA-Bio Inc., catalog number: S1011696A)
Adipose-derived mesenchymal stem cells (Lonza Bioscience, Walkersville, MD, USA, catalog number: PT5006)
Human peripheral blood mononuclear cells (PBMCs) (e.g., derived from healthy donors or patients)
Reagents
MesenPRO RSTM medium (Thermo Fisher Scientific, catalog number: 12746012)
KBM 501 medium (Kohjin Bio, catalog number: 16025015)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, catalog number: 10270-106)
TrypLE Select (Thermo Fisher Scientific, catalog number: 12563-011)
Dulbecco's phosphate-buffered saline (DPBS) (Thermo Fisher Scientific, catalog number: 14190144)
UltraPureTM 0.5 M EDTA, pH 8.0 (Thermo Fisher Scientific, catalog number: 15575020)
StemScaleTM PSC suspension medium (StemScale) (Thermo Fisher Scientific, catalog number: A4965001)
DAPT, gamma-secretase inhibitor (Abcam, catalog number: ab120633)
EPZ004777 (iDOT1L) (Tocris, catalog number: 5567)
Y27632 (FUJIFILM Wako Pure Chemical Corporation, catalog number: 036-24023)
STEM-CELLBANKER® GMP grade (ZENOAQ, catalog number: 11924)
RNeasy Mini kit (Qiagen, catalog number: 74904)
SuperScript IV VILO Master Mix (Thermo Fisher Scientific, catalog number: 11756050)
TaKaRa Ex Taq DNA Polymerase (Takara Bio Inc., catalog number: RR001B)
Primer pair for β-ACTIN and SRV (e.g., Sigma custom oligo; oligo sequence is below)
β-ACTIN (131 bp)
Forward: 5-TCCCTGGAGAAGAGCTACG-3
Reverse: 5-GTAGTTTCGTGGATGCCACA-3
SRV (500 bp)
Forward: 5-ATATGGAGTACGAGAGGACC-3
Reverse: 5-CCTCAGGTTGGAGAGAGTCA-3
Agarose gel (e.g., Nacalai Tesque Inc., catalog number: 01158-85)
Ethidium bromide (e.g., FUJIFILM Wako Pure Chemical Corporation, catalog number: 315-90051)
Solutions
Cell dissociation solution (see Recipes)
PBMC medium (see Recipes)
Reprogramming medium (see Recipes)
Recipes
Cell dissociation solution
Reagent Final concentration Amount
TrypLE Select 0.5× 25 mL
DPBS 0.5× 25 mL
EDTA 0.25 mM 25 µL
Total 50 mL
PBMC medium
Reagent Final concentration Amount
KBM 501 medium 90% 45 mL
FBS 10% 5 mL
Total 50 mL
Reprogramming medium
Reagent Final concentration Amount
StemScale n/a 100 mL
DAPT 5 µM
EPZ004777 3 µM
Y27632 10 µM
Total 100 mL
n/a, not applicable
Laboratory supplies
Microtube 1.5 mL (e.g., Sumitomo Bakelite Co., Ltd., catalog number: MS-4265M)
15 mL centrifuge tube (e.g., IWAKI, catalog number: 2324-015)
30 mL disposable bioreactor (ABLE Biott, catalog number: ABBWVS03A-6)
6-well dish (e.g., IWAKI, catalog number: 3810-006N)
96-well U-bottom dish (e.g., Corning, catalog number: 7007)
PCR tube (e.g., Eppendorf, catalog number: 30124359)
Cryotube (e.g., Thermo Fisher Scientific, catalog number: 377267)
Equipment
CO2 incubator (e.g., Thermo Fisher Scientific, catalog number: 3110, FormaTM series II)
-80 °C freezer (e.g., Panasonic, catalog number: MDF-U33V-PJ)
Centrifuge (e.g., TOMY, catalog number: NIX521)
Bioreactor magnetic stir system base (Able Corp. & Biott Co., catalog number: ABBWBP03N0S-6)
Laser microscope (e.g., Keyence, catalog number: BZ-X810)
Thermal cycler (e.g., Thermo Fisher Scientific, catalog number: 4484073)
Cell counter (e.g., Beckman Coulter, Vi-CELL XR Cell Viability Analyzer System)
Procedure
Transduction of reprogramming factors
Collection of somatic cells
When generating iPSCs from adipose-derived mesenchymal stem cells (AdSCs), collect the cells as a cell pellet after trypsinization with cell dissociation solution for 5 min at 37 °C and centrifugation at 200× g for 3 min.
Count the cells and resuspend them at a density of 1 × 107/mL in 37 °C prewarmed MesenPRO RSTM medium.
Transfer 200 µL of cell suspension (equivalent to 2 × 106 cells) to a new 1.5 mL microtube.
Note: If you attempt to reprogram somatic cells other than AdSCs, resuspend the cells in the appropriate culture medium for that specific cell type. For example, when we reprogram PBMCs, we resuspend the cells in PBMC medium.
Incubation of cells with Sendai virus vector
Thaw SRV iPS-2 vector, tap vigorously, briefly centrifuge at 1,000× g for 1–2 s at 25 °C, and then place on ice.
Add the SRV iPS-2 vector to the cell suspension at a multiplicity of induction (MOI) of 1.
Mix the solution by pipetting five times.
Incubate the cells with the SRV iPS-2 vector in a 1.5 mL microtube at 37 °C and 5% CO2 for 2 h.
Vigorously tap every 20 min.
Notes:
i. Procedures involving viral vectors must be conducted in a Biosafety Level 2 (BSL2) laboratory or higher until SRV-removed iPSC lines are established. Appropriate safety guidelines and regulations for handling and disposal of biohazardous materials must be adhered to.
ii. Two types of SRV are commercially available. One carries four reprogramming factors: OCT4, SOX2, KLF4, and C-Myc. The other carries six reprogramming factors: the aforementioned four factors plus NANOG and LIN28A. The type of SRV to be used depends on the type of cells being reprogrammed. In our experience, fibroblasts can be reprogrammed with four-factor transduction, while PBMCs require six-factor transduction.
iii. The MOI is also a critical factor for successful reprogramming. In our experience, AdSCs can be reprogrammed with an MOI of 1, while PBMCs require an MOI of 3. We recommend starting the reprogramming experiment with an MOI of 1 or 3. If you are unable to reprogram other types of cells, you should optimize the MOI for efficient gene transduction.
Reprogramming somatic cells under 3D conditions
Preparation of bioreactor for reprogramming
Add 20 mL of reprogramming medium to a 30 mL disposable bioreactor.
Pre-incubate at 37 °C, 5% CO2 on a bioreactor magnetic stir system base at 70 rpm placed within the incubator.
Collection of reprogramming factor–transduced cells
After the 2 h incubation of AdSCs with the SRV iPS-2 vector, add 800 µL of MesenPRO RSTM medium to the 1.5 mL microtube containing the cell–vector mixture. This brings the total volume of the cell suspension to 1 mL.
Immediately centrifuge the mixture at 300× g for 3 min and carefully aspirate the supernatant.
Gently resuspend the cell pellet in 1 mL of pre-warmed reprogramming medium.
Transfer the entire cell suspension to the bioreactor pre-placed inside the incubator. Incubate at 37 °C with 5% CO2, rotating at 70 rpm.
Medium change
After 48 h, add 10 mL of reprogramming medium.
Another 48 h, collect 15 mL of cell suspension, centrifuge at 200× g for 3 min at 25 °C, and remove the supernatant.
Resuspend the cell pellet in 15 mL of fresh reprogramming medium.
Add the resuspended cells to the bioreactor.
Change half of the medium every 48 h until iPSC spheres emerge and grow.
Expect the appearance of some cell aggregates approximately 1-month post-seeding.
Continue changing the medium until the cells reach confluence. In this protocol, the iPSC aggregates increase in both size and number without requiring any single-cell dissociation steps.
Expect the cells to reach confluence approximately 2–3 weeks after the primary cell aggregates emerge.
Establishment of iPSC lines under 3D conditions
Preparation of bioreactor for maintenance
Add 30 mL of StemScale (without DAPT, iDOT1L, or Y27632) to a new 30 mL bioreactor.
Pre-incubate at 37 °C, 5% CO2 on a bioreactor magnetic stir system base at 70 rpm.
Selection of SRV vector-negative iPSC spheres (Figure 1)
Once the primary iPSC spheres reach confluence, transfer from 5 to 10 mL of spheres into a 6-well dish.
Under a microscope and using a P1000 pipette, pick up each small sphere (typically ranging in size from 200 to 300 µm) and place it into a well of a 96-well dish. It is important to ensure that no more than one sphere is placed into each well.
Detect the green fluorescent protein (GFP) signal using a laser microscope and mark wells containing GFP-negative spheres, as the SRV iPS-2 vector encodes the EGFP gene in its backbone.
Expect to obtain 5–10 GFP-negative spheres from one 96-well dish. Transfer only GFP-negative spheres to a new 30 mL bioreactor containing StemScale.
Change the medium every 48 h until the iPSC spheres increase in size and number.
Figure 1. Selection and expansion of reprogramming virus vector-free induced pluripotent stem cells (iPSCs) in suspension culture. Cells were observed by fluorescence microscopy for green fluorescent protein (GFP) expression (indicating Sendai virus vector remnant cells). Cells without residual GFP expression were selectively transferred to the next bioreactor. The cells increased in size and number during each passage. GFP-negative spheres indicate the absence of Sendai virus (SeV), as shown by quantitative reverse transcription polymerase chain reaction (RT-PCR) image (right panel). β-ACTIN was the housekeeping gene.
Verification of exogeneous reprogramming vector removal
Extract total RNA from GFP-positive and GFP-negative iPSC spheres and parental somatic cells.
Perform a reverse transcription (RT) reaction using RT enzyme (e.g., Super Script IV Vilo Master Mix).
Perform a polymerase chain reaction (PCR) for a housekeeping gene such as β-ACTIN and the SRV vector. PCR amplification is performed with the appropriate primer pairs and TaKaRa Ex Taq DNA polymerase under the following conditions: initial denaturation at 98 °C for 10 s, followed by 35 cycles of denaturation at 98 °C for 10 s, annealing at 60 °C for 30 s, and extension at 72 °C for 45 s. PCR products are resolved on a 2% agarose gel containing ethidium bromide.
Maintenance of iPSCs
Preparation of bioreactor for maintenance (Figure 2)
Add 30 mL of StemScale (without DAPT, iDOT1L, or Y27632) to a new 30 mL bioreactor.
Pre-incubate at 37 °C, 5% CO2 on a bioreactor magnetic stir system base at 70 rpm.
Figure 2. Reprogramming of somatic cells into induced pluripotent stem cells (iPSCs) in a suspension culture. The reprogrammed cells emerged as small clumps of cells (white arrows) in a spinner flask culture. The spheroids increased in size, cleaved into smaller clumps (white dashed circle), and self-dissociated into smaller pieces over time. The formation of reprogrammed spheroids was observed approximately 30 days after initiating the reprogramming process.
Passage of iPSCs
Transfer a few iPSC spheres (usually 10 spheres are enough) to a new 30 mL bioreactor using a P1000 pipette or 5 mL autopipettor. While our preferred size is approximately 500 µm in diameter, spheres of any size can be transferred.
Incubate at 37 °C and 5% CO2 at 70 rpm, and perform a half-medium change every 48 h until iPSCs reach confluence.
Cryopreservation of 3D-cultured iPSCs
Cell collection
After cell growth, transfer 5 mL of spheroid suspension to a 15 mL centrifuge tube.
Centrifuge the 15 mL tube at 300× g for 3 min or simply let the tube stand for 3 min.
Remove the supernatant.
Cryopreservation
Suspend the iPSC spheres in 5 mL of STEM-CELLBANKER medium, pre-chilled to 4 °C.
Immediately transfer 1 mL of the cell suspension to each cryopreservation tube.
Cryopreserve the cells at -80 °C for short-term storage. For long-term storage, preserve at -276 °C.
Data analysis
As this protocol describes the generation and maintenance of iPSCs under 3D conditions, data analysis was not necessary.
Validation of protocol
This protocol has been validated for the generation of iPSCs from PBMCs. PBMCs were isolated from donors, and 2 × 106 cells were incubated with the SRV iPS vector. For the reprogramming of PBMCs, we employed the SRV iPS-4 vector, which encodes six reprogramming factors. By cultivating the six-factor transduced PBMCs in the reprogramming medium, we observed the emergence of primary iPSC spheres approximately 25 days post-seeding. After cultivating the primary spheres in the same reactor used for reprogramming, the primary iPSC spheres expanded in both size and number without the need for a single-cell dissociation step. Following the selection of GFP-negative spheres and confirmation of transgene removal, we were able to successfully maintain these PBMC-derived iPSCs using the outlined technique. PBMC-derived iPSCs exhibited undifferentiated states and demonstrated the capability to differentiate into all three germ layers, both in vitro and in vivo.
This protocol or parts of it has been used and validated in the following research article(s):
Tsukamoto et al. [7]. A passage-free, simplified, and scalable novel method for iPSC generation in three-dimensional culture. Regenerative Therapy (Figure 4).
General notes and troubleshooting
General notes
A limitation of this protocol lies in the manual selection of GFP-negative spheres under the microscope. This step is labor-intensive and time-consuming compared with other steps in this protocol.
The undifferentiated state of the iPSCs produced and maintained using this protocol is evaluated through standard quantitative PCR and immunostaining procedures. After paraffin-embedding and sectioning, immunostaining can be performed.
The differentiation potential of iPSCs can be assessed in vitro by embryoid body formation and in vivo by teratoma formation.
Troubleshooting
Issue Suggested solution
No iPSC spheres observed
Ensure that the AdSCs do not undergo senescence. Cellular senescence and low growth activity can hinder efficient cell reprogramming.
Verify that the SRV vector was stored at -80 °C or below. We recommend making aliquots to avoid multiple freeze-thaw cycles.
Check that the reprogramming medium is fresh. Use fresh reprogramming medium supplemented with DAPT and iDOT1L. We recommended making aliquots of the StemScale medium and using it within two weeks.
No transgene silencing (no GFP-negative spheres) There are four types of commercially available SRV vectors. The SRV used in this study is either SRV iPS-2 or iPS-4, which are automatically removed from reprogrammed cells by endogenous microRNA-302. If you are using SRV iPS-1 or iPS-3, a small interfering (si)RNA procedure should be performed to remove the SRV from the iPSCs.
Acknowledgments
This work was supported by grants from the Japan Health Research Promotion Bureau Research Fund (2022-B-02) and the Japan Agency for Medical Research and Development (AMED) under grant number 20be0304501h0002 (HA). MT was supported by a research grant from a Grant-in-Aid for JSPS Fellows (22J01591 and 22KJ3169). The protocol was used in Tsukamoto et al. [7] (DOI: 10.1016/j.reth.2024.02.005).
Competing interests
The authors have no conflicts of interest to report.
Ethical considerations
Human PBMCs were collected after obtaining written informed consent. All experiments were approved by the Institutional Review Board of the National Center for Child Health and Development of Japan (permit nos. 385 and 396). All experiments involving human cells were performed in accordance with the tenets of the Declaration of Helsinki (revised 2013).
References
Wiegand, C. and Banerjee, I. (2019). Recent advances in the applications of iPSC technology. Curr Opin Biotechnol. 60: 250–258.
Nagasaka, R., Matsumoto, M., Okada, M., Sasaki, H., Kanie, K., Kii, H., Uozumi, T., Kiyota, Y., Honda, H., Kato, R., et al. (2017). Visualization of morphological categories of colonies for monitoring of effect on induced pluripotent stem cell culture status. Regen Ther. 6: 41–51.
Huang, C. Y., Liu, C. L., Ting, C. Y., Chiu, Y. T., Cheng, Y. C., Nicholson, M. W. and Hsieh, P. C. H. (2019). Human iPSC banking: barriers and opportunities. J Biomed Sci. 26(1): 87.
Lei, Y. and Schaffer, D. V. (2013). A fully defined and scalable 3D culture system for human pluripotent stem cell expansion and differentiation. Proc Natl Acad Sci USA. 110(52): E5039–E5048.
Galvanauskas, V., Grincas, V., Simutis, R., Kagawa, Y. and Kino-oka, M. (2017). Current state and perspectives in modeling and control of human pluripotent stem cell expansion processes in stirred-tank bioreactors. Biotechnol Progr. 33(2): 355–364.
Hookway, T. A., Butts, J. C., Lee, E., Tang, H. and McDevitt, T. C. (2016). Aggregate formation and suspension culture of human pluripotent stem cells and differentiated progeny. Methods. 101: 11–20.
Tsukamoto, M., Kawasaki, T., Vemuri, M. C., Umezawa, A. and Akutsu, H. (2024). A passage-free, simplified, and scalable novel method for iPSC generation in three-dimensional culture. Regen Ther. 27: 39–47.
Article Information
Publication history
Received: Jun 12, 2024
Accepted: Aug 22, 2024
Available online: Sep 10, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Stem Cell > Pluripotent stem cell > Cell-based analysis
Stem Cell > Pluripotent stem cell > Cell induction
Cell Biology > Cell isolation and culture > Cryopreservation
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Genetic Tagging and Imaging of Proteins with iFAST in Candida albicans
JD Jonas Devos
PD Patrick Van Dijck
WG Wouter Van Genechten
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5082 Views: 212
Reviewed by: Shailesh KumarLucy Xie Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Microbiology Mar 2024
Abstract
Candida albicans is the most common human fungal pathogen, able to reside in a broad range of niches within the human body. Even though C. albicans systemic infection is associated with high mortality, the fungus has historically received relatively little attention, resulting in a lack of optimized molecular and fluorescent tools. Over the last decade, some extra focus has been put on the optimization of fluorescent proteins (FPs) of C. albicans. However, as the FPs are GFP-type, they require an aerobic environment and a relatively long period to fully mature. Recently, we have shown the application of a novel type of fluorogen-based FP, with an improved version of fluorescence activating and absorption shifting tag (iFAST), in C. albicans. Due to the dynamic relation between iFAST and its fluorogens, the system has the advantage of being reversible in terms of fluorescence. Furthermore, the combination of iFAST with different fluorogens results in different spectral and cellular properties, allowing customization of the system.
Key features
• Genetic integration and tagging with the iFAST tag in Candida albicans.
• Imaging and localization of a protein of interest tagged with iFAST.
• Reversibility of fluorescence with iFAST.
Keywords: Candida albicans iFAST Fluorescence activating and absorption shifting tag Reversible fluorescence Protein tag Multi-color Fluorescence microscopy
Graphical overview
Background
The ubiquitous commensal Candida albicans is a fungus that resides in the gastrointestinal tract, blood, skin, and other mucosal surfaces [1–3]. In healthy individuals, the immune system is sufficiently capable of eradicating C. albicans, but upon opportunistic circumstances, the fungus can become pathogenic, often with a high morbidity rate [4]. As a species, C. albicans is closely related to the eukaryotic model organism Saccharomyces cerevisiae. However, despite this close relationship, C. albicans has an aberrant codon usage, translating the CUG codon to serine instead of leucine in 96% of cases [5,6]. Consequently, heterologous expression of C. albicans proteins requires codon optimization. Conversely, this also means that molecular tools and protein labeling techniques developed for use in S. cerevisiae cannot easily be translated to C. albicans, as these will suffer from the same mistranslation issues. Regarding this issue, several techniques for codon optimization in C. albicans have been reported [7,8], but studies have shown that these are not generally applicable [9].
In recent years, some fluorescent tools have been developed/optimized for use in C. albicans (and extensively reviewed in Van Genechten et al. [10]). In this protocol, we show the use of a novel type of fluorescent protein, an improved version of fluorescence activating and absorption shifting tag (iFAST), which requires the binding of a fluorogen, instead of chromophore maturation, to be fluorescently active. This novel system was developed by the group of Gautier for use in mammalian cells [11,12]. The iFAST protein is half the size of GFP and has the added benefit of reversibly binding its fluorogens, which are analogs of 4-hydroxybenzyliden-rhodanine (HBR). On their own, iFAST and HBR analogs are relatively non-fluorescent, but upon contact of iFAST protein with its fluorogen, fluorescence can be observed. An added benefit of this system is that the interaction is reversible and does not require oxygen, making this a reversible fluorescent system even in anaerobic conditions, where classical FPs like GFP do not work.
Materials and reagents
Biological materials
SC5314 Candida albicans wild-type strain [13]
SN152 Candida albicans arg4Δ/arg4Δ leu2Δ/leu2Δ his1Δ/his1Δ URA3/ura3Δ::imm434 IRO1/iro1Δ::imm434 [14]
Reagents
Granulated yeast extract (Merk, catalog number: 1.03753)
Bacterial peptone (Oxoid, catalog number: LP0037B)
Glucose (Fluka, catalog number: 49459)
Glycerol (Sigma, CAS number: 58-81-5)
Granulated bacteriological agar (Difco, catalog number: 214530)
Nourseothricin (Jena Bioscience, CAS number: 96736-11-7)
Complete synthetic mixture (CSM) (MP Biomedicals, catalog number: 114500022)
Yeast nitrogen base (YNB) without amino acids, ammonium sulfate, and riboflavin (Formedium, catalog number: YN6501)
Ammonium sulfate (VWR, CAS number: 7783-20-2)
Lithium acetate dihydrate (LiAc·2H2O) (Sigma-Aldrich, CAS number: 6108-17-4)
Polyethylene glycol (PEG) 3350 (Sigma-Aldrich, CAS number: 25322-68-3)
Trizma® base (Sigma-Aldrich, CAS number: 77-86-1)
HCl (Sigma-Aldrich, CAS number: 7647-01-0)
EDTA (Sigma-Aldrich, CAS number: 6381-92-6)
HMBR (Twinkle Factory, catalog number: 480541-250)
HBR-3,5DM (Twinkle Factory, catalog number: 499558-250)
HBR-3,5DOM (Twinkle Factory, catalog number: 516600-250)
HIFI Q5 polymerase (New England Biolabs, catalog number: NEB M0491L)
NEBuilder® HiFi DNA Assembly Master Mix (New England Biolabs, catalog number: NEB E2621X)
ssDNA, MB-grade from fish sperm (Merck, catalog number: 11467140001)
rCutsmart restriction buffer (New England Biolabs, catalog number: B6004S)
Restriction enzymes:
StuI (New England Biolabs, catalog number: R0187S)
NheI-HF (New England Biolabs, catalog number: R3131S)
PstI-HF (New England Biolabs, catalog number: R3140S)
Solutions
YPD (see Recipes)
YPD-NAT agar (see Recipes)
YPD-glycerol (see Recipes)
Low fluorescence medium (see Recipes)
1 M LiAc (see Recipes)
PEG 3350 (50%) (see Recipes)
TE (10×) (see Recipes)
LiAc/TE buffer (see Recipes)
LiAc/PEG buffer (see Recipes)
5 mM fluorogen stock solution (see Recipes)
Recipes
YPD (1 L)
Reagent Final concentration Quantity or Volume
Yeast extract granulated 1% (w/v) 10 g
Bacteriological peptone 2% (w/v) 20 g
Glucose 2% (w/v) 20 g
H2O (demineralized) n/a 1 L
YPD-NAT agar (1 L)
*Note: Add the nourseothricin after heat sterilization and when the medium has cooled down to approximately 50 °C.
Reagent Final concentration Quantity or Volume
Yeast extract granulated 1% (w/v) 10 g
Bacteriological peptone 2% (w/v) 20 g
Glucose 2% (w/v) 20 g
Bacteriological agar 2% (w/v) 20 g
Nourseothricin* 0.02% (w/v) 200 mg
H2O (demineralized) n/a 1 L
YPD-glycerol (100 mL)
Reagent Final concentration Quantity or Volume
Yeast extract granulated 1% (w/v) 1 g
Bacteriological peptone 2% (w/v) 2 g
Glycerol 30% (v/v) 30 mL
H2O (demineralized) n/a 70 mL
Low fluorescence medium (1 L)
Reagent Final concentration Quantity or Volume
CSM 0.079% (w/v) 0.79 g
YNB (without amino acids, without ammonium sulfate, without riboflavin) 0.69% (w/v) 6.9 g
Ammonium sulfate 0.5% (v/v) 5 g
Glucose 2% (w/v) 20 g
H2O (demineralized) n/a 1 L
1 M LiAc (100 mL)
Reagent Final concentration Quantity or Volume
LiAc·2H2O 1 M 10.2 g
H2O (demineralized) n/a 100 mL
PEG 3550 (50%) (100 mL)
*Note: First, add 50 mL of H2O and a stir bar. Let the PEG dissolve overnight and add water up to a total volume of 100 mL.
Reagent Final concentration Quantity or Volume
PEG 3550 50% (w/v) 50 g
H2O (demineralized) n/a Until 100 mL*
TE (10×) (1 L)
*Note: After adding Tris, adjust the pH to 8 using HCl.
Reagent Final concentration Quantity or Volume
Tris 10 mM 12.11 g*
EDTA 1 mM 2.92 g
H2O (demineralized) n/a 1 L
LiAc/TE buffer (1 mL)
Reagent Final concentration Quantity or Volume
LiAc (1M) 0.1 M 100 µL
TE (10×) 1× 100 µL
H2O (MiliQ) n/a 800 µL
LiAc/PEG buffer (1 mL)
Reagent Final concentration Quantity or Volume
LiAc (1 M) 0.1 M 100 µL
TE (10×) 1× 100 µL
PEG 3350 [50% (w/v)] 40% (w/v) 800 µL
Fluorogen stock solution
*Note: The quantity is calculated to make a total of 100 mL. Store this stock solution in aliquots of 20 µL at -20 °C. Make a separate stock solution for each fluorogen; do not mix. Fluorogen stock solutions can withstand multiple freeze-thaw cycles; however, try not to defrost more than five times. Additionally, it is best to protect the fluorogens from light as much as possible when working.
Reagent Final concentration Quantity or Volume
HMBR 5 mM 0.125 g*
HBR-3,5DM 5 mM 0.133 g*
HBR-3,5DOM 5 mM 0.149 g*
Laboratory supplies
Microtubes (1.5 mL) (Eppendorf, catalog number: 0030125150)
Micro pipettes (sizes 1,000, 200, and 10 µL)
Micro pipette tips (standard tips of 1,000, 200, and 10 µL)
Microscope slides (VWR, catalog number: 630-1985)
Glass covers (VWR, catalog number: 631-0122)
Glass culture tubes (10 mL) (Fisher Scientific, catalog number: 11557403)
Caps for culture tubes (VWR, catalog number: LUDI184010631)
Erlenmeyer flasks (300 mL) (VWR, catalog number: 391-0275)
Erlenmeyer flask caps (VWR, catalog number: 391-0950)
Clear cuvette (VWR, catalog number: 97000-590)
Vortex (VWR, catalog number: 444-1372)
NucleoSpinTM Gel and PCR Clean-up Kit (Macherey-Nagel, catalog number: 740611.50)
NucleoSpinTM Plasmid EasyPure Kit (Macherey-Nagel, catalog number: 740727.50)
Equipment
FV1000 microscope (Olympus, model: FV1000-IX81) with 60× UPlanSApo (NA1.35) objective lens (Olympus) or equivalent
ThermoMixer® F1.5 (Eppendorf, catalog number: 5384000020) or equivalent
Shaking incubator (New Brunswick Scientific, model: Innova40) or equivalent
Microtube centrifuge (Eppendorf, model: 5471C) or equivalent
BioPhotometer (Eppendorf, model: 6131)
NanoDropTM 1000 spectrophotometer (Thermo Scientific, model: ND-1000) or equivalent
Software and datasets
FV10-ASW 4.2 software package (Olympus)
ImageJ (v1.53t, 24/08/2022) (https://imagej.net/software/fiji/downloads)
RStudio (R version: 4.3.2, 31/10/2023) (https://cran.rstudio.com/; https://posit.co/download/rstudio-desktop/)
Procedure
Below we provide a stepwise overview of tagging a protein of interest with iFAST in C. albicans. We describe how to do this in order to obtain an endogenous tag or an overexpression construct. This protocol has been used to tag Ftr1 and Erg11, endogenously or as an overexpression construct, respectively [15]. Furthermore, we also describe a protocol to utilize the reversibility of the iFAST system to alter the absorption and excitation maxima of the tagged construct by exchanging the fluorogen. All steps of this protocol were performed at room temperature (22 °C) unless stated otherwise.
Transformation of Candida albicans (see General note 1)
Streak the wild-type C. albicans (SC5314) from the -80 °C stock and plate on YPD agar at 30 °C overnight (ON).
Transfer a single colony to 3 mL of YPD and grow ON at 30 °C in a shaking incubator (at 240 rpm).
Measure the cell density of the culture at 600 nm using the BioPhotometer. Dilute the ON culture to an OD600 of 0.2–0.4 in 50 mL of YPD in a 300 mL Erlenmeyer flask. Incubate again at 30 °C and grow the cells to a density of approximately 1.5; this will take 3–4 h.
Pellet the cells at 1,000× g for 5 min.
Remove the supernatant and resuspend the cells in 0.7 mL of LiAc/TE buffer.
While adding the LiAc/TE buffer, mix the cells well by pipetting up and down.
Pipette in a new Eppendorf tube (in the following order):
100 µL of the cell suspension in LiAc/TE buffer.
DNA (1 µg) that needs to be transformed.
i. In the case of a plasmid: First, linearize the plasmid via a restriction digestion. Then, use the unpurified restriction product.
ii. In case of PCR: Use 50 µL of unpurified linear PCR product.
20 µL of fish sperm ssDNA (10 mg/mL) (preheated to 98 °C for 5 min and cooled to 50 °C).
Leave at room temperature for 10 min.
0.7 mL of LiAc/PEG buffer.
Vortex shortly.
Incubate for 16 h at 30 °C while shaking (300 rpm).
Heat-shock the cell mixture using a heat block for 15 min at 44 °C.
Pellet cells at 5,200× g for 1 min.
Remove the supernatant and resuspend in 1 mL of YPD.
Let the cells recover for 4 h at 30 °C while shaking (300 rpm).
Pellet the cells at 5,200× g for 1 min.
Plate out on YPD agar containing 200 mg/L nourseothricin for both the pFA6-based endogenous tagging and the overexpression strains using the CIp10 plasmid. Incubate for 24 h at 30 °C and re-streak to single colonies on YPD-NAT agar. The plates should contain approximately 20 colonies for the pFA6 transformation and more than 100 colonies for the CIp10 transformation.
Colonies are verified based on the presence and localization of fluorescence under the confocal microscope. Alternatively, the transformation can be checked through colony PCR [16], targeting the tagged gene and iFAST (for iFAST reverse primer, see Table S3). Since the integration efficiency of the linearized CIp10 plasmid was around 90% in our case, testing five colonies proved to be sufficient. The integration efficiency of the PCR product for endogenous tagging of genes was less efficient (approximately 10%–20%), so at least 12 colonies were tested in that case.
Endogenous protein tagging with iFAST
The iFAST-SAT1 construct will be integrated between the end of the gene and the beginning of the terminator. For this, a forward and reverse primer need to be constructed with a sequence that overlaps with specific regions within the genome and binds on the pFA6-iFAST plasmid (available on AddGene: #209414). The genomic sequences of C. albicans genes can be accessed at http://www.candidagenome.org (see General note 2).
Design the forward primer to contain the last 50 bp of the gene of interest (GOI) excluding the stop codon, a double GGGGS linker (“GGAGGTGGAGGTTCTGGTGGAGGTGGTTCA”), and the first 20 bp of iFAST (“ATGGAACATGTTGCCTTTGG”).
The reverse primer consists of the first 50 complementary bp of the GOI terminator and the last 20 bp of the SAT1 gene (“TTAGGCGTCATCCTGTGCTC”).
Using the constructed primers, amplify the iFAST-SAT1 construct from the pFA6-iFAST plasmid using Q5 HiFi DNA polymerase (New England Biolabs) using the protocol in Table 1. The master mix was made following the manufacturer’s instructions.
Table 1. Thermocycling conditions for Q5 PCR reaction
Step Temperature (°C) Duration Number of cycles
Initial denaturation 98 30 s 1
Denaturation 98 15 s
30
Back to step 2
Annealing 64 25 s
Extension 72 30 s per kb
Final extension 72 5 min 1
Hold 10 ∞ -
Follow the protocol described in section A to transform the iFAST-SAT1 construct in C. albicans wild type. Select on medium containing 200 mg/L nourseothricin (see General note 3).
Pick a colony and re-streak again to obtain single colonies on YPD-NAT agar before continuing.
Colonies are verified based on fluorescence and its localization under the confocal microscope or by PCR, targeting the GOI and the iFAST gene (for iFAST reverse primer, see Table S3).
To check protein localization, grow a single colony overnight in low fluorescence medium at 30 °C. Dilute the culture in fresh medium to OD600 0.2–0.4 and grow for 4–5 h until the mid-exponential phase (~OD600 1). Incubate the strain for 15 min in low fluorescence medium with 50 µM of fluorogen. Afterward, add 1–2 µL of culture to a standard microscopy slide, cover with a coverslip, and check under the confocal microscope. Choose the settings of the microscope in accordance with the fluorogen (Table 2).
Transfer verified colonies to YPD-glycerol medium and place at -80 °C for long-term storage.
Overexpression tagging with iFAST
In silico Gibson cloning
The construction of the plasmid for overexpression tagging is based on Gibson assembly [17]. In the NEBuilder tool (https://nebuilder.neb.com/#!/) or a similar in silico cloning tool, insert the sequence of the plasmid backbone (CIp10-NAT1; AddGene: #225583) (Text S1). Process the plasmid for restriction digestion with PstI and NheI to allow the integration of two inserts.
As the first insert, add the full DNA sequence of the GOI, excluding the stop codon.
As the second fragment to insert, add the full iFAST sequence (Text S2). In between these fragments, add a double [GGGGS]2 linker.
In vitro cloning
Using the primers constructed by the in silico cloning tool, amplify the GOI and iFAST using Q5 HiFi DNA polymerase using the protocol in Table 1 (see General note 4).
Amplify the sequence of the GOI from isolated gDNA.
iFAST can be amplified from an iFAST-containing plasmid (pFA6a-iFAST available on AddGene: #209414) or gBlock.
Afterward, purify the PCR fragments using the NucleoSpinTM Gel and PCR Clean-up kit and measure the concentration using Nanodrop.
After the restriction of the backbone plasmid with PstI and NheI, purify the restriction mixture using the NucleoSpinTM Gel and PCR Clean-up kit. Measure the concentration of the purified, restricted plasmid using Nanodrop.
Construct the overexpression plasmid by ligating the linearized vector with the two PCR products (GOI and iFAST). The ligation mixture to do this consists of 50 ng of plasmid and a molar ratio of plasmid:fragments of 1:3 with 10 µL of NEBuilder® HiFi DNA Assembly Master Mix and the total volume adjusted to 20 µL. This mixture was incubated at 50 °C for up to 1 h (see General note 5).
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Amplify the constructed plasmid using chemically competent E. coli [18]. Afterward, purify the plasmids using the NucleoSpinTM Plasmid EasyPure kit.
Before transformation into C. albicans, linearize the plasmid using restriction digestion by StuI. Transformation into C. albicans is done according to an adapted Gietz protocol (section A).
After transformation, select on YPD-NAT. Re-streak to single colonies and check successful tagging using fluorescence microscopy. Alternatively, colonies can be checked by PCR targeting the GOI and the iFAST gene. The FW primer within the GOI is unique, whilst the RV primer for the iFAST gene is “TTGCCAATCACTTGTTTGGG”.
To check protein localization, grow a single colony overnight in low fluorescence medium at 30 °C. Dilute the culture in fresh medium to OD600 0.2–0.4 and grow for 4–5 h until the mid-exponential phase (~OD600 1). Incubate the strain for 15 min in low fluorescence medium with 50 µM of fluorogen. Afterward, add 1–2 µL of culture to a standard microscopy slide, cover with a coverslip, and check under the confocal microscope. Choose the settings of the microscope in accordance with the fluorogen (Table 2).
Transfer verified colonies to YPD-glycerol medium and place at -80 °C for long-term storage.
Reversible fluorescence of iFAST
Grow the transformed cells overnight at 30°C in low fluorescence medium (Figure 1).
Dilute the transformed strain in fresh low fluorescence medium to OD600 of 0.2 and grow for 4–5 h until the mid-exponential phase (~OD600 1).
Before starting the experiment, predetermine the time points at which you want to image the cells (see General note 6).
Prior to adding any fluorogen, add 1–2 µL of culture on a standard microscopy slide, cover with a coverslip, and image using the fluorescent microscope (see section E). Use this during the analysis as the pre-treatment condition.
Add the fluorogen to the culture for a final concentration of 50 µM of fluorogen and a final volume of 1 mL.
For each predetermined time point repeat the following steps:
Add 1–2 µL of the culture on a standard microscopy slide and cover with a coverslip.
Let the rest of the culture incubate at 30 °C with gentle shaking (300 rpm) until the next time point.
Image the sample under the confocal microscope (see section E).
After the last time point, wash the cells twice by spinning down (1 min at 5,200× g), replacing the supernatant with medium lacking fluorogen. Image 1–2 µL of the culture on a standard microscopy slide after each of the washing steps (see General note 7).
If desired, the same culture can be used to repeat this experiment with another fluorogen to exploit the multicolor characteristic of the iFAST protein. For this, use the washed culture and repeat from step D4 (Figure 1).
Figure 1. Diagram of the experimental steps of section D. All incubation steps are performed at 30 °C.
Sample preparation and confocal microscopy
For handling the microscope, we used a previously published protocol [19]. For the selection of the laser lines, excitation filter, and appropriate emission filters for the different fluorogens, see Table 2.
Aside from the fluorescent channel, concurrent DIC or brightfield images are taken (see General note 8).
From the FV-10 software (or other user-specific software), images of both channels are exported to a (multi-)TIFF format (see General note 9).
Table 2. Overview of the validated fluorogens and their appropriate microscopy settings
Fluorogen λexcmax/λemmmas Excitation laser Excitation filter Emission filter
HMBR 480/541 nm 488 nm DM405/488 BA505–605 nm
HBR-3,5DM 499/558 nm 488 nm DM405/488 BA505–605 nm
HBR-3,5DOM 516/600 nm 515 nm DM458/515 BA575–620 nm
Data analysis
To evaluate the reversibility of fluorescence, the fluorescence intensity of the cells needs to be calculated and can be compared over time. All image analysis was done using the ImageJ2 software [20]. To do this, install and open the ImageJ2 software. Open a multi-TIFF image: File > Open…
Make a new macro to facilitate the image processing: Plugins > New>Macro. Past in the code written below.
run("Set Measurements...", "area mean display redirect=None decimal=3");
// This if statement clears out the ROI manager if it is not empty
if(roiManager("count")!= 0){
roiManager("Delete");
}
// The Min and Max numbers are chosen in accordance with our system, adjust these //if needed
setMinAndMax(0, 4095);
run("Stack to Images");
This macro makes sure that the correct measuring methods are set. Further, it will check if the ROI manager is empty; if not, it will clear the ROI manager. Additionally, it sets the minimal and maximal brightness of the image. Lastly, it separates the two channels of a multi-TIFF image (see General note 10). To run the macro, select the Run button.
Using the Freehand selection tool, select the regions of interest (ROIs). Select all C. albicans cells in the DIC or brightfield image and copy the ROIs to the ROI manager. To achieve this, select a cell and press the T button on the keyboard, and the ROI manager will open automatically.
In addition to the cells, select seven regions on the DIC image that only include background and no cells. These seven regions should be of similar size as the cells. These will be used to calculate the background fluorescence of the image.
Once all cells and background regions on the DIC or brightfield image have been selected and copied to the ROI manager, select the image of the fluorescent channel and measure via the ROI manager: Window > ROI manager > Measure. This will open a dialog box titled “Results.” Copy these results to an Excel file.
For the analysis of the fluorescence intensity at each timepoint, including the pre-treatment control, follow the following steps:
For each image, calculate the average intensity of the seven background ROI’s and subtract this value from the cell values.
To calculate the fold change in fluorescence, the data is normalized to the pre-treatment control (see General note 11). Calculate the average of the background-corrected cell measurement of the pre-treatment condition.
Divide every background-corrected cell measurement with this normalization value, including the cell measurements of the pre-treatment condition (see General note 12).
The fold change data of all cells is put into one big dataset, including information on which fluorogen was used and at which time point the data was collected (an example of such dataset can be found at https://github.com/Jonas-Ds/iFAST-RStudio-Figures).
For the visualization and statistical analysis, RStudio with the ggplot and stats packages were used, respectively. RStudio script with example data can be found here: https://github.com/Jonas-Ds/iFAST-RStudio-Figures/. At each time point, the two different strains (iFAST tagged strain vs. wild-type control) were compared using a Student's t-test.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Devos et al. [15]. A multi-colour fluorogenic tag and its application in Candida albicans. Microbiology (Figures 2–4; Figure S3).
General notes and troubleshooting
General notes
Transformation of C. albicans is based on a LiAc/PEG method, which promotes the uptake of DNA by the yeast cells.
Following are two example primers for the endogenous tagging of FTR1 in Candida albicans. The full genetic sequence including 1 kb up/downstream can be found on the Candida Genome Database (CGD) (www.candidagenome.org/). For the forward primer, the last 50 bp of the gene, excluding the stop codon, are in italics; the linker is underlined; and the first 20 bp of iFAST are in bold.
AAGTTGATGAAACTTCATCAAACAAATTGATCGAATCCAAAGAAAACAAAGGAGGTGGAGGTTCTGGTGGAGGTGGTTCAATGGAACATGTTGCCTTTGG
For the reverse primer, the reverse complement of the first 50 bp of the terminator of FTR1 is in italics, and the reverse complement of last 20 bp of the SAT1 gene is in bold.
CACAGTCTCTTGCCTTATTCTTTTAGTTGTTGAATAATAATTAACTAAGTTTAGGCGTCATCCTGTGCTC (Table S3).
When using these primers for a different gene, only the italics part of the primers needs to be adjusted to the gene of interest.
No purification of the PCR product is performed, as this will lower the yield of transformation. C. albicans is not efficient in keeping plasmids even under selective pressure, therefore no purification is required.
The melting temperature for the PCRs is calculated using the Tm calculator tool of New England BioLabs (https://tmcalculator.neb.com/#!/main). In case a different DNA polymerase is used, the thermocycling conditions of Table 1 should be adjusted according to the DNA polymerase.
The NEBuilder® HiFi DNA Assembly Master Mix contains an exonuclease, a polymerase, and a DNA ligase. Due to the combination of these enzymes, no further processing of the PCR fragments is required after the restriction of the plasmid.
In the original publication of this protocol, a couple of different imaging and timing strategies can be found [15]. These include imaging at 2, 5, 10, and 15 min followed by two washing steps. This imaging and washing combination can then be followed by the addition of a different fluorogen and subsequent fluorescence imaging. The exact time points of imaging depend on the type of experiment and biological question. It is recommended that the imaging of the samples is done between timepoints and not to wait until the end of the experiment. For each time point, is it sufficient to take 1–2 µL of culture, so the same culture can be used for the entire experiment. Wash steps are performed on the whole culture.
It is recommended to image the culture after each wash step so that the decline in fluorescence due to the removal of the fluorogen can be followed and later represented during data analysis.
When the cells are in focus, it is recommended to take three pictures, each time of a different group of cells, to get more information about the cells at each time point. We recommend a minimum of 25 cells per strain per time point.
Here, we export as a multi-TIFF file, as this is our personal preference. The data analysis will also work fine if you export each channel as a separate TIFF file. If your software does not allow exporting to a TIFF file, you can use any other picture format that is compatible with image analysis in ImageJ2.
In case the channels of the fluorescent microscope were exported as separate TIFF files, remove the last line of the macro script.
In case of using multiple fluorogens in succession, the last image before the addition of the new fluorogen, of the second wash step, is used as the pre-treatment control for the new fluorogen.
The pre-treatment cell measurements are taken along to take into account the variation even before the addition of the fluorogen.
Troubleshooting
Problem 1: Failed C. albicans transformation
Possible cause: Expired nourseothricin, insufficient transformed DNA, PEG concentration changes, no fresh TE mixture.
Solution:
In the case of endogenous tagging, the selection of correct transformants is performed through the antibiotic nourseothricin. If you have too many incorrect transformants when transforming and selecting with nourseothricin, it may be that the nourseothricin is expired, added to the medium when it wasn’t cooled down sufficiently, or that your specific background strain is relatively resistant to nourseothricin. In these cases, we would advise using new nourseothricin, cooling the agar medium sufficiently, or increasing the nourseothricin concentration.
On the other hand, if no colonies are acquired after transformation, several adjustments could be made. First, prepare novel PEG and TE mixtures, since long-term evaporation might lead to concentration adjustments, as mentioned by Gietz and Schiestl [21]. Another possible explanation is insufficient PCR product or linearized plasmid. Check concentrations utilizing a Nanodrop spectrophotometer or equivalent instrument. If the concentration proved to be sufficient, you could still utilize more DNA, e.g., up to 5 µg. Using agar gel electrophoresis, it is also possible to verify whether the restriction digestion of the plasmid has worked.
In the case of endogenous construction, overhangs of 50 bp for homolog recombination might not be sufficient. It is possible to increase the overhang length; we have utilized overhangs of up to 100 bp.
Problem 2: Confocal microscopy
Possible cause: Insufficient fluorescence signal due to insufficient excitation or fluorogen.
For confocal microscopy, it is important to keep in mind that each genetically tagged strain will behave slightly differently, and this will require some optimization for each experiment. When experiencing low fluorescence from the tagged strains, we recommend adjusting the laser power and the voltage on the photomultiplier tube of the microscope, as this will influence the brightness and contrast of the images. However, keep in mind that higher laser power will also cause more photobleaching; in this protocol, we use an argon laser at 100 µW and the high voltage of the photomultiplier tube set to 580. Low fluorescence might also be caused by the fluorogen stock solution that has been through too many freeze-thawing cycles or has been exposed to light too much. For this, try to repeat the experiment with a new aliquot for the fluorogen stock. Increasing the fluorogen concentration might also solve the issue of low fluorescence; however, keep in mind that increasing the fluorogen concentration will also increase background fluorescence and can have a slight effect on the growth of the strains.
Additionally, we have not encountered that certain fluorogens are unable to reach specific organelles or are quenched at those organelle conditions. It is therefore a valid strategy to test other HMBR variants that are utilized in this protocol.
Acknowledgments
These protocols were used to acquire the data and results that were reported in our publication in Microbiology (Devos et al. [15], https://doi.org/10.1099/mic.0.001451). The protocol was developed using internal funding of the KU Leuven Research Council (grant # C14/22/075).
Competing interests
The authors declare that there are no competing interests.
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Bougnoux, M. E., Diogo, D., François, N., Sendid, B., Veirmeire, S., Colombel, J. F., Bouchier, C., Van Kruiningen, H., d'Enfert, C., Poulain, D., et al. (2006). Multilocus Sequence Typing Reveals Intrafamilial Transmission and Microevolutions of Candida albicans Isolates from the Human Digestive Tract. J Clin Microbiol. 44(5): 1810–1820.
Findley, K., Oh, J., Yang, J., Conlan, S., Deming, C., Meyer, J. A., Schoenfeld, D., Nomicos, E., Park, M., et al. (2013). Topographic diversity of fungal and bacterial communities in human skin. Nature. 498(7454): 367–370.
Brown, G. D., Denning, D. W., Gow, N. A. R., Levitz, S. M., Netea, M. G. and White, T. C. (2012). Hidden Killers: Human Fungal Infections. Sci Transl Med. 4(165): e3004404.
Gomes, A. C., Miranda, I., Silva, R. M., Moura, G. R., Thomas, B., Akoulitchev, A. and Santos, M. A. (2007). A genetic code alteration generates a proteome of high diversity in the human pathogen Candida albicans. Genome Biol. 8(10): R206.
Santos, M. A. and Tuite, M. F. (1995). The CUG codon is decoded in vivo as serine and not leucine in Candida albicans. Nucleic Acids Res. 23(9): 1481–1486.
Daniel, E., Onwukwe, G. U., Wierenga, R. K., Quaggin, S. E., Vainio, S. J. and Krause, M. (2015). ATGme: Open-source web application for rare codon identification and custom DNA sequence optimization. BMC Bioinf. 16(1): 303.
Puigbo, P., Guzman, E., Romeu, A. and Garcia-Vallve, S. (2007). OPTIMIZER: a web server for optimizing the codon usage of DNA sequences. Nucleic Acids Res. 35: W126–W131.
Van Genechten, W., Demuyser, L., Dedecker, P. and Van Dijck, P. (2020). Presenting a codon-optimized palette of fluorescent proteins for use in Candida albicans. Sci Rep. 10(1): 6158.
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Supplementary information
The following supporting information can be downloaded here:
Text S1. CIp10-NAT1plasmid map
Text S2. iFAST sequence
Table S3. Primers
Article Information
Publication history
Received: May 30, 2024
Accepted: Aug 23, 2024
Available online: Sep 10, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > Protein > Detection
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5,083 | https://bio-protocol.org/en/bpdetail?id=5083&type=0 | # Bio-Protocol Content
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Sorghum bicolor Extracellular Vesicle Isolation, Labeling, and Correlative Light and Electron Microscopy
DA Deji Adekanye *
TC Timothy Chaya *
JC Jeffrey L. Caplan
(*contributed equally to this work)
Published: Vol 14, Iss 19, Oct 5, 2024
DOI: 10.21769/BioProtoc.5083 Views: 276
Reviewed by: Xiaofei LiangAnuradha SinghMasayoshi Nakamura
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Original Research Article:
The authors used this protocol in Plant Physiology Feb 2024
Abstract
Extracellular vesicles are membrane-bound organelles that play crucial roles in intercellular communication and elicit responses in the recipient cell, such as defense responses against pathogens. In this study, we have optimized a protocol for isolating extracellular vesicles (EVs) from Sorghum bicolor apoplastic wash. We characterized the EVs using fluorescence microscopy and correlative light and electron microscopy.
Key features
• Allows the isolation of extracellular vesicles from the monocot plant Sorghum bicolor.
• Labels isolated extracellular vesicles with fluorescent dyes for easy characterization with light microscopy.
• Validates dye labeling and further characterizes extracellular vesicles using a correlative light and electron microscopy approach.
Keywords: Sorghum bicolor Monocot Extracellular vesicles Correlative light and electron microscopy
Background
Extracellular vesicles (EVs) are membrane-bound organelles secreted by cells [1,2]. Many studies have shown that EVs are packaged with RNA, lipids, proteins, and other metabolites; these EV contents play an important role in mediating intercellular signaling [1–3].
The observation of plant and fungal EVs deposited at fungal infection sites suggested a possible mediation of the plant–pathogen interaction by EVs [4–6]. The notion of EVs mediating plant–pathogen interactions is supported by multiple studies showing enhanced plant EVs secretion after fungal infection, silencing of host plant’s immunity genes by sRNA packaged in fungal EVs, and modulation of fungal gene expression by host plant mRNAs [6–12]. Proteomic analysis of isolated plant EVs revealed they contain plant defense proteins [13–15], and a non-canonical secretory pathway has been proposed as the route for secretion of defense factors and virulence factors by the plants and fungal pathogens, respectively [16,17]. However, despite plant EVs having been described since the 1960s [5], methods for their isolation were only developed fairly recently for eudicot plants. Eudicot plant EVs have been largely isolated from the apoplastic wash of leaves through differential centrifugation, immuno-affinity capture, and density gradient centrifugation [3,15,18].
Here, we provide a detailed protocol that we used previously [19] for the isolation and characterization of EVs from the monocot plant Sorghum bicolor. To detect EVs by fluorescence microscopy, we designed a method for staining the EVs using an esterified dye, Calcein AM Green, and a lipophilic dye, Potomac Gold. To further characterize EVs, we developed a correlative light and electron microscopy method to examine the morphology of EVs stained with our fluorescent labeling approach. The new methods developed here complement standard EV characterization methods, such as nanoparticle tracking analysis and transmission electron microscopy.
Materials and reagents
Pro-mix BK55 soil (Premier Tech Horticulture, catalog number: 5060055RG)
6” standard pot (Grower’s, catalog number: 03EJ-SP600)
Sorghum bicolor seeds (BTx623 inbred line)
Scissors (W.B. Mason, catalog number: ACM13529)
Parafilm®, 4 in. × 250 ft (Bemis, catalog number: PM999)
Pyrex dish (Corning, catalog number: 3160100)
60 mL syringe (BD medical, catalog number: BD301036)
8-inch half-round file (Harbor Freight, catalog number: 96629)
Open-top thick-wall polypropylene tubes, 50 mL, 29 mm × 104 mm (Beckman Coulter, catalog number: 357007)
250 mL Nalgene bottles, (Millipore Sigma, catalog number: B9157)
2-(morpholin-4-yl)ethane-1-sulfonic acid (MES) hydrate (Sigma-Aldrich, catalog number: M8250)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: SS7653)
Calcium chloride (CaCl2) (Sigma-Aldrich, catalog number: C1016)
1 L Glass beaker (Pyrex, catalog number: 100-1L-PK)
Miracloth 475855-1R (Millipore, catalog number: 3446249)
1 M Tris-Hydrochloride (Tris-HCl), pH 7.5 (Fisher Bioreagents, catalog number: BP1757-500)
KimwipesTM (Kimberly-Clark, catalog number: 34120)
25 ml serological pipettes (Thermo Fisher Scientific, catalog number: 170357N)
OptiPrep density gradient (60% iodixanol) (Sigma-Aldrich, catalog number: D1556)
Thin-walled ultra-clear centrifuge tubes (14 mm × 89 mm) (Beckman Coulter, catalog number: 344059)
Calcein Green, AM (Thermo Fisher Scientific, catalog number: C34852)
Potomac Gold (Lavis Lab, HHMI Janelia)
Poly-L-lysine (Sigma-Aldrich, catalog number: P4707)
Trypan blue (Sigma-Aldrich, catalog number: T6146)
HPLC grade water (Thermo Fisher Scientific, catalog number: 022934)
Microscopes slides, 75 mm × 25 mm × 1 mm (Thermo Fisher Scientific, catalog number: 12-544-2)
Cover glass, 22 mm × 22 mm, No. 1.5 (Thermo Fisher Scientific, catalog number: 12541B)
Secure seal spacer, 9 mm diameter (Thermo Fisher Scientific, catalog number: S24737)
Isopropyl alcohol (IPA) (Millipore Sigma, catalog number: W292907)
5% (3-aminopropyl)triethoxysilane (APTES) (Millipore Sigma, catalog number: 440140-100ML)
25% Glutaraldehyde (Electron Microscopy Supplies, catalog number: 16220)
1.5 mL Polypropylene microfuge tubes (Beckman Coulter, catalog number: 357448)
Ethanol 200 proof (Thermo Fisher Scientific, catalog number: T038181000)
Molecular-grade ethanol (Electron Microscopy Sciences, catalog number: 15055)
HPLC-grade ethanol (Thermo Scientific Chemicals, catalog number: AS611050040)
Ibidi Grid-50 cover glasses (Ibidi, catalog number: 10817)
Attofluor cell chamber (Thermo Fisher, catalog number: A7816)
Aluminum mounts to fit in SEM (Electron Microscopy Sciences, catalog number: 75185)
Ultra-smooth carbon adhesive tabs, 25 mm diameter (Electron Microscopy Sciences, catalog number: 778727-25)
PELCO conductive silver paint (Ted Pella, Inc, catalog number: 16062-15)
Wash-N-DryTM coverslip rack (VWR, catalog number: 490007-150)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D4540)
Solutions
Vesicle isolation buffer (VIB) (see Recipes)
Calcein AM stain (see Recipes)
Potomac gold stain (see Recipes)
Trypan blue (see Recipes)
Glutaraldehyde fixative (see Recipes)
Ethanol dehydration solutions (see Recipes)
Recipes
Vesicle isolation buffer (VIB)
20 mM MES
2 mM CaCl2
0.1 M NaCl
Adjust pH to 6.0. Autoclave at 121 °C for 30 min
Calcein AM Stain
Resuspend 50 µg of Calcein Green, AM in 50 µL of DMSO.
Dilute to 1 µM in HPLC water.
Potomac gold stain
Resuspend 100 µg of Potomac Gold in 100 µL of DMSO.
Make 5 µL aliquots.
Dilute to 1 µM in HPLC water.
Trypan blue
Make a 100 nM stock solution in HPLC water.
Glutaraldehyde fixative
Make fresh.
Filter 50 mL of 20 mM glutaraldehyde, pH 7.5, through a 0.2 µm filter into a 50 mL conical tube.
Make 1% and 2% glutaraldehyde in filtered 20 mM Tris-HCl, pH 7.5.
Ethanol dehydration solutions
Make a 25%, 50%, 75%, and 95% ethanol concentration series by diluting HPLC-grade ethanol in HPLC water.
Equipment
Light meter (Li-Cor, model: Quantum LI-190-R and LI-350)
Vacuum pump (Cole Parmer Welch, model: 2585B-50)
Vacuum desiccator chamber (Millipore Sigma, model: F42400-2221, catalog number: BAF424002221)
Avanti J-20 centrifuge (Beckman Coulter, catalog number: 368608)
JA-14 fixed-angle rotor (Beckman Coulter, model: JA-14, catalog number: 339247)
JA-17 fixed angle rotor (Beckman Coulter, model: JA-17, catalog number: 369691)
OptimaTM L-90k ultracentrifuge (Beckman Coulter, catalog number: 2043-30-1191)
SW 41 Ti rotor, swinging bucket, titanium, 6 × 13.2 mL (Beckman Coulter, catalog number: 331362)
OptimaTM MAX tabletop ultracentrifuge (Beckman Coulter, catalog number: 393315)
TLA 55 fixed-angle rotor package (Beckman Coulter, catalog number: 366725)
Total internal reflection fluorescence microscope (TIRF) (Oxford Instruments, model: Andor Dragonfly 600)
Note: Other sensitive microscopes can be used; however, TIRF and spinning disk confocal microscopes are recommended. Here, we used Borealis TIRF on an Andor Dragonfly 600 microscope with an Andor Zyla 4.2 PLUS sCMOS camera and a Leica Plan Apo 63× oil immersion TIRF objective (numerical aperture, 1.47). We used a 2,000mW, 488nm excitation laser, and 521/38nm bandpass (BP) filter for Calcein AM and a 1,000mW, 561nm excitation laser, and 594/43nm BP filter for Potomac Gold.
Stainless steel tweezers (Tronex, catalog number: SA316L)
Cover glass drying rack (Millipore Sigma, catalog number: Z743684)
Sonicator (Elma, catalog number: 51000065766)
Critical point dryer (Tousimis, model: Autosamdri®-815B, Series A, catalog number: 8780B)
Apreo VolumeScopeTM scanning electron microscope with Maps 3.9 software
Software and datasets
ImageJ
Procedure
Note: An overview of the sorghum EV isolation workflow can be found in Figure 1.
Figure 1. General workflow. Images showing the continuous processes for sorghum extracellular vesicles (EV) isolation and subsequent density gradient purification. Total process time is ~24 h. Vesicle isolation buffer (VIB), vacuum apoplastic wash (vAW), centrifuge apoplastic wash (cAW).
Growing sorghum and collecting leaf tissue
Sow sorghum seeds in a 6” pot containing pro-mix BK55 growing medium.
Grow for 14 days in a greenhouse or growth chamber with 16/8 h light/dark cycle at a temperature of 27 and a light intensity of 350 µmol/m2/s. Maintain 65% relative humidity.
Before harvesting leaves, cut 0.5 cm of the leaf tips of each plant to allow for better vacuum infiltration of the VIB buffer.
Harvest the plants above the soil with scissors and then rinse in water twice to remove damaged plant materials and soil residue.
Leaf infiltration and vacuum apoplastic wash collection
Plants should be submerged in batches for vacuum infiltration in a 1 L beaker containing 300 mL of VIB buffer. Place the beaker inside the vacuum chamber at -0.1 MPa for 20 s; this should be repeated five times, shuffling the leaves after every cycle.
Note: Ensure all the plants are fully submerged in VIB buffer. Place a glass Petri dish on the plant to keep the leaves fully submerged.
Collect the plants in small batches and wrap them into bouquets with a small strip of parafilm.
Filter the remaining buffer through Miracloth to remove debris and store the vacuum apoplastic wash (vAW) on ice.
Centrifugal apoplastic wash collection, debris removal, and EV pelleting
Modify a 60 mL syringe by cutting off the end and wrapping it with 3 cm of parafilm (Figure 2).
Figure 2. Apoplastic wash collection. Image showing 250 mL Nalgene bottles with 60 mL syringe modification for centrifuge apoplastic wash (cAW) isolation from Sorghum bicolor plants. The syringe is wrapped with parafilm (3 cm) to create a stopper between the tube and the syringe. The layer of parafilm is approximately 3 mm thick.
Widen the opening of the 250 mL Nalgene bottle with an 8-inch half-round file until the 60 mL syringe smoothly slides into the bottle.
Place the bouquets of infiltrated leaves stem down inside 60 mL syringes and load them into 250 mL centrifuge bottles.
Centrifuge the 250 mL bottles containing the infiltrated leaves at 700× g for 30 min at 4 to generate the centrifuge apoplastic wash (cAW) using a JA-14 rotor in an Avanti J-26S XP centrifuge.
Combine the cAW with the vAW and load into 50 mL polypropylene centrifuge tubes.
Centrifuge at 10,000× g for 1 h at 4 using the JA-17 rotor in an Avanti J-26S XP centrifuge.
Decant the supernatant as soon as the tubes are removed from the rotor into new 50 mL polypropylene centrifuge tubes.
Centrifuge at 39,800× g for 2 h at 4 °C. Carefully decant the supernatant and discard, resuspend the pellets in chilled VIB buffer, and pool together.
Note: Before decanting, mark the side of the tube close to the wall of the rotor with a sharpie; that is where the majority of the pellets are settled. This step is done beside the centrifuge to prevent agitation of the pellet settled on the side of the tube.
Centrifuge the resulting solution at 39,800× g for 2 h at 4 °C.
Decant the supernatant and resuspend the pellet (P40) in 100–250 µL of 0.2 µm-filtered 20 mM Tris-HCl, pH 7.5.
Density gradient and EV fraction separation
Using 60% aqueous iodixanol stock solution, prepare 40% (v/v), 20% (v/v), 10% (v/v), and 5% (v/v) iodixanol dilutions in VIB buffer.
Prepare a discontinuous density gradient column by carefully layering 3 mL of 40% solution, 3 mL of 20% solution, 3 mL of 10% solution, and 2 mL of 5% solution in a 14 mm × 89 mm (13.2 mL) thin-walled ultra-clear centrifuge tube.
Add 100–250 µL of the crude EV pellet resuspended in 20 mM Tris-HCl, pH 7.5 to the top of the discontinuous density gradient.
Centrifuge at 100,000× g for 17 h at 4 °C and at the maximum acceleration and deceleration using a SW 41 Ti rotor in an Optima L-90K.
After ultracentrifugation, discard 4.5 mL of the solution from the top of the gradient and collect the next 2.1 mL fraction from the gradient. Divide this fraction into six 350 µL aliquots in Beckman Coulter 1.5 mL microfuge tubes. Bring the volume in each tube up to 1.5 mL using VIB buffer.
Centrifuge the tubes at 100,000× g for 1 h at 4 °C using a TLA 55 rotor and an Optima Max ultracentrifuge.
Resuspend the pellets from the tubes and pool them together with 250 µL of Tris-HCl, pH 7.5. Bring the volume up to 1.5 mL using Tris-HCl, pH 7.5.
Centrifuge again at 100,000× g for 1 h at 4 °C. The final pellet containing the gradient-purified EV (GPEV) fraction will be resuspended in 50 µL of 20 mM Tris-HCl, pH 7.5. Keep the purified EVs at 4 °C and ensure downstream assays are done within 48 h.
Labeling and imaging of EVs
Prior to imaging, place 0.5 mL of poly-L-lysine on a clean cover glass and incubate for 30 min at room temperature (RT) (see cover glass cleaning protocol in Section F).
Aspirate poly-L-lysine off the cover glass.
Note: Ensure this is done from the edge of the cover glass to avoid scratching the surface of the cover glass after poly-L-lysine has been added.
Wash the cover glass three times with copious HPLC-grade water.
Note: This can be done by dipping the cover glass in HPLC-grade water using forceps.
Arrange the cover glass in a dish with the coated side up.
Bake at 50 °C for 1 h. Let the cover glass cool to RT.
Cut a secure seal spacer in half, remove the paper backing, and mount on a cover glass (Figure 3).
Figure 3. Picture showing stained extracellular vesicle (EV) samples mounted on a microscope slide partitioned with a secure seal spacer and sealed with a poly-L-lysine-treated 22 × 22 cover glass. EV samples were stained with Calcein AM and Potomac Gold. Trypan blue was added to quench unbound Potomac Gold dye.
In a 1.5 mL tube, mix 5 µL of EVs with 5 µL of 1 µM Calcein AM stain. Incubate at RT for 30 min.
Apply 1 µL of labeled EVs to an individual secure seal spacer well on the poly-L-lysine-coated cover glass.
Add 1 µL of 1 µM Potomac Gold stain to a final concentration of 500 nM.
Quench the background fluorescence of Potomac Gold adding an equal volume of 100 nM trypan blue for a final concentration of 50 nM.
Note: Using only 4 µL prevents mixing between wells.
Remove the waxed paper secure seal spacer cover to expose the other side of the adhesive sticker and apply a clean microscope slide on top.
Image the samples using total internal reflection fluorescence (TIRF) or spinning disk confocal microscopy (Figure 4). Use typical excitation and emission settings for green-fluorescent dyes for Calcein AM and red-fluorescent dyes for Potomac Gold.
Note: See Figure 4 for green and red fluorescent imaging parameters we used on the Andor Dragonfly 600. Standard 488 nm and 561 nm excitation can be used for green and red, respectively, and the fluorescence signal should be excited and collected sequentially.
Figure 4. Calcein AM and Potomac Gold staining of extracellular vesicles (EVs) isolated from Sorghum bicolor. Trypan blue (50 nM) was added to quench the unbound Potomac Gold. Samples were imaged on an Andor Dragonfly 600 microscope with total internal reflection fluorescence (TIRF) using a 500 ms exposure time and four averaging on a Zyla 4.2 PLUS sCMOS camera. For Calcein AM, 3% 488nm excitation (3.3 mW) and a 521/38 nm bandpass (BP) filter (TR-DFLY-F521-038) were used. For Potomac Gold, 5% 561 nm laser (4.0 mW) and 594/43 nm BP filter (TR-DFLY-F594-043) were used.
Cover glass cleaning and modification for correlative light and electron microscopy
Put the cover glasses in a Wash-N-DryTM coverslip rack and change the solutions in a 100 mL glass beaker.
Sonicate cover glasses for 30 min in 95% IPA, covering the beaker with parafilm to reduce evaporation.
Wash five times with 10 mL of Milli-Q H2O for a total of 50 mL.
Sonicate with 5 N NaOH and cover with parafilm.
Note: Do not use aluminum foil for this step. The NaOH will react with the foil and create aluminum oxide and hydrogen gas.
Wash five times with Milli-Q H2O.
Sonicate with 100% EtOH for 30 min.
Wash with copious amounts of HPLC water and place in a drying rack, skipping spaces between cover glasses to allow for airflow.
Air dry at RT for at least 1 h or bake at 50 °C for 30 min.
Place the cover glass grid side up (arrow on the left-hand side pointing away from you) on standard microscope slides that are taped to a 10 cm Pyrex Petri dish (Figure 5).
Figure 5. A 10 cm glass Pyrex Petri dish with standard microscope glass slides taped in place. A) Ibidi cover glasses with an edge hanging off of a slide to allow for easier manipulation with forceps. B) Representative image showing solutions added to the top of the cover glass. C) Petri dish covered during incubation steps to reduce evaporation. Cover the cover glass with 1 mL of 100% IPA. Incubate for 1 min and remove.
Bake off the excess IPA at 50 °C for 30 min and cool the dried cover glass to RT.
Passivate the clean cover glasses with 0.2 µm-filtered 0.1 N NaOH for 20 min.
Hold the cover glass with forceps and rinse three times with 15 mL of HPLC water.
Remove excess water.
Bake at 50 °C and let cool to RT.
Apply 5% APTES in 95% IPA for 10 min.
Using a pipette, aspirate off the remaining APTES solution.
Hold the cover glass with forceps and rinse 5× with 15 mL of HPLC water. Between rinses, put down the cover glass and pick up from a different location to ensure no APTES is left behind.
Notes:
If modification was successful, water should slide off the cover glass.
If APTES is not washed off completely, the residual will generate a brown precipitate that will interfere with imaging.
Let the cover glass dry completely for at least 1 h at RT.
Apply 1 mL of 1% glutaraldehyde fixative (see Recipes). Incubate for 1 h at RT.
Remove glutaraldehyde fixative.
EV attachment and imaging by light microscopy
Add 5 µL of 1 µM Calcein AM stain to 5 µL of EVs and incubate at RT for 30 min.
Add 5 µL of 1 µM Potomac Gold. Incubate for 5 min.
Add 10 µL of 100 nM trypan blue. Incubate for 5 min.
Remove excess and gently wash 3× with 20 mM Tris-HCl, pH 7.5.
Note: This will remove the vesicles that have not covalently attached to the cover glass. Unattached vesicles will show up on light microscopy but will vanish in electron microscopy processing.
Immediately add 0.5 mL of additional Tris-HCl, pH 7.5 to prevent drying.
Note: Do not let vesicles dry to glass, it will change the morphology.
Place into an Attofluor cell chamber and add additional Tris-HCl, pH 7.5 as necessary.
Use total internal reflection microscopy (TIRFM) or spinning disc confocal with a 63× 1.4 NA oil objective, taking a tiled z-stack to capture the grid location on the cover glass.
Deconvolve and stitch the images.
Carefully remove most of the Tris-HCl and slowly add 200 µL of 2% glutaraldehyde fixative.
Note: For all of the remaining steps, do not pipette over the sample to prevent dislodging the EVs.
Incubate for 30 min.
Remove glutaraldehyde fixative and gently wash 3× with HPLC water.
Critical point drying
Remove HPLC water and gently add 25% HPLC-grade ethanol. Incubate for 5 min.
Remove solution and add 50% HPLC-grade ethanol. Incubate for 5 min.
Remove solution and add 75% HPLC-grade ethanol. Incubate for 5 min.
Remove solution and add 95% HPLC-grade ethanol. Incubate for 5 min.
Remove solution and add 100% molecular-grade ethanol. Incubate for 5 min.
Remove solution and add 100% molecular-grade ethanol. Keep in ethanol until loaded into a critical point dryer.
Fill the Tousimis Autosamdri critical point dryer with 100% molecular grade ethanol.
Run samples following the manual with the purge timer set to 3.
Scanning electron microscopy (SEM) correlative imaging
Mount samples on an aluminum SEM stub with ultra-smooth carbon adhesive tabs and coat with a 5 nm layer of platinum using a sputter coater (Leica ACE600 coater) (Figure 6).
Brush conductive silver paint on the edge of the coated cover glass to reduce charging on the glass.
Load the sample into the SEM and image (Figure 6).
Figure 6. Sputter-coated and mounted cover glass. PELCO conductive silver paint was brushed on to connect the top and bottom of the glass to prevent charging on the sample.
Image processing and correlation
Load the .ims image file taken on the Dragonfly microscope and stitch them using the Imaris Stitcher 9.0. Save the merged image as a new file.
Open the .ims file in FIJI and find the five slices of the Z-stack that contain Calcein AM and Potomac Gold stained EVs. Subset these slices.
Run Z project, select Max intensity, and hit OK.
Adjust the brightness and contrast to clearly show the vesicles.
If this is a multichannel image, save both the merged as well as splitting each channel individually.
Select each individual channel image and merged channels and run Flatten. This will set the display settings and merge the channels into a single flat image. This is necessary to view the image in the MAPS software.
Save the new image as a .tif.
Mount the cover glass on a stub (Figure 6) and load into the SEM. Put under vacuum, image with a fast scan to navigate, and find the grid. Select a region that has a good distribution of EVs and take a tiled high-resolution image with a 4,096 × 4,096 image size, a 2 nm pixel, and 1 µs dwell time.
Note: For increased correlation success, ensure the tiled region overlaps with the laser-etched portions of the grid.
Import the maximum intensity projection from ImageJ into MAPS 3.9
In the MAPS software (v3.9) on the Apreo SEM, follow the correlation wizard. Use the corners of the LM and SEM grid squares for the first rough alignment, and then refine the alignment using the EVs (Figure 7).
Figure 7. Correlative light and electron microscopy with Calcein AM and Potomac Gold labeled extracellular vesicles (EVs). A) Large overview scanning electron microscopy (SEM) image gridded section that was previously imaged via spinning disk or total internal reflection fluorescence (TIRF) microscopy. B) An overlay image showing the TIRF image of Calcein AM (Green) and Potomac Gold (magenta) labeled EVs data aligned with the SEM image. Images were aligned in MAPS software version 3.9. C) Inset showing a magnified view of the boxed area in (B).
Image correlation with FIJI ImageJ (alternate protocol)
Note: For our analysis, we used the MAPS software as described in section J (steps 7–10). However, the MAPS software is limited to Thermo Fisher SEMs and, therefore, we have provided an alternate protocol for image correlation using FIJI ImageJ.
Load both the SEM and light microscopy images into FIJI.
Convert both images to 16-bit.
Using the line tool, draw a line connecting two points in each image that should correlate.
Run Align image by ROI. Set the LM image as the source and the SEM image as the target. Select Scale and Rotate. Click OK. This will generate a 32-bit grayscale image in a new window.
Convert the new image to 16-bit.
Run Merge Channels.
Set the aligned image as a colored channel and the SEM image as gray. Check Create composite and Keep source images. Keeping the source images is necessary if you want to go back and refine the aligned image.
Repeat using the line tool and Align image by ROI to refine the alignment.
Validation of protocol
This protocol or a part of it has been used and validated in the following research articles:
Chaya et al. [19]. The extracellular vesicle proteomes of Sorghum bicolor and Arabidopsis thaliana are partially conserved. Plant Physiology (Figures 1–5).
Acknowledgments
This protocol was adapted from Chaya et al. [19]. This work was supported by a grant from the Department of Energy (DOE) Office of Biological and Environmental Research (BER) program DOE (DE-SC0020348) and in part by a seed grant from the College of Agriculture and Natural Resources at the University of Delaware. Microscopy equipment and data storage was acquired by shared instrumentation grants from the National Institutes of Health (S10 OD030321, S10 OD025165, and NIH S10 OD028725), and microscopy access was supported by grants from the National Institute of General Medical Sciences (P20 GM103446 and P20 GM139760) and the State of Delaware.
Competing interests
There are no conflicts of interest or competing interest.
References
Baldrich, P., Rutter, B. D., Karimi, H. Z., Podicheti, R., Meyers, B. C. and Innes, R. W. (2019). Plant Extracellular Vesicles Contain Diverse Small RNA Species and Are Enriched in 10- to 17-Nucleotide “Tiny” RNAs. Plant Cell. 31(2): 315–324.
Cui, Y., Gao, J., He, Y. and Jiang, L. (2020). Plant extracellular vesicles. Protoplasma. 257(1): 3–12.
Cai, Q., Qiao, L., Wang, M., He, B., Lin, F. M., Palmquist, J., Huang, S. D. and Jin, H. (2018). Plants send small RNAs in extracellular vesicles to fungal pathogen to silence virulence genes. Science. 360(6393): 1126–1129.
Abubakar, Y. S., Sadiq, I. Z., Aarti, A., Wang, Z. and Zheng, W. (2023). Interplay of transport vesicles during plant-fungal pathogen interaction. Stress Biol. 3(1): 35.
Halperin, W. and Jensen, W. A. (1967). Ultrastructural changes during growth and embryogenesis in carrot cell cultures. J Ultrastruct Res. 18: 428–443.
Rutter, B. D. and Innes, R. W. (2023). Extracellular Vesicles in Phytopathogenic Fungi. Extracell Vesicles Circ Nucl Acids. 4(1): 90–106.
Hua, C., Zhao, J. H. and Guo, H. S. (2018). Trans-Kingdom RNA Silencing in Plant–Fungal Pathogen Interactions. Mol Plant. 11(2): 235–244.
Huang, C. Y., Wang, H., Hu, P., Hamby, R. and Jin, H. (2019). Small RNAs – Big Players in Plant-Microbe Interactions. Cell Host Microbe. 26(2): 173–182.
Iwakawa, H. o. and Tomari, Y. (2013). Molecular Insights into microRNA-Mediated Translational Repression in Plants. Mol Cell. 52(4): 591–601.
Wang, S., He, B., Wu, H., Cai, Q., Ramírez-Sánchez, O., Abreu-Goodger, C., Birch, P. R. and Jin, H. (2024). Plant mRNAs move into a fungal pathogen via extracellular vesicles to reduce infection. Cell Host Microbe. 32(1): 93–105.e6.
Weiberg, A., Wang, M., Lin, F. M., Zhao, H., Zhang, Z., Kaloshian, I., Huang, H. D. and Jin, H. (2013). Fungal Small RNAs Suppress Plant Immunity by Hijacking Host RNA Interference Pathways. Science. 342(6154): 118–123.
Ghosh, S., Regmi, K. C., Stein, B., Chen, J., O’Connell, R. J. and Innes, R. W. (2024). Infection of Alfalfa Cotyledons by an Incompatible but Not a Compatible Species of Colletotrichum Induces Formation of Paramural Bodies and Secretion of EVs. bioRxiv. doi.org/10.1101/2024.04.28.591504.
De Palma, M., Ambrosone, A., Leone, A., Del Gaudio, P., Ruocco, M., Turiák, L., Bokka, R., Fiume, I., Tucci, M., Pocsfalvi, G., et al. (2020). Plant Roots Release Small Extracellular Vesicles with Antifungal Activity. Plants. 9(12): 1777.
Garcia-Ceron, D., Lowe, R. G. T., McKenna, J. A., Brain, L. M., Dawson, C. S., Clark, B., Berkowitz, O., Faou, P., Whelan, J., Bleackley, M. R., et al. (2021). Extracellular Vesicles from Fusarium graminearum Contain Protein Effectors Expressed during Infection of Corn. J Fungi. 7(11): 977.
Rutter, B. D. and Innes, R. W. (2016). Extracellular Vesicles Isolated from the Leaf Apoplast Carry Stress-Response Proteins. Plant Physiol. 173(1): 728–741.
Robatzek, S. (2007). Vesicle trafficking in plant immune responses. Cell Microbiol. 9(1): 1–8.
Rodrigues, M. L., Nosanchuk, J. D., Schrank, A., Vainstein, M. H., Casadevall, A. and Nimrichter, L. (2011). Vesicular Transport Systems in Fungi. Future Microbiol. 6(11): 1371–1381.
Liu, Y., Wu, S., Koo, Y., Yang, A., Dai, Y., Khant, H., Osman, S. R., Chowdhury, M., Wei, H., Li, Y., et al. (2020). Characterization of and isolation methods for plant leaf nanovesicles and small extracellular vesicles. Nanomed Nanotechnol Biol Med. 29: 102271.
Chaya, T., Banerjee, A., Rutter, B. D., Adekanye, D., Ross, J., Hu, G., Innes, R. W. and Caplan, J. L. (2023). The extracellular vesicle proteomes of Sorghum bicolor and Arabidopsis thaliana are partially conserved. Plant Physiol. 194(3): 1481–1497.
Article Information
Publication history
Received: Jun 7, 2024
Accepted: Aug 19, 2024
Available online: Sep 10, 2024
Published: Oct 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Plant Science > Plant cell biology > Organelle isolation
Plant Science > Plant biochemistry > Lipid
Biochemistry > Lipid > Extracellular lipids
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5,084 | https://bio-protocol.org/en/bpdetail?id=5084&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
An Improved Focus-Forming Assay for Determination of the Dengue Virus Titer
MM Maharah Binte Abdul Mahid
PB Pradeep Bist
KS Kristmundur Sigmundsson
MM Muhammad Danial Bin Mohd Mazlan
Satoru Watanabe
MC Milly M. Choy
SV Subhash G. Vasudevan
KC Kitti Wing Ki Chan
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5084 Views: 333
Reviewed by: Luis Alberto Sánchez VargasRan ChenDebashis Dutta
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Abstract
Dengue virus (DENV), a common and prevalent mosquito-borne endemic disease, is caused by four serotypes (DENV-1–4) and has spread rapidly on a global scale over the past decade. A crucial step in the development of antiviral therapeutics requires the utilization of in vitro cell-based techniques, such as plaque assays and focus-forming assays (FFA) for virus quantification. Vero cells have been widely used for FFA and plaque assay; however, there are instances when their efficacy and efficiency in the detection of certain clinical DENV isolates are low. Here, we showed that BHK-21 cells are more sensitive than Vero cells in the detection of all DENV-1–4 plaques and foci. In addition, we developed an improved FFA protocol for the quantification of all four DENV serotypes. Using a pan-flavivirus envelope (E) antibody, we reduce the possibility of false positives by defining a focus to consist of a minimum of eight infected cells. We outlined a protocol using the Operetta® high-content imaging system to automate the digital capture of these infected cells. A pipeline was also designed using the CellProfilerTM automated image analysis software to detect these foci. We then compare the results of the improved FFA with plaque assay. Notably, the improved FFA detected clear foci of the DENV-4 strain that does not form distinct plaques. We subsequently demonstrated the potential application of the improved FFA protocol in antiviral testing, utilizing a nucleoside inhibitor of DENV, NITD008 as a control. The protocol is amenable to a diverse array of applications, including high-throughput compound screening (HTS).
Key features
• An improved focus-forming assay that has the potential to quantify the DENV-4 strain, which was previously hard to plaque.
• Improvements have been made to reduce the possibility of false positives.
• Improved workflow using automated digital imaging process and counting of foci.
• Applicable to antiviral compounds screening and is amenable to high-throughput screening.
Keywords: DENV clinical isolates Plaque assay Focus-forming assay BHK-21 Vero Automated imaging Automated foci counting Antiviral compound screening
Background
Dengue represents a significant challenge to global public health systems, with over 40% of the global population at risk of infection. It is estimated that approximately 400 million people are infected with dengue virus (DENV) each year, of whom 100 million manifest clinical symptoms of dengue hemorrhagic fever with 25,000 deaths reported annually [1,2]. The symptoms include the abrupt onset of fever, which is followed by a multitude of other critical complications, including circulatory failure, hemorrhages in the skin and gastrointestinal tract, and shock. DENV comprises four antigenically distinct serotypes, exhibiting variations in both structural and nonstructural proteins. The four antigenically distinct serotypes of dengue virus are designated DENV-1, DENV-2, DENV-3, and DENV-4 [3,4]. The necessity for efficient, accurate, and robust assays for the quantification of infectious viruses, which contribute to the discovery of antiviral drugs, has thus increased. A variety of techniques have been explored and developed over the years, with plaque and/or focus-forming assays (FFA) still representing the gold standard for such assays. A variety of cell lines have been employed in these assays, with Vero (African green monkey kidney fibroblasts) cells being the most frequently utilized in plaque assay [5]. Furthermore, a number of optimization protocols for FFA have been documented for a range of viruses, including DENV [6,7]. However, this method typically necessitates a great number of steps, including the use of multiple distinct monoclonal antibodies for foci visualization; moreover, the method was not extended to DENV-3 and DENV-4 detection. In this work, we delineate the methodology for employing the FFA for the quantification of infectious virus particles using E protein detection in all serotypes and compare it with plaque assay quantification of infectious virus production. We demonstrate that the FFA approach was more sensitive than the plaque assay in detecting DENV-4, which exhibited poor plaque formation. In addition, all four DENV serotypes form clearer plaques and foci in BHK-21 cells than in Vero cells. Moreover, improvements have been made to the existing FFA. For example, we defined a focus to comprise a minimum of eight cells, thereby reducing the possibility of false positives. We also outlined a protocol using the Operetta® high-content imaging system and a pipeline using the CellProfilerTM software to automate foci imaging and foci counting. In addition, we demonstrate the immediate application of FFA as a tool for assessing the antiviral efficacy of compounds using a known adenosine nucleoside DENV inhibitor, NITD008 [9,10], as a control. Altogether, the improved FFA is a precise and reliable assay not only for quantifying clinical isolates but also for high-throughput screening of compounds for dengue drug discovery.
Materials and reagents
Biological materials
Representative Dengue virus strains from EDEN study [8]: DENV-1 clinical isolate (EDEN1: GenBank accession EU081230), DENV-2 clinical isolate (EDEN2: GenBank accession EU081177), DENV-3 clinical isolate (EDEN3: GenBank accession EU081190), DENV-4 clinical isolate (EDEN4: GenBank accession GQ398256)
Aedes albopictus C6/36 cell line (ATCC, CRL-1660TM)
Baby hamster kidney fibroblast, BHK-21 (ATCC, CCL-10TM)
African green monkey kidney fibroblast, Vero cell line (ATCC, CCL-81TM)
Hybridoma cells; 4G2 (ATCC, HB-112TM)
Reagents
RPMI1640 medium (Thermo Fisher Scientific, Gibco®, catalog number: 11875093)
DMEM (1×), high glucose, pyruvate medium (Thermo Fisher Scientific, Gibco®, catalog number: 11995-040)
Heat-inactivated fetal bovine serum (FBS) (Thermo Fisher Scientific, Gibco®, catalog number: 10082147)
200 mM L-glutamine (Thermo Fisher Scientific, Gibco®, catalog number: 25030081)
Penicillin and streptomycin (pen/strep) (Thermo Fisher Scientific, Gibco®, catalog number: 15140122)
1 M HEPES (Thermo Fisher Scientific, Gibco®, catalog number: 15630-080)
0.25% Trypsin-EDTA (Thermo Fisher Scientific, Gibco®, catalog number: 25200-056)
Protein-free hybridoma medium (PFHM-II medium) (Thermo Fisher Scientific, Gibco®, catalog number: 12040-077)
1× PBS (1st BASE, catalog number: BUF-2040-10X1L)
Ethanol (EtOH) molecular biology grade (Merck, Sigma-Aldrich, catalog number: 51976)
Hydrochloric acid (Merck, Sigma-Aldrich, catalog number: 258148)
RPMI1640 powder (Thermo Fisher Scientific, Gibco®, catalog number: 31800-022)
DMEM powder, high glucose (Thermo Fisher Scientific, Gibco®, catalog number: 12100046)
Sodium Bicarbonate (Thermo Fisher Scientific, catalog number: 25080094)
Tris (1st BASE, catalog number: BIO-1400)
Glycine (Merck, Sigma-Aldrich, catalog number: G7126)
Triton X-100 (Bio-Rad, catalog number: 161-0407)
Bovine serum albumin (BSA), heat shock isolation (Bio Basic, catalog number: AD0023)
Sodium azide (Merck, Sigma-Aldrich, catalog number: S8032)
DAPI (Merck, Sigma-Aldrich, catalog number: D9542)
37% Formaldehyde solution (Merck, Sigma-Aldrich, catalog number: F1635)
Crystal violet (Merck, Sigma-Aldrich, catalog number: C3886)
Alexa FluorTM 488 F(ab’) 2 fragment of goat anti-mouse IgG (H + L) (Thermo Fischer Scientific, Invitrogen, catalog number: A11017)
Alexa FluorTM 594 F(ab’) 2 fragment of goat anti-mouse IgG (H + L) (Thermo Fischer Scientific, catalog number: A-11020)
Alexa FluorTM 488 Phalloidin (Thermo Fisher Scientific, catalog number: A12379)
Solutions
0.8% methyl-cellulose medium supplemented with 2% FBS (see Recipes)
1% crystal violet (see Recipes)
0.1 M glycine (pH 2.7) (see Recipes)
1 M Tris- HCl (pH 9.0) (see Recipes)
3.7% formaldehyde (see Recipes)
10% formaldehyde (see Recipes)
Recipes
0.8% methyl-cellulose medium supplemented with 2% FBS
Add 8 g of methyl-cellulose powder into 500 mL of water and autoclave twice to dissolve the powder.
Prepare 500 mL of 2× RPMI1640 or DMEM media by dissolving RPMI1640 powder or DMEM powder in water followed by supplementing with 4% heat-inactivated FBS, 4 mM L-glutamine, 200 U/mL pen/strep, 0.075% sodium bicarbonate solution, and 50 mM HEPES.
After filtration of 2× RPMI1640 or DMEM media through a 0.2 μm membrane filter unit, mix well with 500 mL prepared methyl-cellulose.
Store at 4 °C.
1% crystal violet
Add 5 g of crystal violet to 100 mL of 100% EtOH and mix well to dissolve powder.
Add 400 mL of water.
Store at room temperature.
0.1 M glycine (pH 2.7)
Add 3.75 g of glycine into 500 mL of water.
Adjust the pH to 2.7 using 1 N HCl.
Store the solution at 4 °C
1 M Tris-HCl (pH 9.0)
Add 30.3 g of Tris into 250 mL of water.
Adjust the pH to 9.0 using 1 N HCl.
Store the solution at room temperature.
3.7% formaldehyde
Add 500 mL of 37% formaldehyde into 4,500 mL of Milli-Q water.
10% formaldehyde
Add 200 mL of 37% formaldehyde into 540 mL of Milli-Q water.
Laboratory supplies
NunclonTM MULTIDISH 24 (Thermo Fisher Scientific, catalog number: 142475)
NunclonTM MULTIDISH 48 (Thermo Fisher Scientific, catalog number: 150687)
PerkinElmer cell carrier ultra microplates, treated, black, 96-well with lid (Genomax Technologies, catalog number: 6055302)
Greiner Bio-One MASTERBLOCKTM 96 deep-well conical-bottom 2 mL storage plate (Fisher Scientific, catalog number: 07-000-873)
Cryotubes vials for freezing viruses (Thermo Fisher Scientific, catalog number: 368632)
175 cm2 angled-neck easy flasks (Nunc, catalog number: 159920)
50 mL centrifuge tubes (BD Falcon, catalog number: 357550)
Filter unit 0.45 µm (Merck, Sigma-Aldrich, Millex®-HP, catalog number: SLHPR33RS)
Filter unit 0.2 μm (Thermo Fisher Scientific, catalog number: 567-0020)
HiTrap protein G HP-5 mL (GE Healthcare, catalog number: 170-0405-01)
Snakeskin dialysis tubing 10 kDa (Thermo Fisher Scientific, catalog number: 68100)
15 mL centrifuge tubes (Merck, Sigma-Aldrich, catalog number: CLS430791)
Aluminum foil
Equipment
Incubator without CO2 atmosphere at 28 °C (Sanyo, model: MIR-262)
Humidified incubator with 5% CO2 atmosphere at 37 °C (Nuaire, model: NU-5710E)
VACUSAFE aspiration system (Integra Biosciences, model: 158310)
ProBlotTM 25 economy rocker single platform (Bio Laboratories, model: S2025-B-230V)
Operetta® high content imaging system (PerkinElmer, model: Operetta)
Cermax® Xenon fiber-optic light source (Excelitas Technologies, model: XL3000)
DELL computer swinging rotor centrifuge (for cells) (Thermo Electron)
-80 °C freezer (Thermo Fisher Scientific)
Visi-White transilluminator (Analytik Jena US, model: TW-26)
Autoclave (TOMY, model: SX-700)
AKTA purifierTM UPC 10 (GE Healthcare, catalog number: 28406268)
pH meter (Satorius, catalog number: PY-PW-A)
NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, catalog number: ND-2000)
Software and datasets
Harmony v4.8 (Operetta, 1/1/2013)
CellProfilerTM (Broad institute, 03/12/2022)
Prism v10.0 (GraphPad, 08/23/2023)
Procedure
Generation of DENV stocks
Grow C6/36 cells in a 175 cm2 flask with RPMI1640 medium supplemented with 10% FBS, 2 mM L-glutamine, 100 U/mL pen/strep, and 25 mM HEPES and incubate at 28 °C under non-CO2-atmosphere-condition until ~90% confluency.
Note: Growth media supplemented with 20% FBS improves cell growth if C6/36 cells do not proliferate desirably.
Thaw virus stock and dilute with serum-free RPMI1640 medium.
Remove the medium from the culture flask and inoculate with 10 mL of virus inoculum at MOI of 0.1 into a 175 cm2 flask.
Incubate for 1 h at 28 °C under non-CO2 atmosphere conditions.
Discard the virus inoculum and add 25 mL of RPMI1640 medium supplemented with 2% FBS into the 175 cm2 flask.
Incubate for 5–7 days at 28 °C under non-CO2 atmosphere conditions.
Note: The cytopathic effect is observed depending on the virus strain (seen only in DENV-2 infection but not in other DENV strains infection). To ascertain the optimal duration for obtaining a high virus titer, it is recommended that the virus titer in the supernatant be determined at different time points. For example, the incubation period for DENV2 is five days, whereas it is seven days for DENV-1, DENV-3, and DENV-4.
Scrape the cells and transfer them into a 1.5 mL microcentrifuge tube.
Spin down the cells at 1,800× g for 10 min at 4 °C. Pass the supernatant through a 0.45 μm membrane filter and collect it in a fresh microcentrifuge tube.
Aliquot virus supernatant into cryotubes and store in a -80 °C freezer until use.
Determination of viral titer by plaque assay (Figure 1, Table 1, Figure 2)
Grow BHK-21 cells in RPMI1640 supplemented with 10% FBS, 2 mM L-glutamine, and 100 U/mL pen/strep or Vero cells in DMEM supplemented with 10% FBS, 4.5 g/L D-glucose, 2 mM L-Glutamine, and 110 mg/L sodium pyruvate and 100 U/mL pen/strep in a humidified incubator with 5% CO2 atmosphere at 37 °C.
Seed cells at 2 × 105 cells per well in 500 μL of supplemented RPMI1640 or DMEM medium into a 24-well plate.
Note: For detachment of cells during the cell seeding procedure, 2–3 mL of trypsin is used. Cells may be seeded at 5 × 104 or 2 × 104 cells per well in 500 μL for 100% confluency prior to infection after 2 or 3 days of incubation, respectively. Gently move the plate using short, back-and-forth, and side-to-side motions prior to placing it into the incubator to ensure an even distribution of cells.
Incubate cells overnight at 37 °C in a 5% CO2 incubator to allow cells to attach and reach 100% confluency.
Dilute the virus in a series of 10-fold dilutions in serum-free RPMI1640 or DMEM.
Note: Dilution in microfuge or centrifuge tubes is preferred to enable vertexing and spinning down in the centrifuge after diluting for a homogenous suspension. If unable to dilute in tubes, resuspend virus suspension with a pipette for a minimum of 30 times upon diluting.
Discard culture supernatant of BHK-21 cells or Vero cells and add 200 μL of the diluted virus into each well.
Note: Virus should be added immediately after removing culture supernatant to avoid cells drying out.
Figure 1. Flow diagram of plaque and improved focus-forming assays (FFA) to quantify dengue virus (DENV). The sequential steps were employed during plaque and FFA to quantify DENV1-4 clinical isolate titers. Following the seeding of BHK-21 or Vero cells onto 24- or 96-well plates, the plates were incubated overnight at 37 °C in 5% CO for the formation of a cell monolayer. The following day, the cells were infected with the respective virus and incubated for a period of 4–5 days (BHK-21) and 6–8 days (Vero) to allow for the development of plaques, and for a period of 2 days for both BHK-21 and Vero FFA plates, respectively. The plaque assay plates were fixed in 3.7% formaldehyde for a period of 12 h, stained with 1% crystal violet, and the number of plaques was counted manually. In contrast, FFA plates were fixed in 10% formaldehyde, permeabilized with 0.2% Triton X-100, blocked with 1% bovine serum albumin (BSA), and incubated with α-DENV E protein (4G2) (overnight) followed by the addition of Alexa FluorTM 488 antibody. Plates were then imaged using the Operetta® high-content imaging system and foci were counted using the automated System Virus (SV) Foci Counter imaging pipeline.
Table 1. Comparative analysis of the plaque assay and the improved focus-forming assay. The table illustrates the differences between the two assays with regard to virus quantification from a clinical standpoint, where virus quantification is of paramount importance. In such a scenario, FFA is a highly credible method and serves as a useful tool.
Plaque assay Focus forming assay (FFA)
Plate format 24-well 96-well
Sample volumes 200 µL 50 µL
Reagent volumes More volume of all reagents is required overall Less volume of all reagents is required overall
Assay duration Two weeks 1 week
Assay sensitivity for identification of viral strains Low (DENV-4 plaques are not clear) High (DENV-4 foci are clear)
Assay read-out Manual Automated
High throughput screening No Yes
Figure 2. BHK-21 and Vero cell lines serve as in vitro cell-based infection models for DENV1-4 clinical isolates quantification. (A) BHK-21 and Vero cells were used as in vitro infection models to quantify infectious virus production by plaque assay and focus-forming assay (FFA). Viral genomic copies were detected by quantitative reverse transcription polymerase chain reaction (qRT-PCR) as a positive indication of infection. After 2 days of incubation*, DENV-1–4 foci in BHK-21 were clearer, brighter, and larger compared to the foci seen in Vero cells for all the clinical isolates. DENV-1 formed larger foci in both BHK-21 and Vero cells, while single-cell infections were seen mostly for DENV-2 and DENV-4 in Vero cells. Dashed circles indicate representative foci for each serotype in the different cell lines. After 5 days of incubation, DENV-1–3 plaques were clear and large, while DENV-4 formed unclear plaques in BHK-21. No plaques were seen for DENV-1, DENV-3, and DENV-4; however, small plaques were seen for DENV-2 in Vero after 5 days. Plaques for DENV-1–3 in Vero cells were seen after 8 days of incubation. In addition, the presence of DENV-1–4 viral RNA was detected in the respective samples. Of note, the BHK-21 infectious model is more efficient and is preferred over Vero, and FFA is a viable alternative to detect DENV-4 that does not form proper plaques. Note: All plates were incubated at 37 °C in 5% CO2. FFU: focus-forming unit; PFU: plaque-forming unit. Green: E protein.
Incubate the plate for 1 h at 37 °C in a 5% CO2 incubator.
Note: Pre-equilibrate the 0.8% methyl-cellulose medium supplemented with 2% FBS to room temperature prior to use.
Discard the virus and overlay the cells with 500 μL of 0.8% methyl-cellulose medium supplemented with 2% FBS.
Note: The viscosity of methyl-cellulose medium necessitates the use of a Pasteur pipette for overlaying the medium onto the cells.
Incubate plate for 4–8 days at 37 °C in 5% CO2 incubator.
Note: The plaque size is affected by the virus replication rate. Check the plaque size visually before fixation to obtain clear plaque morphology. BHK-21-seeded plates of DENV-1 infection are normally incubated until day 4 post-infection, whereas plates of DENV-2–4 are usually incubated until day 5 post-infection. Vero seeded plates of DENV-1,3,4 infection were incubated for 8 days and DENV-2 was incubated for 5 days. However, it is recommended to incubate Vero-seeded plates of DENV-2 infection for more than 5 days as the plaques were small.
Fix cells with 500 μL of 3.7% formaldehyde (diluted in Milli-Q water) for a minimum of 3 h.
Note: Fixation overnight may be required if 3.7% formaldehyde is not freshly made.
Rinse the plate with a copious amount of running water to remove methyl-cellulose medium completely.
Add 1–2 drops of 1% crystal violet into each well and stain for 1 min.
Rinse the plate with copious amounts of running water to remove the excess crystal violet stain.
Air dry plates on a paper towel.
Visualize plaques using the white light transilluminator apparatus and count the number of plaques for determination of virus titer as follows:
Virus titer [plaque-forming units (pfu)/mL] = average number of plaques × 1,000 µL/200 µL inoculum × reciprocal of dilution factor
Preparation of α-DENV E protein antibodies (4G2) from hybridoma cells
Note: Antibodies specific to the detection of E protein of flaviviruses including DENV can be found commercially.
Culture 4G2 hybridoma cells in 50 mL of PFHM-II medium in 175 cm2 flasks in a humidified incubator with 5% CO2 atmosphere at 37 °C.
Note: It is recommended to culture the cells in RPMI1640 supplemented with 10% FBS, 2 mM L-glutamine, 100 U/mL pen/strep during the initial period of several days after thawing cells. Once the cells grow well, replace the media gradually by increasing the proportion of PFHM-II media to RPMI1640 from 10% (vol/vol) to 100% PFHM-II media.
Collect cell suspension into a centrifugal tube when the cells become confluent (the color of culture media turns to orange or yellow).
Centrifuge cells at 900× g for 5 min at room temperature.
Collect the supernatant and store it at 4 °C without filtration until a sufficient volume of supernatant is obtained.
Continue culturing the cells and repeat steps C2–C4 if a large volume of the supernatant is required.
Filter the supernatant through a 0.45 μm membrane filter unit.
Load the 4G2 supernatant onto a 5 mL Protein G column pre-equilibrated in PBS (pH 7.2).
Note: 4G2 antibody is purified using the AKTA purifier. Refer to the manufacturer’s instruction guides regarding sample loading specifications.
Wash the column with PBS using five times the column volume (i.e., 25 mL).
Prepare a Grenier 96-well master block containing 60 μL 1 M Tris-HCl pH 9.0 for collection.
Note: The standard ratio of Tris-HCl to glycine (100:6) for neutralization is subject to change depending on the concentration of buffers prepared. The volume of Tris-HCl required for neutralization (pH 7) can be adjusted by pH paper testing.
Elute antibodies using 100% 0.1 M glycine pH 2.7 and collect 1 mL fractions into the wells of the block.
Note: Check the purity of the antibody by running an SDS-PAGE.
Select fractions of high purity and collect them into a dialysis membrane; then, dialyze against PBS overnight.
Quantitate the concentration of the purified antibody using NanoDrop.
Determination of viral titer by FFA (Figure 1, Table 1, Figure 2, Figure 3)
Figure 3. Application of the BHK-21 infectious model in evaluating the efficacy of antiviral compound NITD008 against DENV1-4 clinical isolates using focus-forming assay (FFA). Huh7 cells were infected with DENV-1–4 at a multiplicity of infection (MOI) of 0.3 and subsequently treated with varying concentrations of NITD008 (100, 50, 10, 1, 0.1, and 0.01 µM). Post 48 h, viral supernatants were assayed on BHK-21 cells for FFA to estimate the virus titers. The half-maximal effective concentration (EC50) of NITD008 was calculated through the plotting of a sigmoidal dose-response curve using GraphPad Prism. The known EC50 of NITD008 lies between the range of 0.16–2.61 µM [9,10], and the range of calculated EC50 from both assays was 0.62–1.18 µM, indicating the efficiency of FFA. The data was obtained from three biological replicates and verified by an independent experimenter as being reproducible. The data presented here represents a single experiment out of a total of three (n = 3, SD = 0.02–0.78).
D1. Preparation of FFA samples for image acquisition
Maintain BHK-21 cells in RPMI1640 supplemented with 10% FBS, 2 mM L-glutamine, and 100 U/mL pen/strep or Vero cells in DMEM supplemented with 10% FBS, 4.5 g/L D-glucose, L-Glutamine and 110mg/L Sodium Pyruvate and 100 U/mL pen/strep in a humidified incubator with 5% CO2 atmosphere at 37 °C.
Seed cells at 5 × 104 cells per well in 100 μL into the PerkinElmer cell carrier 96 ultra microplates.
Note: Cells may be seeded at 1.25 × 104 or 5 × 103 cells per well in 100 μL for 100% confluency prior to infection after 2 days or 3 days incubation, respectively. A single-cell monolayer formation is crucial to allow the capture of focused images for accurate analysis.
Incubate cells overnight at 37 °C in a 5% CO2 incubator to allow cells to adhere and reach 100% confluency.
Prepare 10-fold serial dilutions of the virus in serum-free RPMI1640 or DMEM.
Remove the culture supernatant of BHK-21 cells or Vero cells and add 50 μL of diluted virus into each well.
Note: Virus should be added immediately after removing culture supernatant to avoid cells drying out.
Incubate the plate for exactly 1 h at 37 °C in a 5% CO2 incubator.
Remove the virus and add 125 μL of 0.8% methyl-cellulose medium supplemented with 2% FBS. Incubate the plates with infected BHK-21 cells at 37°C in a 5% CO2 incubator for two days.
Note: Plates with infected Vero cells will require more than 2 days incubation, as foci are not as clear as those seen in BHK-21 with mostly single-cell infections observed (Figure 2).
Add 100 µL/well of 10% formaldehyde (diluted in Milli-Q water) and incubate for 1 h at room temperature or 20 min in the 37 °C incubator.
Wash off the 10% formaldehyde thoroughly with a copious amount of water in a container. Shake the plate vigorously to remove the methyl-cellulose medium completely.
Add 100 µL/well of 0.2% Triton X-100 in PBS and incubate on ProBlotTM 25 economy rocker with maximum speed at room temperature for 20 min.
Note: For the prevention of microbial growth, you can optionally add 0.01% sodium azide to the 0.2% Triton-X-100.
Discard 0.2% Triton X-100 in 1× PBS and wash wells three times with 1× PBS.
Note: Remove as much 1× PBS as possible when washing each time and avoid drying of cells by ensuring wells are always filled with 1× PBS.
Add 125 µL/well of 1% BSA in 1× PBS and incubate on ProBlotTM 25 economy rocker at maximum speed at room temperature for 1–2 h.
Discard 1% BSA in 1× PBS and wash wells three times with 1× PBS.
Add 50 µL/well of purified mouse anti-E 4G2 antibody (final concentration of 2 µg/mL) in 1% BSA/PBS and incubate for at least 1 h on ProBlotTM 25 economy rocker at 60–70 rpm at room temperature or overnight at 4 °C.
Collect the purified mouse anti-E 4G2 antibody from each well in a new tube after incubation and keep at -30 °C.
Note: Diluted antibody can be reused approximately 2–3 times.
Wash wells three times with 150 µL of 1× PBS.
Add 50 µL/well of Alexa FluorTM 488 goat anti-mouse antibody with a 1:1,000 dilution in 1% BSA/PBS and incubate for 1 h on ProBlotTM 25 economy rocker at 60–70 at room temperature.
Note: Protect the antibody and sample plate from light by wrapping the tube holding the diluted antibody and sample plates with aluminum foil from this step onward. Do not incubate for more than 1 h as this may result in a high background signal.
Remove the secondary antibody (Alexa FluorTM 488 goat anti-mouse antibody) and wash wells three times with 1× PBS.
Add 50 µL/well of DAPI at 1:10,000 dilution and incubate for 5 min on ProBlotTM 25 economy rocker at room temperature in the dark.
Wash wells three times with 150 µL of 1× PBS.
Maintain wells in approximately 125 µL of 1× PBS at 4 °C until imaging.
D2. Image acquisition from FFA samples using the Operetta® high content imaging system
Place the microplate with FFA samples into the instrument.
Note: Ensure the light on the instrument turns blue before opening the lid and inserting the microplate.
Select the Setup tab and choose the following parameters:
Plate type: 96 PerkinElmer CellCarrier Ultra
Objective: 20× high NA
Note: Magnification of the objective used affects the imaging of whole wells and the time taken to acquire an image. Choose a high-magnification objective for whole well imaging; however, more time will be needed for acquisition. The automated SV Foci Counter pipeline has been optimized for the analysis of images captured with the 20× high NA objective.
Optical (Opt.) mode: Non-Confocal
Excitation: 50%
Transmission: 0%
Select channels:
Alexa 488:
Time: 20 ms
Height: 0.0 µM
Note: The term "time (ms)" is used to describe the exposure time, which plays a crucial role in determining the brightness of the image. The exposure time may require adjustment due to batch-to-batch variability in staining intensity. Height (µM) refers to the focus height above the plate bottom. The height indicated here is the default setting and may require amendment to achieve a more focused image.
DAPI:
Time: 10 ms
Height: 0.0 µM
Layout selection:
Number of wells: 96
Well:
Number of fields: 97
Overlap: 0%
Stack:
First plane at: 0.0 µM
Number of planes: 0
Distance: 0.0 µM
Last plane at: 0.0 µM
Overall height: 0.0 µM
Image control:
Coloring: highlight
Flatfield correction: None
Select the Run Experiment tab and press start.
D3. Determination of focus-forming units (FFU) by CellProfilerTM (Figure 4)
Figure 4. Characterization of foci formation in the BHK-21 infectious model. (A) The correlation between the number of plaque-forming units (PFU) and the number of foci is presented herewith. A DENV-2 inoculum of 100 PFU was added to a BHK-21 cell monolayer, and a plaque assay and focus-forming assay (FFA) were subsequently conducted. The number of plaques and foci was determined through manual counting. The number of foci was determined by employing a minimum of one, two, four, or eight infected cells per foci. As illustrated in the bar diagram (dotted lines), the foci identified with a minimum of eight cells demonstrated a stronger correlation with PFUs. In some cases, larger foci may consist of cells that are more than eight in number and may appear to be fusing together. In such instances, the foci will continue to be regarded as two discrete entities. (B) Images of foci were captured using the Operetta® high-content imaging system, and the automated foci counting process was conducted using the SV foci counter, a pipeline designed using the CellProfilerTM software. A foci plot was generated, and the number of foci was counted automatically using the SV foci counter imaging pipeline, which estimated the size of the foci to be eight cells. n = number of foci. (C) Representative images of DENV-1–4 individual foci using 20× objective. The presence of a dotted circle indicates the focus for DENV-1–4, with the number of instances quantified as follows: 3, 3, 2, and 5. Red: E Protein; green: f-actin; blue: nuclei. Scale bar: 300 µm. Data is representative of two technical replicates (n = 2).
Note: See supplementary materials for an example customized image analysis pipeline termed “SV Foci Counter.”
Load images in CellProfilerTM and categorize images into DAPI- and Alexa FluorTM 488-labeled images as Nuclei and Foci under NamesAndTypes function.
Add the following modules and customize the respective settings:
CropGreen and CropBlue
Cropping shape: Rectangle
Cropping method: Coordinates
Apply which cycle’s cropping pattern: Every
Left and right rectangle positions: 0 - end – Absolute
Top and bottom rectangle positions: 0 - end – Absolute
Remove empty rows and columns: All
Identify Primary Objects:
Use advanced settings: Yes
Select the input image and name the primary objects to be identified: Foci
Typical diameter of objects in pixel units (Min, Max): 10, 1,000
Note: Diameter of foci was estimated based on 8 cells minimum per foci.
Discard objects outside the diameter range: Yes
Discard objects touching the border of the image: No
Threshold strategy: Global
Thresholding method: Manual
Manual threshold: 0.1–0.16
Note: Depending on the quality of antibody staining done, there may be a need to adjust the threshold correspondingly.
Threshold smoothing scale: 1.3488
Method to distinguish clumped objects: Intensity
Method to draw dividing lines between clumped objects: Shape
Automatically calculate the size of the smoothing filter for declumping: Yes
Automatically calculate the minimum allowed distance between local maxima: Yes
Speed up by using a lower-resolution image to find local maxima: Yes
Display accepted local maxima: No
Fill holes in identified objects: After both thresholding and declumping
Handling of objects if an excessive number of objects identified: Continue
Export to spreadsheet.
Data analysis
All experiments were conducted with a minimum of two technical replicates and biological replicates. To ensure the accuracy and reliability of the results, an independent experimenter conducted a verification process. The determination of focus-forming units from images captured with the Operetta® high-content imaging system was analyzed using the Harmony program and CellProfilerTM. Dose-response analysis for half maximal effective concentration (EC50) determination can be performed by plotting the dose-response curve using the GraphPad Prism software.
Validation of protocol
Verification of the protocol has been shown in Figures 2–4 in the procedure section. FFA usage in BHK-21 and Vero cells has been shown in Figure 2, section B. Application of the BHK-21 infectious model in evaluating the efficacy of antiviral compound NITD008 against DENV1-4 clinical isolates using FFA is shown in Figure 3, section D. Usage of the Operetta® high-content imaging system and the SV foci counter in determining foci formation is shown in Figure 4, section D3.
Acknowledgments
This research is supported by the IAF-ICP I2301E0019 administrated by Agency for Science, Technology and Research. We would like to acknowledge and give our warmest thanks to all the members of Subhash Vasudevan’s laboratory without whom the protocol established would have not been possible.
Competing interests
Authors declare no competing interests.
References
Bhatt, S., Gething, P. W., Brady, O. J., Messina, J. P., Farlow, A. W., Moyes, C. L., Drake, J. M., Brownstein, J. S., Hoen, A. G., Sankoh, O., et al. (2013). The global distribution and burden of dengue. Nature. 496(7446): 504–507.
Messina, J. P., Brady, O. J., Golding, N., Kraemer, M. U. G., Wint, G. R. W., Ray, S. E., Pigott, D. M., Shearer, F. M., Johnson, K., Earl, L., et al. (2019). The current and future global distribution and population at risk of dengue. Nat Microbiol. 4(9): 1508–1515.
James, M. N. and Dubovi, E. J. (2017). Chapter 29 - Flaviviridae. (p. 525–545). In: Fenner's Veterinary Virology (Fifth Edition). Academic Press.
Bruno, F., Abondio, P., Bruno, R., Ceraudo, L., Paparazzo, E., Citrigno, L., Luiselli, D., Bruni, A. C., Passarino, G., Colao, R., et al. (2023). Alzheimer’s disease as a viral disease: Revisiting the infectious hypothesis. Ageing Res Rev. 91: 102068.
Baer, A. and Kehn-Hall, K. (2014). Viral Concentration Determination Through Plaque Assays: Using Traditional and Novel Overlay Systems. J Visualized Exp.: e52065.
Bolívar-Marin, S., Bosch, I. and Narváez, C. F. (2022). Combination of the Focus-Forming Assay and Digital Automated Imaging Analysis for the Detection of Dengue and Zika Viral Loads in Cultures and Acute Disease. J Trop Med. 2022: 1–11.
Payne, A. F., Binduga-Gajewska, I., Kauffman, E. B. and Kramer, L. D. (2006). Quantitation of flaviviruses by fluorescent focus assay. J Virol Methods. 134: 183–189.
Low, J. G., Ooi, E. E., Tolfvenstam, T., Leo, Y. S., Hibberd, M. L., Ng, L. C., Lai, Y. L., Yap, G. S., Li, C. S., Vasudevan, S. G., et al. (2006). Early Dengue Infection and Outcome Study (EDEN) – Study Design and Preliminary Findings. Ann Acad Med Singap. 35(11): 783–789.
Yin, Z., Chen, Y. L., Schul, W., Wang, Q. Y., Gu, F., Duraiswamy, J., Kondreddi, R. R., Niyomrattanakit, P., Lakshminarayana, S. B., Goh, A., et al. (2009). An adenosine nucleoside inhibitor of dengue virus. Proc Natl Acad Sci USA. 106(48): 20435–20439.
Touret, F., Baronti, C., Goethals, O., Van Loock, M., de Lamballerie, X. and Querat, G. (2019). Phylogenetically based establishment of a dengue virus panel, representing all available genotypes, as a tool in dengue drug discovery. Antiviral Res. 168: 109–113.
Supplementary information
The following supporting information can be downloaded here:
SV foci counter
Article Information
Publication history
Received: May 6, 2024
Accepted: Aug 21, 2024
Available online: Sep 29, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Microbiology > Antimicrobial assay > Antiviral assay
Cell Biology > Cell imaging > Fluorescence
Drug Discovery > Drug Screening > Anti-infective agents
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Expansion and Precise CRISPR-Cas9 Gene Repair of Autologous T-Memory Stem Cells from Patients with T-Cell Immunodeficiencies
XL Xun Li
VC Van Trung Chu
CK Christine Kocks
KR Klaus Rajewsky
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5085 Views: 521
Reviewed by: Alka MehraScott McCombSalma Merchant
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Original Research Article:
The authors used this protocol in Science Immunology Feb 2024
Abstract
The adoptive transfer of autologous, long-lived, gene-repaired T cells is a promising way to treat inherited T-cell immunodeficiencies. However, adoptive T-cell therapies require a large number of T cells to be manipulated and infused back into the patient. This poses a challenge in primary immunodeficiencies that manifest early in childhood and where only small volumes of blood samples may be available. Our protocol describes the ex vivo expansion of potentially long-lived human T memory stem cells (TSCM), starting from a limited number of peripheral blood mononuclear cells (PBMCs). Using the perforin gene as an example, we provide detailed instructions for precise gene repair of human T cells and the expansion of TSCM. The efficiency of precise gene repair can be increased by suppressing unintended non-homologous end-joining (NHEJ) events. Our protocol yields edited T-cell populations that are ready for phenotyping, genome-wide off-target analysis, and functional characterization.
Key features
• Expansion and enrichment of TSCM from PBMCs using IL-7 and IL-15.
• Phenotyping of TSCM.
• Design of “off-the-shelf” gene-repair strategies based on knock-in of a single exon or complete cDNA and design of effective guide RNAs and DNA donor templates.
• High-efficiency gene targeting using CRISPR-Cas9, recombinant adeno-associated virus serotype 6 (rAAV6), and a selective small molecule inhibitor of DNA-dependent protein kinase (DNA-PK).
Keywords: Humans Memory T cells Immunodeficiencies Gene therapy Perforin CRISPR-Cas9 systems Ribonucleoproteins Adeno-associated virus 6 (AAV6) Genome editing Gene editing efficiency
Graphical overview
Background
Familial hemophagocytic lymphohistiocytosis (FHL) is a group of inherited, potentially fatal inflammatory diseases caused by loss-of-function mutations in the NK-cell and T-cell cytotoxic machinery. Patients with FHL2 carry mutations in the perforin-1 gene (PRF1). This protocol is an extension of our previous work with primary, perforin-deficient T cells from a genetic mouse model that mimics FHL2 [1,2]. We isolated memory T cells from these mice, repaired their perforin genes, and showed that treatment with these repaired T cells was able to overcome lethal hyperinflammation [2]. The repaired T cells efficiently killed EBV-triggered malignant B cells in vivo, thereby removing the inflammatory stimulus and restoring immune homeostasis [2].
In the same study, we also showed that human T cells can be repaired in a similar way, setting the stage for translation to the clinic [2]. Ideally, T-cell therapy is performed with antigen-experienced memory T cells that are able to differentiate in vivo into different effector subsets, are long-lived, and possess self-renewing capacities [3]. T memory stem cells (TSCM) reside in CD8+ and CD4+ T-cell precursor subsets that express CD62L [4]. For the CD8 T-cell compartment, it has been shown that CD62L+ cells possess the capacity for immune reconstitution, self-renewal, and persistence in vivo [5]. A combination of recombinant IL-7 and IL-15 can be used to generate such TSCM by activating and expanding a pool of precursor T cells present in blood [6–8]. IL-7 is required for the development of these cells, IL-15 maintains their proliferation, and their gene expression signature resembles naturally occurring TSCM [6].
This protocol describes a highly efficient gene repair strategy for human primary T cells, starting from the typically limited number of peripheral blood mononuclear cells (PBMCs) that can be obtained from critically ill, immunodeficient children in a clinical setting. Ex vivo expansion and enrichment of potentially long-lived TSCM cells is followed by nucleofection of ribonucleoprotein complexes (RNPs) [9] composed of guide RNA (gRNA) and Cas9 nuclease protein to introduce site-specific DNA double-strand breaks. The T cells are then infected with recombinant adenovirus-associated virus serotype 6 (rAAV6) to deliver the DNA template for host cell–mediated homology-directed repair (HDR) [10] and are treated with a small molecule non-homologous end-joining (NHEJ) inhibitor to increase HDR efficiency [8,11]. This strategy achieves highly efficient gene repair in T cells from patients with FHL, resulting in CD8+ T-cell populations carrying more than 85% mCherry reporter knock-ins (see Suppl. Figure S2 in Li et al. [2]).
For effective gene repair, it is desirable to develop “off-the-shelf” strategies that repair as many known disease mutations as possible. We illustrate this principle using the human PRF1 locus, whose exons 2 and 3 encode a 555 amino acid polypeptide [12,13]. By replacing a mutated exon 3 with a corrected version, approximately two-thirds of ~60 known PRF1 disease mutations can be repaired [13]. By replacing exon 3 with the full-length coding region of PRF1, 20 additional disease mutations in exon 2 can be repaired. In both cases, the genetic makeup of the repaired T cells is only minimally altered, and the repaired PRF1 gene is expressed under its physiological transcription and translation control elements.
PBMC samples from FHL2 patients are difficult to obtain and therefore precious, and contain only a limited number of cells. Therefore, we also provide a protocol for cryopreservation of expanded patient TSCM cells. Our protocol does not require sorting steps for T-cell isolation and can be performed repeatedly if needed, allowing optimization of gene editing conditions, scaling up of T-cell culture, and potentially multiple rounds of patient treatment. Expanded patient T cells showed no signs of T-cell exhaustion [14–16], neither before nor after gene editing.
Materials and reagents
Biological materials
Peripheral blood mononuclear cells (PBMCs) from FHL patients and healthy donors (controls)
Reagents
OPTIONAL, see step A1 note: Addgene plasmid #64324 [pU6-(BbsI) CBh-Cas9-T2A-mCherry]
OPTIONAL, see step A1 and A2 notes: Addgene plasmid #86457 [MSCV-hU6-(BbsI)-CcdB-(BbsI)-PGK-Puro-T2A-BFP]
FuGENE® HD transfection reagent (Promega, catalog number: E2311)
Opti-MEMTM serum-reduced medium (Fisher Scientific, catalog number: 31985062)
QuickExtractTM DNA extraction solution (Lucigen, catalog number: QE0905T)
Alt-R® CRISPR-Cas9 crRNA 2 nmol [Integrated DNA Technologies (IDT)] (see Table 1 and Figure S1)
Table 1. Guide RNA (gRNA) sequences for editing the coding part of exon 3 of the PRF1 gene [gene ID 5551; Homo sapiens chromosome 10, GRCh38.14, NC_000010.11(70597348..70602741, complement]. Only the 20 target-specific nucleotides are shown in the third column (see also Figure S1). The preferred protospacer-adjacent motif (PAM) for Cas 9 is NGG, shown in bold and underlined.
Species Name Sequence (5'→3') Genome_coordinates[strand]
Human gPRF1.1 GCGGGGGAGTGTGTACCACATGG Chr10:70599156:70599178[+]
Human gPRF1.2 GGAGCTGGGTGGCCGCATATCGG Chr10:70599018:70599040[-]
Alt-R® CRISPR-Cas9 tracrRNA 100 nmol (IDT, catalog number: 1072534) (see Figure S1)
Alt-RTMS.p. Cas9 nuclease V3 or Alt-RTMS.p. HiFi Cas9 nuclease V3 (IDT, catalog number: 1081059 or 1081061)
De novo synthesized codon-modified DNA repair template (GeneArt Gene Synthesis Services, Thermo Fisher Scientific)
pAAV-GFP Control Vector (Cell BioLabs, catalog number: VPK-400)
NEBuilder® HiFi DNA Assembly Cloning Kit (New England BioLabs, catalog number: E5520S)
One ShotTM Stbl3TM chemically competent E. coliTM (Thermo Fisher Scientific, catalog number: C737303)
ImmunoCultTM-XF T cell expansion medium (STEMCELL Technologies, catalog number: 10981)
Human recombinant IL-7 (5 ng/mL) (PeproTech, catalog number: AF-200-07)
Human recombinant IL-15 (5 ng/mL) (PeproTech, catalog number: AF-200-15)
ImmunoCultTM human CD3/CD28/CD2 T-cell activator (STEMCELL Technologies, catalog number: 10970)
CryoStor® CS10 cryopreservation medium with 10% DMSO (STEMCELL Technologies, catalog number: 07930)
Phosphate-buffered saline (PBS) pH 7.2, without calcium/magnesium (Life Technologies, catalog number: 20012-068)
Bovine serum albumin fraction V (BSA) (Carl Roth, catalog number: 8076.3)
EDTA (Ethylendiaminetetraacetic acid disodium salt) solution pH 8 (0.5 M) (Sigma-Aldrich, catalog number: 03690)
LIVE/DEADTM Fixable Near IR (876) Viability Kit (Thermo Fisher Scientific, catalog number: L34981)
Human recombinant IL-2 (25 ng/mL) (PeproTech, catalog number: 200-02)
P3 Primary Cell 4D-NucleofectorTM X kit L (Lonza, catalog number: V4XP-3012)
Dimethyl sulfoxide (DMSO) 10 mL (Sigma-Aldrich, catalog number: D2438)
M3814 (Nedisertib; DNA-PK inhibitor; CAS number: 1637542-33-6) (MedChemExpress, catalog number: HY-101570)
Ethanol absolute without additives, BioUltra 500 mL (Sigma-Aldrich, catalog number: 51976)
Phorbol-12-myristate-13-acetate (PMA) 1 mg (Sigma-Aldrich, catalog number: P8139)
Ionomycin calcium salt from S. conglobatus 1 mg (Sigma-Aldrich, catalog number: I0634)
Brefeldin A solution (1,000×) (BioLegend, catalog number: 420601)
BD Cytofix/CytopermTM Fixation/Permeabilization Kit (BD Biosciences, catalog number: 554714)
QuickExtractTM DNA extraction solution (LGC Biosearch Technologies/Lucigen, catalog number: QE09050)
PrimeSTAR® GXL DNA polymerase (TaKaRa, catalog number: R050B)
CloneJET PCR cloning kit (Thermo Fisher Scientific, catalog number: K1231)
Antibodies:
Brilliant Violet 785TM anti-human CD8, clone SKI (BioLegend, catalog number: 344740)
APC anti-human CD62L, clone DREG-56 (BioLegend, catalog number: 304810)
PE/Cyanine7 anti-human CD45RA, clone HI-100 (BioLegend, catalog number: 304126)
FITC anti-human CD95 (Fas), clone DX2 (BioLegend, catalog number: 305606)
Brilliant Violet 421TM anti-human CD279 (PD-1), clone NAT105 (BioLegend, catalog number: 367422)
PE/Dazzle TM 594 anti-human TIGIT (VSTM3), clone A15153G (BioLegend, catalog number: 372716)
PE anti-human Perforin, clone B-D48 (BioLegend, catalog number: 353304)
Optional, see step C15: PE anti-human CD366 (Tim-3), clone F38-2E2 (BioLegend, catalog number: 345006)
Optional, see step C15: Brilliant Violet 711TM anti-human CD223 (LAG-3), clone 11C3C65 (BioLegend, catalog number: 369319)
Optional, see step C15: APC/Cyanine7 anti-human CD39, clone A1 (BioLegend, catalog number: 328226)
Solutions
Bovine serum albumin (BSA) stock solution (10%) (see Recipes)
MACS buffer (see Recipes)
P3 electroporation buffer (see Recipes)
M3814 (Nedisertib) stock solution (25 mM) (see Recipes)
Phorbol myristate acetate (PMA) stock solution (50 ng/mL) (see Recipes)
Ionomycin stock solution (500 ng/mL) (see Recipes)
Recipes
Bovine serum albumin (BSA) stock solution (10%)
Layer 10 g of BSA over 100 mL of PBS in a glass beaker and let it dissolve overnight at 4 °C. Sterile-filter through a 0.2 µm PES bottle-top filter. Store at 4 °C or freeze in aliquots at -20 °C.
MACS buffer (PBS pH 7.2 containing 0.5% BSA and 2 mM EDTA)
To 500 mL of PBS pH 7.2 (without calcium and magnesium), add 25 mL of sterile 10% BSA stock solution and 2 mL of 0.5 M EDTA solution pH 8. Sterile-filter through a 0.2 µm PES bottle-top filter.
CRITICAL: LIVE/DEADTM fixable stains are only compatible with flow cytometry buffers containing less than 1% of protein.
P3 electroporation buffer
Just before use, freshly prepare 20 µL of P3 electroporation buffer by adding 3.6 µL of Supplement 1 to 16.4 µL of P3 primary cell NucleofectorTM solution (P3 Primary Cell 4D-NucleofectorTM X kit, Lonza).
M3814 (Nedisertib) stock solution (25 mM)
Dissolve 5 mg of M3814 in 415 µL of DMSO (25 mM = 5,000×). Store in 10 µL aliquots at -80 °C. Dilute the stock solution 1:100 in PBS pH 7.2 to make a 50× working stock solution (250 µM = 50×). Aliquot the working stock solution and store at -20 °C for up to 4 weeks.
Phorbol myristate acetate (PMA) stock solution (50 ng/mL)
Dissolve 1 mg of PMA in 20 mL of ethanol (50 ng/mL final concentration). Store in 10 µL aliquots at -20 °C.
CAUTION: Phorbol ester is a tumor promoter. Wear personal protective equipment including gloves.
Ionomycin stock solution (500 ng/mL)
Dissolve 1 mg of ionomycin calcium salt in 2 mL of ethanol (500 ng/mL final concentration). Store in 10 µL aliquots at -20 °C.
Laboratory supplies
Tissue culture 96-well plates, flat bottom (Sarstedt, catalog number: 83.3924)
Tissue culture 96-well plates, U bottom (Sarstedt, catalog number: 83.3926.500)
Tissue culture 48-well plates (Sarstedt, catalog number: 83.3923)
Tissue culture 24-well plates (Sarstedt, catalog number: 83.3922)
Tissue culture 12-well plates (Sarstedt, catalog number: 83.3921)
Tissue culture 6-well plates (Sarstedt, catalog number: 83.3920)
Tissue culture dish 100 × 20 mm (Sarstedt, catalog number: 83.3902)
PIPETMAN® single-channel pipettes P10, P20, P200, P1000 (Gilson)
Pipette filter tips [Sarstedt, catalog numbers: 70.3010.275 (10 µL), 70.3030.265 (20 µL), 70.3030.375 (100 µL), 70.3030.110 (200 µL), 70.3060.275 (1,000 µL)]
0.2 mL PCR tubes with domed lids, 8-well strips (neoLab, catalog number: 7-5207)
1.5 mL tubes (Eppendorf, catalog number: 0030 120.086)
2.0 mL tubes (Eppendorf, catalog number: 0030 120.094)
15 mL conical centrifugation tubes FalconTM 352097 (Thermo Fisher, catalog number: 10136120)
Mr. FrostyTM freezing container (Thermo Fisher, catalog number: 5100-0036)
2 mL CryoPure tubes (freezing vials) (Sarstedt, catalog number: 72.380.992)
Disposable PES filter units 0.2 µm, 150 mL (Thermo Scientific Nalgene, catalog number: 596-3320)
Disposable PES filter units 0.2 µm, 500 mL (Thermo Scientific Nalgene, catalog number: 595-3320)
Standard 96-well microtiter plate, U bottom (Thermo Scientific Abgene, catalog number: AB0796)
Equipment
Tissue culture incubator set at 37 °C, 5% CO2
Refrigerated centrifuge with adapters for plates and 15 mL conical tubes (e.g., Eppendorf Centrifuge 5810R)
Benchtop centrifuge for 1.5 mL/2 mL tubes (e.g., Eppendorf Centrifuge 5424)
BD LSRFortessa cell analyzer flow cytometer (BD Bioscience)
TC20TM automated cell counter (Bio-Rad, catalog number: 1450102)
4D-Nucleofector® X Unit (Lonza, catalog number: AAF-1003B)
Bright-LineTM hematocytometer (Sigma-Aldrich, catalog number: Z359629)
Optional; see steps A2 and A3: BD FACSAria III Cell Sorter (BD Biosciences)
Software and datasets
CrispRGold (free) (https://crisprgold.mdc-berlin.de/) or CRISPOR (free) (http://crispor.tefor.net/)
ICE CRISPR Analysis Tool (Synthego) (free)
https://www.synthego.com/products/bioinformatics/crispr-analysis
GeneOptimizer Algorithm (GeneArt Gene Synthesis Services, Thermo Fisher Scientific) (free)
https://www.thermofisher.com/de/de/home/life-science/cloning/gene-synthesis/geneoptimizer.htmL
FlowJo (v.10, FlowJo, BD Biosciences) (commercial analysis software, requires a license)
ApE (A Plasmid Editor) (free) (https://jorgensen.biology.utah.edu/wayned/ape/)
Procedure
Design and experimental assessment of gRNA efficiency
Note: gRNAs contain the target-specific sequence for guiding Cas9 protein to a genomic location. We use a two-part guide RNA annealed to form an overlapping duplex (Figure S1). One part of this two-part gRNA consists of a non-variable 67-nucleotide-long tracrRNA; the other consists of a partly variable 36-nucleotide-long crRNA that harbors 20 target-specific nucleotides. Alternatively, you can use a format that combines these RNAs in a 100-nucleotide-long single-guide RNA. We prefer the former option because it reduces costs. See General note 1.
Design gRNAs with CrispRGold (https://crisprgold.mdc-berlin.de/) [17] around the selected genomic target site [coding part of exon 3 of the PRF1 gene (Gene ID 5551; Homo sapiens chromosome 10, GRCh38.14, NC_000010.11[70597348..70602741, complement)]. Alternatively, gRNAs can be designed with CRISPOR (http://crispor.tefor.net/) [18]. The software outputs the 20 target-specific nucleotides in the gRNA. For examples of gRNAs targeting exon 3 of the PRF1 gene, see Table 1, Figure 1, and Figures S2–S4.
Note: To reduce costs, we often pre-select efficient gRNAs by first ordering oligo duplexes, cloning them into mammalian expression vectors, and transfecting them into cells [19]. Alternatively, to save time, you can skip step A2 and order several crRNAs right away from Integrated DNA Technologies (IDT), assemble the corresponding RNPs (steps D1–D5), electroporate them into activated primary human T cells (steps F1–F12), and then proceed as described in steps A3–A5 below.
Order forward and reverse DNA oligos [with cloning-compatible overhangs as configured by CrisprGold (Figure S3) or CRISPOR] corresponding to the 5–6 most high-ranking gRNAs (specificity score ≥ 12 in CrispRGold; Figure S3), phosphorylate, anneal, and clone them via BbsI sites (Figure S5) into a guide RNA/Cas9 expression plasmid, and transfect into HEK293T cells. We use the plasmid pU6-(BbsI) CBh-Cas9-T2A-mCherry (Addgene plasmid #64324, https://www.addgene.org/64324/; Chu et al. [20]). It was modified by us to express the fluorescent marker mCherry and is originally based on Addgene plasmid #42230, https://www.addgene.org/42230/; Cong et al. [21], Ran et al. [22]). For transfection, we use 5 × 105 HEK293T cells in a 6-well plate and transfect them with 9 µL of FUGENE® HD transfection reagent with 3 µg of sgRNA-expression plasmid (ratio 3:1) completed with Opti-MEMTM to a final volume of 150 µL. We routinely obtain a transfection efficiency of >80%.
Note: In order to be able to select for transfected cells by fluorescence-activated cell sorting (FACS), we often use the expression plasmid MSCV-hU6-(BbsI)-CcdB-(BbsI)-PGK-Puro-T2A-BFP carrying a blue fluorescent protein (BFP) reporter (Addgene Plasmid #86457, https://www.addgene.org/86457/; Chu et al. [19]) and transfect into Cas9-expressing suspension cells, such as BJAB lymphoma cells by electroporation. (Any human Cas9-expressing cell line can be used for this purpose. A large selection of Cas9-expressing cell lines is commercially available.)
Three days after transfection, isolate genomic DNA from the transfected cells [along with unedited (non-transfected) control cells] (50 µL of QuickExtractTM DNA Extraction Solution per 105 cells) and PCR-amplify a 500–1,000-bp region around the targeted genome site. Send the PCR amplicons to Sanger sequencing.
Note: Alternatively, one day after transfection, first isolate BFP-expressing (transfected) cells by FACS and expand the sorted cells for two days, before isolating genomic DNA. gRNA action through Cas9 RNP delivery reaches a maximum after approximately 24 h (Kim et al. [23], see Figure 6B).
Assess the genome editing efficiency of your guide RNAs by Inference of CRISPR Edits (ICE) [24]. Upload Sanger trace data (.ab1 files) to the free ICE CRISPR Analysis Tool (https://www.synthego.com/products/bioinformatics/crispr-analysis) (Figure S6).
CRITICAL: High-quality Sanger sequencing data are required for this step. See also General note 1.
Order the two highest-scoring crRNAs, as well as tracrRNA, and S.p. Cas9 or S.p. HiFi Cas9 from a commercial supplier such as IDT. (For applications where high fidelity of gene repair is critical, we recommend HiFi Cas9.)
CRITICAL: Good gRNAs are extremely important for achieving efficient and reliable gene editing. The higher the predicted knockout efficiency, the better. Generally, we aim for an ICE indel percentage of 60% or higher (Figure S6).
Design of DNA donor templates
Gene correction strategy examples for the PRF1 gene are shown in Figure 1.
Figure 1. Examples of two gene correction strategies for the PRF1 gene. A. Schematic representation of the human PRF1 gene and the encoded 555 amino acid perforin polypeptide. UTR, untranslated region; SP, signal peptide (cleaved off after translocation of the nascent protein into the ER); MACPF, membrane attack complex/perforin domain (includes the central machinery of pore formation); EGF, EGF-like domain (forms a shelf-like assembly connecting MACPF and C2 domain); C2, C2 domain (calcium-dependent phospholipid binding). B. Two gene correction strategies. With both strategies, the native exon 3 coding part gets replaced by a repaired version whose sequence has been diversified (codon-modified, see Procedure step B2). Repair templates are codon-modified to prevent unwanted HDR events that do not lead to gene repair. Strategy 1 replaces the mutated version of exon 3 with a repaired version. For a more comprehensive gene correction by cDNA knock-in [25], strategy 2 replaces exon 3 with the full-length PRF1 coding sequence (CDS): In place of the 5' UTR, we engineered the cDNA to start with a T2A “self-cleaving peptide” preceded by a flexible serine-glycine-linker (TCC.GGC.AGC.GGC) [26], that is followed by the PRF1 ATG start codon, the signal peptide, and the complete PRF1 coding sequence ending with the TGA stop codon. Strategy 2 places the PRF1 CDS under endogenous transcriptional and translational control and allows correction of the ~60 known pathogenic PRF1 mutations (with the exception of frameshift or nonsense mutations in exon 2) [12,13,27]. Cleavage sites for the two guide RNAs (gRNA) are indicated. We use two gRNAs for gene repair in primary T cells because this increases the gene repair efficiency (see Li et al. [2] Suppl. Table S1). HA, homology arm; HDR; homology-directed repair. Adapted from Figure 6 in Li et al. [2].
Design a DNA donor template carrying 5' and 3' homology arms of at least 0.5–1.5 kb upstream and downstream of the coding part of exon 3 of PRF1 (Figure 1). Design primers and PCR-amplify homology arms from the genomic DNA from a healthy donor, incorporating 20–30 bases overlap to exon 3 (or the T2A-cDNA) and the NotI-linearized backbone vector (Figure S7). These overlaps are required for the “one-step cloning” as described below in step B4.
Note: Generally, we aim for a minimum length of 500 bp but up to 1 kb or more if the packaging capacity of the adeno-associated virus (AAV) vector allows it (maximally 4–4.4 kb insert size excluding inverted terminal repeats;https://www.vectorbiolabs.com/faq-aav/). See also Figure S7.
Modify the nucleotide sequence of the replaced exon or the cDNA in the DNA repair template by using the free Gene Optimizer Algorithm (GeneArt Gene Synthesis Services, Thermo Fisher Scientific) [28]. We refer to this modified exon 3 repair template as “codon-modified” (see Figure 1).
CRITICAL: This step is intended to retain high expression of the repaired gene while diversifying the nucleotide sequence to prevent unwanted, small HDR events that would not lead to gene repair. These can occur in DNA stretches with extended homologies between the repair template and the native genomic locus.
Order a de novo synthesized DNA fragment corresponding to the DNA repair template.
Assemble the whole HDR template for “one-step cloning” into the AAV vector: Mix NEBuilder® HiFi DNA Assembly Master Mix with 0.5 pmole each of the 5' and 3' homology arms, the codon-modified exon 3 (or T2A-cDNA), and the pAAV-GFP backbone vector (linearized with NotI) (Figure S7), and incubate the reaction at 50 °C for 60 min.
Transform into chemically competent E. coli such as the HB101-derived strain Stbl3, which supports cloning of inserts containing repetitive elements, and purify the plasmids containing the HDR repair templates. (Owing to the high efficiency of NEBuilder® HiFi DNA Assembly, we usually obtain approximately 30% of colonies with inserts at this step.)
Note: Alternatively, you can assemble the HDR template by one-step cloning into a NotI linearized intermediate cloning vector, and only clone the assembled NotI fragment into pAAV in a second step. We also deposited cloning plasmids containing the assembled HDR templates for PRF1 targeting (NotI-fragment) (Addgene plasmids # 209075 and 209076).
Confirm the sequence of the HDR repair templates by Sanger sequencing.
CRITICAL: The viral vector carries an inverted terminal repeat (ITR) at each end. These sequences are repetitive and make cloning difficult and the pAAV plasmids unstable. Since this can affect AAV production and targeting efficiency downstream, we recommend submitting the plasmid to full plasmid next-generation sequencing to a commercial supplier such as Genewiz or Eurofins Genomics LLC.
Expansion and enrichment of T memory stem cells from peripheral blood mononuclear cells (PBMCs) using IL-7 and IL-15
A general workflow for the expansion of T memory stem cells is shown in Figure 2.
Figure 2. Workflow for expanding and enriching T memory stem cells (TSCM) from peripheral blood mononuclear cells (PBMCs). Starting from frozen aliquots of PBMCs, cells are cultured in bulk and the T cells (including all T-cell subsets) are selectively activated in the presence of IL-7 and IL-15 for 3 days, then further expanded in the presence of IL-7 and IL-15. IL-7 is required for the development of TSCM and IL-15 maintains their proliferation without promoting their differentiation into effector subsets [6]. Cells are then frozen down for later use.
Thaw a vial of frozen PBMCs (from a patient with FHL2 or a healthy donor) and transfer cells to 5 mL of prewarmed Immunocult XF T-cell expansion medium in a 15 mL conical tube. Isolation and cryopreservation of PBMCs is a routine procedure in clinical settings; see Puleo et al. [29] for more details.
Activate bulk T cells (T cells containing all subsets): Pellet cells at 300× g for 5 min at 4 °C, gently remove supernatant, and resuspend the cell pellet in ImmunoCultTM-XF T cell expansion medium with human IL-7 (5 ng/mL) and IL-15 (5 ng/mL) in the presence of ImmunoCultTM human CD3/CD28/CD2 T-cell activator (25 μL/mL) at a density of 1 × 106/mL in a well of a 12- or 6-well plate (for up to 2 or 4 mL, respectively). Incubate cells at 37 °C and 5% CO2 for approximately 72 h.
Collect activated bulk T cells and pellet cells at 300× g for 5 min at 4 °C, gently removing the supernatant.
Resuspend cells with fresh ImmunoCultTM-XF T cell expansion medium with human IL-7 (5 ng/mL) and IL-15 (5 ng/mL) at a density of approximately 0.5 × 106 cells/mL in wells of a 6-well plate (up to 4 mL) or 100 mm culture dishes (up to 10 mL).
Incubate for another 5–7 days. During this period, change the medium and split cells to a density of approximately 0.5 × 106/mL every 2–3 days, as needed. We recommend a maximum density of approximately 1 × 106 cells per milliliter. Approximately 95% of the cells are CD3+ T cells at this point, half of which are CD8+ T cells that have expanded 600–700-fold, as estimated from two independent experiments. Within the CD8+ T-cell compartment, most of the cells were TSCM cells; see step C15 and Figure 3.
Figure 3. Phenotype of CD8+ T memory stem cells (TSCM) from a patient with familial hemophagocytic lymphohistiocytosis (FHL) after selective expansion and enrichment from peripheral blood monocytic cells (PBMCs). A. Flow cytometry analysis of CD8+ T cells before (Day 0) and after expansion (Day 10) from PBMCs isolated from a FHL patient. CD8+ T cells expanded 600–700-fold, starting from 104 to 4 × 104 cells (n = 2). Expanded CD8+ T cells stained positive for the naïve T-cell markers CD45RA and CD62L, but were almost exclusively CD95+, distinguishing them from CD45R+CD62L+CD95- naïve T cells and identifying them as TSCM-like. B. The expanded CD8+ T cells did not express the T-cell exhaustion markers PD-1 and TIGIT compared with the control staining. Thus, expanded CD8+ T cells from FHL patients showed no signs of T-cell exhaustion, an important prerequisite for T-cell therapy. TEMRA, effector memory T cells re-expressing CD45RA; TEM, effector memory T cells; TN, naïve T cells; TSCM, T memory stem cells; TCM, central memory T cells.
Prepare the expanded patient T cells for cryo storage: Reserve some cells for flow cytometry analysis (approximately 2–5 × 105 cells in a 96-well U-bottom plate; enough for two to maximum five staining reactions), pellet the remaining cells at 300× g for 5 min at 4 °C, gently remove supernatant, resuspend the cell pellet in ice-cold CryoStor® CS10 solution, and transfer the cell suspension to cryovials (5 × 105 to 1 × 106 cells in 0.5 mL per vial).
Cryopreserve cells using a Mr. FrostyTM freezing container, store them at -80 °C overnight, and transfer them to a liquid nitrogen tank on the next day.
Prepare the cells for flow cytometry analysis: Wash the reserved cells (from step C6) once with 100 μL of MACS buffer and pellet at 300× g for 5 min at 4 °C; then, carefully remove the supernatant without disturbing the cell pellet.
Reconstitute a vial of the lyophilized fluorescent, amine-reactive LIVE/DEADTM dye with DMSO in 500 µL (instead of the 50 µL volume recommended by the manufacturer). Add 0.2 μL of the reconstituted LIVE/DEADTM dye to 50 μL of MACS buffer per sample (1:250).
Note: You can store the remaining reconstituted LIVE/DEADTM dye in aliquots at -20 °C.
Resuspend cell pellet with 50 μL of LIVE/DEADTM dye containing MACS buffer and incubate at 4 °C for 10 min, protected from light.
Prepare the staining solution by mixing the following antibodies in 50 µL of MACS buffer per sample: BV785-CD8 (diluted 1:200), APC-CD62L (1:400), PE/Cy7-CD45RA (1:400), FITC-CD95 (1:400), BV421-PD1 (1:50), and PE/Dazzle594-TIGIT (1:100). (CD62L, CD45RA and CD95 are markers for TSCM cells, while PD1 and TIGIT serve as markers for exhausted T cells; see Figure 3A and B.)
Wash the cells once with MACS buffer and resuspend in 50 μL of staining solution.
Stain cells with the staining solution for 15 min at 4 °C, protected from light.
Wash cells once with 200 μL of MACS buffer and resuspend in 100 µL of MACS buffer prior to flow cytometric analysis (Figure 3). Keep the cells on ice.
Flow cytometry analysis: Acquire at least 1 × 104 to 2 × 104 events (live cells) by first gating on lymphocytes (using the forward and side scatters FSC-A/SSC-A), then gating on singlet cells (by excluding doublets, clumps, and debris using FSC-H/FSC-W and SSC-H/SSC-W), and then gating on live cells (by excluding dead cells based on the LIVE/DEADTM stain signal). For data analysis, first pre-gate on CD8+ cells and then analyze the other T-cell subsets (see Figure 3).
CRITICAL: Be sure to include a positive control for the antibody staining and the setting of the flow cytometry parameters. We use activated T cells from healthy donors (stimulated two days before flow analysis), and follow the procedure described in step F1 and F2. For clinical applications, we recommend that you characterize your T cells for the expression of additional T-cell exhaustion markers (TIM-3, LAG-3, and CD39) (see Suppl. Figure S5 in Li et al. [2]).
Prepare RNP complexes
Generate gRNA complexes by mixing 20 μL of a given crRNA (100 pmol) with 20 μL of tracrRNA (100 pmol) at a molarity ratio of 1:1 in a PCR tube.
Heat the mixture at 95 °C for 5 min.
Cool to room temperature (15–25 °C) on the benchtop for at least 10 min.
Store gRNA complexes at -20 °C for up to 1 year.
Right before electroporation, generate RNP complexes by mixing 2 μL of gRNA complex (100 pmol) and 0.75 μL of Cas9 (~50 pmol) at a molarity ratio of 2:1 in a PCR tube per 5 × 105 activated human T cells and incubate the mixture at room temperature for 10–20 min on the benchtop.
Note: We use two gRNAs for gene repair in primary T cells because this increases the gene repair efficiency (Li et al. [2] Suppl. Table S1).
Mix the two RNP complexes immediately before electroporation.
Production and purification of adeno-associated virus (AAV) donor vectors
Transfect, purify, concentrate, and calculate the copy number of rAAV6 donor particles according to our previous protocol [30]. See General note 2.
Genome editing of human T cells and phenotyping of edited T cells by flow cytometry
The general workflow for gene editing of expanded patient-derived T cells is shown in Figure 4.
Figure 4. Workflow for CRISPR/Cas9 gene repair in expanded CD8+ T memory stem cells (TSCM) from patients with familial hemophagocytic lymphohistiocytosis (FHL). Starting from aliquots of expanded, patient-derived frozen TSCM-like cells, T cells are reactivated and expanded for 2 days with IL-2, then electroporated with RNPs, and immediately infected with recombinant adenovirus-associated virus (AAV) serotype 6 (carrying the DNA donor repair templates) in the presence or absence of a small molecule NHEJ inhibitor [DNA-PK inhibitor M3814 (Nedisertib)]. Transfected, infected T cells are first expanded for 4 days with IL-2 and then for two more days in the presence of IL-7 and IL-15.
Thaw frozen vials from step C7 and culture T cells in ImmunoCultTM-XF T cell expansion medium with human IL-2 (25 ng/mL) in the presence of ImmunoCultTM human CD3/CD28/CD2 T-cell activator (25 μL/mL) at a density of 1 × 106/mL.
Forty-eight hours post reactivation, harvest expanded T cells and count the cells using a hemacytometer.
Prewarm ImmunoCultTM-XF T cell expansion medium (at least 1 mL per sample).
Transfer 5 × 105 T cells to a 1.5 mL Eppendorf tube and spin down at 300× g for 5 min at room temperature.
Note: We generally electroporate 5 × 105 to 106 activated human T cells per electroporation reaction. The minimum cell number of T cells per electroporation is 3 × 105.
Remove the supernatant and wash the cells once at room temperature with 1 mL of PBS.
Pellet cells at 300× g for 5 min at room temperature and carefully remove the supernatant without disturbing the cell pellet.
Suspend cells in 20 μL of P3 buffer (see Recipes) by gently pipetting up and down 10 times.
Add freshly prepared RNPs (see steps D5 and D6) to the cell suspension and mix well by pipetting up and down 10 times.
Transfer the mixture to a well of a 16-well nucleocuvette strip.
Electroporate the cells using the EH-100 program of Lonza 4D-Nucleofector.
After electroporation, add 75 μL of prewarmed ImmunoCultTM-XF T cell expansion medium containing human IL-2 (25 ng/mL) per well and gently mix cells.
Transfer resuspended cells to an Eppendorf tube containing 500 µL of prewarmed culture medium.
Transfer 1/5 to 2/5 of the cells in 200 μL of prewarmed ImmunoCultTM-XF T cell expansion medium containing human IL-2 (25 ng/mL) to a well of a 96-U bottom plate (corresponding to an estimated 1 × 105 to 2 × 105 cells based on the input).
Add rAAV6 donor particles (from Section E) to the well containing the electroporated cells at a MOI of 1 × 106 to 2 × 106 genome copy/cell.
Add 4 µL of M3814 (Nedisertib) inhibitor from a 50× working stock solution (250 µM; see Recipes) to a final concentration of 5 μM.
After approximately 16 h, gently mix cells by pipetting up and down 10 times and pellet cells at 300× g for 5 min at 4 °C; then, carefully remove the supernatant without disturbing the cell pellet.
Add 200 µL of prewarmed ImmunoCultTM-XF T cell expansion medium containing human IL-2 (25 ng/mL) and culture cells in a 96-well U bottom plate.
After approximately 24 h, transfer cells to a 48-well plate and culture cells in 350–400 μL of medium.
Note: To achieve sufficient expansion of edited human T cells, we recommend gradually expanding cells from a 96-well U-bottom plate to a bigger plate (48-well plate, 24-well plate, 12-well plate, and 6-well plate) and monitoring the culture every day until the cells are used for downstream analysis or application.
Four days post-infection, change the medium to fresh ImmunoCultTM-XF T cell expansion medium with human IL-7 (5 ng/mL) and IL-15 (5 ng/mL) for another two days (1.5 mL for a 12-well plate or 3 mL for a 6-well plate).
Analyze the cells by flow cytometry (stain cells as described in steps C9–C15) (see Figure 5).
Figure 5. Efficient repair of perforin mutations CD8+ T memory stem cells (TSCM) from a patient with familial hemophagocytic lymphohistiocytosis (FHL). A. The efficiency of PRF1 gene repair was assessed by flow cytometry analysis of intracellular perforin expression in CD8+ T cells from a patient with FHL. T cells were reactivated two days before the analysis, then stimulated with PMA and ionomycin for 6 h in the presence of brefeldin A, before fixation, permeabilization, and staining for precursor and mature forms of perforin with monoclonal antibody B-D48 [31]. Left: Representative flow cytometry plots after the indicated treatments. An inactive precursor form of perforin is detectable in the CD8+ T cells of this patient (patient alleles) because the patient had a c.1349C>T(T450M) missense mutation, known to impair the proteolytic maturation of perforin [32]. By contrast, perforin staining is almost absent in CD8+ T cells that received RNP without AAV6 (null alleles, generated by NHEJ events causing frameshift mutations). Right: Frequency of perforin-expressing CD8+ T cells after perforin repair by RNP/AAV6/M3814 treatment. Each dot represents one independent experiment, horizontal lines indicate medians. [Note: In these experiments, we also used human BJAB lymphoma cells and a CD19-CD3 bispecific antibody [33] to activate the patient’s T cells, but we later found (using T cells from healthy donors) that this additional activation mode is not necessary (Li et al. [2], Figure 5D)]. B. Left: Representative phenotypes by flow cytometry of repaired patient-derived CD8+ T cells after the indicated treatments. CD45RA+CD62L+ T cells were almost exclusively CD95+, differentiating them from naïve T cells and identifying them as TSCM-like. Right: Frequency of CD45RA+CD62L+CD8+ T cells after the indicated treatments. Each dot represents one independent experiment, horizontal lines indicate medians. C. Repaired CD8+ T cells did not overexpress the T-cell exhaustion markers PD-1 and TIGIT. Other T-cell exhaustion markers (TIM-3, LAG-3, and CD39) were also tested and found not to be overexpressed (see Suppl. Figure S5 in Li et al. [2]). Frequency of PD1-TIGIT- CD8+ T cells after the indicated treatments. Each dot represents one independent experiment, horizontal lines indicate medians. RNP, ribonucleoprotein complexes of gRNA and Cas9 nuclease; AAV6, adeno-associated virus serotype 6 providing the DNA repair template; M3814, DNA-PK inhibitor M3814 (Nedisertib). Non-targeted cells correspond to untransfected patient cells. Reprinted/adapted from Figure 6 in Li et al. [2].
Assessment of gene-targeting efficiency by intracellular perforin staining
Reactivate 5 × 105 to 1 × 106 edited T cells per well of a 24-well plate with ImmunoCultTM-XF T cell expansion medium with human IL-2 (25 ng/mL) in the presence of ImmunoCultTM human CD3/CD28/CD2 T-cell activator (25 μL/mL) at a density of 1 × 106/mL for two days.
Stimulate 1 × 105 to 2 × 105 reactivated T cells with PMA (50 ng/mL) and ionomycin (500 ng/mL) for 6 h in the presence of 1× Brefeldin A solution in a 96-well U bottom plate.
Pipette several times and pellet cells at 300× g for 5 min at 4 °C in the same 96-well U-bottom plate. Then, carefully remove the supernatant without disturbing the cell pellet.
Wash the cells once with 100 μL of MACS buffer and pellet cells at 300× g for 5 min at 4 °C. Carefully remove the supernatant without disturbing the cell pellet.
Add 0.2 μL of the reconstituted LIVE/DEADTM dye per 50 μL of MACS buffer.
Resuspend cell pellet with 50 μL of LIVE/DEADTM dye containing MACS buffer and incubate at 4 °C for 10 min, protected from light.
Wash the cells once with MACS buffer and resuspend in 50 μL of MACS buffer.
Stain for surface markers as described in step C11 and incubate for 15 min on ice, protected from light.
Wash the cells once with MACS buffer.
Thoroughly resuspend cells in 100 μL of Cytofix/CytopermTM solution for 20 min at 4 °C in the dark.
Prepare the perforin staining solution by diluting 1 µL of anti-perforin antibody in 50 µL of 1× Perm/WashTM solution.
Note: We use the monoclonal antibody B-D48 that detects both precursor and mature forms of perforin (Hersperger et al. [31]; see Li et al. [2] Figure 6D).
Wash cells two times in 1× Perm/WashTM solution, pellet, and remove supernatant.
Thoroughly resuspend fixed/permeabilized cells in 50 μL of 1× Perm/WashTM solution containing anti-perforin antibody. Incubate at 4 °C for 30 min in the dark.
Wash cells two times with 200 μL of 1× Perm/WashTM solution and resuspend in 100 μL of MACS buffer prior to flow cytometry analysis (Figure 5).
Assessment of knock-in efficiencies by on-target PCR, cloning, and sequencing of PRF1 alleles
Note: This step serves to confirm the correct integration site and to quantify the fractions of HDR and NHEJ events. Alternatively to the cloning procedure below, you could do PacBio long-read amplicon sequencing.
Extract genomic DNA from 5 × 105 to 1 × 106 targeted bulk T cells with 500 µL of QuickExtractTM DNA extraction solution (at a density of 1 × 106 to 2 × 106 cells/mL).
Mix well and briefly vortex to completely lyse cells. Transfer 50–100 µL of lysed cells to a PCR tube.
Denature the genomic DNA using a PCR machine and the following temperature cycle: 65 °C for 15 minutes, 68 °C for 15 min, 95 °C for 15 min, and 4 °C. The denatured genomic DNA is ready for PCR amplification.
Amplify the targeted site using PrimeSTAR® GXL DNA polymerase with gene-specific primers that bind outside of the homology arms (HA) (PRF1.5HA-Fw and PRF1.3HA-Rv; Li et al. [2] Suppl. Table S8). PCR conditions: 30 cycles (98 °C for 10 s, 60 °C for 15 s, 68 °C for 1 min per kb) in 25 µL containing 5 µL of 5× PrimeSTAR® GXL buffer, 0.5 µL of PrimeSTAR® GXL polymerase, 2 µL of NTP mixture (2.5 mM each), plasmid template 1 µL (2.5–250 ng), primers 1 µL each (10 µM), filled up to 25 µL with water.
Clone the purified PCR products into sequencing vector pJET1.2/Blunt (CloneJET PCR cloning kit).
Pick approximately 100 colonies into 96-well microtiter plates (MTP) (pre-filled with 200 µL of LB agar containing 100 µg of ampicillin), incubate overnight at 37 °C, and send the MTP with suitable primers to MTP sequencing (Sanger Sequencing Services Premium Run, LGC Biosearch Technologies).
Open sequence files one by one with ApE software [34] and count nontargeted, HDR, and NHEJ events (Table 2 and Suppl. Figure S8).
Note: Using two sgRNAs and NHEJ inhibitor M3814, we observed median allele editing frequencies of approximately 60% with strategy 1 and 40% with strategy 2 (n ≥ 3; see also Li et al. [2] Figure 6B and Suppl. Table S2).
Table 2. Knock-in efficiencies measured by sequencing of targeted alleles. Percentage of HDR, NHEJ, and non-targeted events were calculated as illustrated in Figure S8.
Strategy 1 (n = 4) Strategy 2 (n = 3)
% HDR 61.8 73.0 61.5 61.7 38.2 63.3 41.1
% NHEJ 35.3 27.0 35.9 36.2 58.8 23.3 58.9
% Non-targeted 2.9 0 2.6 2.1 2.9 13.3 0
Genome-wide identification of off-target sites with GUIDE-seq
For Tn5-modified GUIDE-seq experiments and data analysis [35], please follow our previous Bio-protocol (Tran et al. [36]; see Section H. Tn5 transposase-mediated GUIDE-seq). See General note 3.
Data analysis
Genome editing efficiency of gRNAs: See Figure S6 for an example of an “inference of CRISPR edits” (ICE) analysis. Flow cytometry data are analyzed with the commercial software FlowJo. Basic expertise in flow cytometry is required to assess the phenotype of patient T cells before and after gene repair and to quantitatively assess cell yields of edited T cells and the gene repair efficiency by measuring the percentage of repaired T cells expressing perforin in response to T-cell activation.
Phenotyping of T cells: After 10 days of culture with IL-7 and IL-15, more than 90% of CD8+ T cells were TSCM cells without signs of T-cell exhaustion; after gene editing, >75% of the T cells retained this phenotype (Figure 3, Figure 5; see also Li et al. [2] Suppl. Figure S5).
Yield of edited cells: Without NHEJ inhibitor M3814, T cells from healthy donors yielded 8 × 105 to 9 × 105 edited T cells after 6 days (starting from 105 T cells) (see Li et al. [2] Figure 5C). The addition of M3814 led to a 30%–40% reduction in cell yield. See General note 4. Using two gRNAs and NHEJ inhibitor M3814, we observed median gene editing efficiencies of approximately 80% and 70% with strategy 1 and strategy 2, respectively (n = 4; see also Li et al. [2] Figure 6B and Suppl. Table S2).
Knock-in efficiencies by sequencing and percentage of HDR, NHEJ, and non-targeted alleles: These data are given in Table 2. Please see Figure S8 in this Bio-Protocol for how to identify HDR, NHEJ, and non-targeted alleles by sequencing and how to calculate the contribution of each to the total number of alleles after gene editing. We recommend sequencing a minimum of 25–30 alleles to obtain representative results. To make sure gene targeting efficiencies are reproducible, experiments should be repeated at least two times.
Validation of protocol
Validation experiments for both gene editing strategies (Figure 1) are shown in our article Li et al. [2], as follows: PRF1 gene editing conditions were optimized using T cells from healthy donors [Figure 5 and Table S1; one independent experiment per donor (n = 3)]. Efficient repair of perforin mutations in T cells from a patient with FHL2 was demonstrated in Figure 6, Table S1, and Suppl. Figure S4 [four independent experiments (n = 4)]. Patient T cells showed a TSCM-like phenotype after gene repair [Figure 6D; four independent experiments (n = 4)] and did not show increased levels of the T-cell exhaustion markers PD-1, TIGIT, TIM-3, LAG-3, and CD39 [Figure 6E and Suppl. Figure S5; four independent experiments (n = 4)].
General notes and troubleshooting
General notes
Because gRNAs can vary widely in gene editing efficiency, it is important to test this parameter experimentally. There are different methods available (see Tran et al. [36] steps A1–A4). For example, gRNAs can be tested in vitro using synthetic crRNAs (or in vitro–transcribed gRNAs), Cas9 protein, and PCR amplicons. gRNAs can also be tested in cell lines or in primary cells, which is considered more reliable than in vitro testing. Popular assays for quantitative assessment of the efficiency of target site cleavage are the Surveyor/T7 endonuclease I (T7EI) assay [22] or Sanger sequencing followed by ICE analysis [24] or TIDE analysis [37]. Commercial kits for testing gRNA efficiency are available as well, such as the GeneArtTM Genomic Cleavage Detection Kit (Invitrogen), the Alt-RTM Genome Editing Detection Kit (IDT), or the Guide-itTM Complete sgRNA Screening System (Takara Bio, Inc.).
An alternative to AAV production that may be less costly and less complex to scale up is non-viral T-cell engineering using CRISPR-Cas9 and virus-free single-stranded DNA as a template donor, as recently demonstrated for IL-7-IL-15-expanded, blood-derived TSCM at a clinical scale [8]. Another advantage of single-stranded DNA template donors is that they can be synthesized to approximately 8,000 nucleotides in length, enabling the use of longer repair templates than AAV, which has only a packaging capacity of 4–4.4 kb.
For clinical applications, it is necessary to establish safety profiles for the gRNAs used and to carefully assess potential genome-wide off-target sites [38,39]. To this end, we have successfully used Tn5 transposase-mediated GUIDE-seq (see Li et al. [2]; Tran et al. [35]). To improve the fidelity of gene repair, an engineered high-fidelity Cas9 such as Alt-RTMS.p. HiFi Cas9 Nuclease V3 from IDT should be used [40].
For scale-up and good manufacturing practice (GMP)-compliant experiments, we suggest following the procedure described by Shy et al. [8] (Figure 5C). Shy et al. [8] expanded electroporated TSCM in G-Rex 100 M gas-permeable culture vessels (Wilson Wolf Manufacturing) in media supplemented with 100 U/mL of IL-7 and 10 U/mL IL-15 every 2–3 days for up to 10 days.
Troubleshooting
Problem Potential solution
Failed ICE analysis (step A4) Try to improve the PCR, the sequencing quality, or both. A clearly defined PCR band and high-quality sequencing data are required for successful ICE analysis.
Low gene knock-out efficiency
(step A4)
Efficient gRNAs are required: Try different gRNAs. Please test several gRNAs to identify highly efficient ones.
Inefficient AAV production/low AAV titers (section E) Use single-stranded DNA as a donor template if the size of the donor template is close to 4.4 kb.
Inefficient expansion of T cells after gene editing (step F18/F19) Expand cells gradually from a 96-well U-bottom plate to a bigger plate (48-, 24-, 12-, and 6-well plates) and monitor the culture every day until cells have sufficiently expanded to be used in downstream analysis or applications.
Acknowledgments
Funding: This research was supported by the European Research Council, Advanced Grant (268921, to K.R.), the Helmholtz Association, Immunology and Inflammation (ZT-0027, to K.R.), the Berlin School of Integrative Oncology (BSIO) graduate program (Ph.D. position) (to X.L.), and the library of the Max Delbrück Center for Molecular Medicine in the Helmholtz Association, Open Access Fund (to K.R.). We thank Yue Zong and Ralf Kühn for helpful comments on the manuscript. The Graphical Overview and Figures 2 and 4 were made in BioRender.com. The original research paper [2] is available for download from Klaus Rajewky’s web site: https://www.mdc-berlin.de/k-rajewsky. The Supplementary Material of Li et al. [2] is freely available.
Original research paper: The protocol was used and validated in Sci Immunol. (2024) 9(92), eadi0042. doi: 10.1126/sciimmunol.adi0042.
Competing interests
X.L., V.T.C., and K.R. are inventors on an international patent (PCT/EP2022/073414, publication date 02 March 2023; Cas9 nickase-mediated gene editing). The remaining author (C.K.) declares that she has no financial or non-financial competing interests.
Ethical considerations
Informed consent was obtained from the legal guardians of the patient with FHL2 and the IRB of the Hamburg State Medical Association approved the study (ID PV5777).
References
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Supplementary information
The following supporting information can be downloaded here:
Figure S1. Two-part gRNA hybridized to form a crRNA:tracr RNA complex.
Figure S2. gRNA design with free CrispRGold software (Graf et al., 2019).
Figure S3. gRNA design with free CrispRGold software (Graf et al., 2019).
Figure S4. Output of CrispRGold (Graf et al., 2019) gRNA design request with off-target sites shown in expanded mode.
Figure S5. Cloning oligo nucleotides with BbsI sticky ends for gRNA standard vectors (pX330, MSCV retroviral vectors, etc.).
Figure S6. Analysis of the genome editing efficiency of the guide RNA PRF1.2 with the free web-based tool “inference of CRISPR edits” (ICE) (Conant et al., 2022).
Figure S7. Plasmid map of the pAAV-GFP control vector.
Figure S8. Example of HDR, NHEJ, and WT events after gene repair as determined by Sanger sequencing.
Article Information
Publication history
Received: May 8, 2024
Accepted: Aug 16, 2024
Available online: Sep 25, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Improving Stability of Spiroplasma citri MreB5 Through Purification Optimization and Structural Insights
Vani Pande
PG Pananghat Gayathri
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5086 Views: 372
Reviewed by: Joana Alexandra Costa Reis Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology May 2022
Abstract
MreB is a prokaryotic actin homolog. It is essential for cell shape in the majority of rod-shaped cell-walled bacteria. Structural and functional characterization of MreB protein is important to understand the mechanism of ATP-dependent filament dynamics and membrane interaction. In vitro studies on MreBs have been limited due to the difficulty in purifying the homogenous monomeric protein. We have purified MreB from the cell-wall-less bacteria Spiroplasma citri, ScMreB5, using heterologous expression in Escherichia coli. This protocol provides a detailed description of purification condition optimization that led us to obtain high concentrations of stable ScMreB5. Additionally, we have provided a protocol for detecting the presence of monovalent ions in the ScMreB5 AMP-PNP-bound crystal structure. This protocol can be used to obtain a high yield of ScMreB5 for carrying out biochemical and reconstitution studies. The strategies used for ScMreB5 show how optimizing buffer components can enhance the yield and stability of purified protein.
Key features
• The protocol is a useful approach to standardize purification of nucleotide-dependent cytoskeletal filaments and other nucleotide-binding proteins.
• The mechanistic basis of how different ions could stabilize a protein, and hence improve yield in purification, has been demonstrated.
• The change in buffer conditions/salt enabled us to get sufficient yield for biochemical and structural characterization.
Keywords: MreB Cytoskeleton proteins ADP Thermal shift assay Affinity chromatography Size exclusion chromatography Crystallization X-Ray adsorption spectroscopy
Graphical overview
Background
There is an array of prokaryotic filamentous proteins that play a pivotal role in maintaining overall cellular functioning [1]. In the majority of bacteria, complex cell wall synthesis machinery plays a major role in cell shape maintenance. Studies have extensively analyzed how shape is regulated in E. coli and B. subtilis. Genetic studies show that deleting mreB causes loss of rod shape and eventual lysis, indicating the pivotal role of the MreB protein in bacterial cell shape and maintenance [2–4].
MreB forms antiparallel double protofilaments in the presence of nucleotide, ATP, or ADP that align parallel to the long axis of the rod-shaped cells [5]. The processive movement of MreB along with the other components of cell wall synthesis machinery drives the insertion of peptidoglycan, thus enforcing the cell shape [6,7].
MreB5 is responsible for determining helicity and motility in the cell-wall-less bacteria Spiroplasma citri [8]. In our recent work, we reported the structure and biochemical characterization of ScMreB5 and its mutants [9]. We showed that the nucleotide state of ScMreB5 determines the filament dynamics and membrane interaction. MreBs are notoriously challenging to purify because they tend to aggregate when overexpressed. Obtaining monomeric and homogeneous proteins for biochemical and structural studies has been challenging, resulting in few MreBs being characterized in vitro.
For in vitro characterization and structure determination, the scmreb5 gene was cloned into the pHis17 expression vector, under the T7 promoter, between NdeI and BamHI restriction endonuclease site. The cloned gene was codon-optimized by mutating the TGA codon to TGG, since in S. citri, UGA is recognized as tryptophan [10]. A restriction-free cloning strategy [11] was chosen for cloning, and the clone was confirmed by DNA sequencing. The cloned plasmid was transformed into the BL21-AI expression strain, which has a T7 RNA polymerase tightly regulated by the araBAD promoter and induced with arabinose for overexpression.
This protocol paper describes optimizing ScMreB5 protein purification, where precipitation in NaCl-containing buffer was prevented by adding excess ADP and MgCl2 to stabilize the protein. This addition prevented any further biochemical characterization of ScMreB5. To eliminate this issue, we optimized the buffer conditions for protein purification using a thermal shift assay [12,13]. The optimized buffer containing KCl removed the necessity for ADP and MgCl2 addition during the purification process. To understand the structural significance of KCl in stabilizing the protein, we performed X-ray adsorption spectroscopy on ScMreB5-AMP-PNP crystals. This technique is widely used for detecting metal ions within a protein crystal. Since the X-ray absorption edges of different elements occur at specific energies, it allows the identification of specific metal ions within the crystal. The presence of potassium ion bridging the protein with the nucleotide provided a structural basis for why the stability of ScMreB5 improved in the presence of KCl.
Materials and reagents
For transformation and large culture expression
100 mm Petri dish (Himedia, catalog number: PW1305-1x100NO)
Luria Bertani agar, LB agar (Himedia, catalog number: M1151-1KG)
Luria Bertani broth powder (Himedia, catalog number: M1245-1KG)
BL21-AI chemically competent E. coli cells (Invitrogen, catalog number: C6070-03)
ScMreB5 cloned pHis17 plasmid (Hexa His tag at the C-terminus)
Ampicillin (Sigma-Aldrich, catalog number: A9518-25G)
100 mL conical flask (Borosil)
2 L conical flask (Borosil)
0.22 µm syringe filter (Cytiva, catalog number: WHA-9914-2502)
10 mL syringe for single use (Dispovan, catalog number: JK-10ml-0110)
L-Arabinose (Himedia, catalog number: GRM037-25G)
For purification
150 mL beaker (Borosil, catalog number: 1000D18)
0.5 mL centrifuge tubes (Tarsons, catalog number: 500000)
1.5 mL centrifuge tubes (Tarsons, catalog number: 500010)
0.2 mL PCR tubes (Tarsons, catalog number: 510051)
50 mL conical centrifuge tubes (Tarsons, catalog number: TAR-500041)
100 mL conical flasks (Borosil, catalog number: 5020016)
500 mL conical flasks (Borosil, catalog number: 5020024)
2 L conical flasks (Borosil, Catalog number: 5020030)
Test tubes, 15 × 125 mm (Borosil, catalog number: 9800U04)
HiIndicator pH paper, pH range 6.5–9.0 (Himedia, catalog number: LA321-1PK)
5 mL His-Trap column (Cytiva, catalog number: 17524801)
Bradford 1× dye reagent (Bio-Rad, catalog number: 5000205)
Bovine serum albumin (BSA) (Himedia, catalog number: TC548-5G)
Durapore membrane filter, 0.22 µm, hydrophilic PVDF, 47 mm membrane (Sigma-Aldrich, catalog number: GVWP04700)
Superdex 200 Increase 10/300 GL (Cytiva, catalog number: 28990944)
Oak Ridge centrifuge tubes, 50 mL (Thermo Scientific, catalog number: 3118-0050)
Thick-walled Polycarbonate tubes, 11 × 34 mm, 1 mL (Beckman Coulter, catalog number: 343778)
Hydrochloric acid (Qualigens, catalog number: Q29145)
Tris, free base, for molecular biology (Himedia, catalog number: MB029-5KG)
Glycine (Sigma-Aldrich, catalog number: G8898-1KG)
Sodium dodecyl sulphate (Sigma-Aldrich, catalog number: 436143)
TEMED (Tetramethylethylenediamine) (Sigma-Aldrich, catalog number: T9281-25ML)
Ammonium persulphate (Sigma-Aldrich, catalog number: A3678)
Acrylamide (Sigma-Aldrich, catalog number: A8887-500G)
Bis-acrylamide (Sigma-Aldrich, catalog number: 146072-500G)
0.75 mm spacer plate (Bio-Rad, catalog number: 1653310)
Short plates (Bio-Rad, catalog number: 1653308)
10-well comb, 0.75 mm (Bio-Rad, catalog number: 1653354)
15-well comb, 0.75 mm (Bio-Rad, catalog number: 1653355)
Coomassie Brilliant Blue (R-250) (Sigma-Aldrich, catalog number: 27816)
2× Laemmli sample buffer (Bio-Rad, catalog number: 1610737)
DTT (Sigma-Aldrich, catalog number: D9779-25G)
Ethanol for molecular biology (Omnis)
Glacial acetic acid (Qualigens, catalog number: Q11007)
Sodium chloride (NaCl) (Qualigens, catalog number: Q15918)
Potassium chloride (KCl) (Qualigens, catalog number: Q13305)
Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: M8266-100G)
ADP adenosine 5'-diphosphate sodium salt (Sigma-Aldrich, catalog number: A2754-500MG)
Imidazole (Sigma-Aldrich, catalog number: I2399-500G)
Glycerol (Fisher Scientific, catalog number: 15457)
Dialysis tubing 10 kDa MWCO (Thermo Fisher Scientific, catalog number: 68100)
Dialysis tubing clamps (Sigma-Aldrich, catalog number: Z371092-10EA)
Vivaspin® Turbo 15 PES 10 kDa MWCO (Sartorius, catalog number: VS15T01)
Precision Plus Protein Dual Color Standards (Bio-Rad, catalog number: 1610394)
Micro test plate, flat Bottom (Tarsons, catalog number: 941196)
Liquid nitrogen
For thermal shift assay
MultiplateTM 96-well PCR plates, low profile, unskirted, white (Bio-Rad, catalog number: MLL9651)
Microseal 'B' PCR plate sealing film, adhesive, optical (Bio-Rad, catalog number: MSB1001)
SYPRO orange (Sigma-Aldrich, catalog number: S5692)
For crystallization
48-well crystallization plate (Hampton Research, catalog number: HR3-180)
Sodium phosphate monobasic monohydrate (Sigma-Aldrich, catalog number: 71507-250G)
Sodium phosphate dibasic (Sigma-Aldrich, catalog number: S9763-100G)
AMP-PNP Adenosine 5'-(β, γ-imido) triphosphate lithium salt hydrate (Sigma-Aldrich, catalog number: 10102547001)
Magnesium chloride (details mentioned above)
PEG 3350 (Sigma-Aldrich, catalog number: 202444-500G)
Glycerol (Sigma-Aldrich, catalog number: G5516)
CrystalCap SPINE HT, 0.05-0.1 mm CryoLoop (Hampton Research, catalog number: HR8-120)
Stock solutions (see Recipes)
100 mg/mL ampicillin
1 mg/mL BSA
2 M MgCl2
1 M unbuffered Tris
100 mM ADP
100 mM AMP-PNP
20% Arabinose (w/v)
5 M NaCl
3 M KCl
2 M Tris pH 8
2 M Imidazole
30% Acrylamide solution (w/v)
10% Ammonium persulphate (w/v)
10% SDS
1.5 M Tris pH 6.8
1.5 M Tris, pH 8.8
1 M sodium phosphate monobasic monohydrate
1 M sodium phosphate dibasic
1 M sodium phosphate buffer, pH 7.8
40% PEG 3350 (w/v)
20% Glycerol (v/v)
50× SYPRO Orange
Solutions and buffers for purification (see Recipes)
10× TGS buffer for SDS-PAGE gels
SDS-PAGE staining solution
SDS-PAGE destaining solution
5% SDS-PAGE stacking gel
12% SDS-PAGE resolving gel
Lysis buffer (Na)
Buffer A (Na)
Buffer B (Na)
Storage buffer
Lysis buffer (K)
Buffer A (K)
Buffer B (K)
Crystallization conditions
Recipes
Materials to be autoclaved
LB Agar
Dissolve 4 g of LB Agar in RO (reverse osmosis) water to a final volume of 100 mL in a 500 mL conical flask. Autoclave at 121 °C for 15 min at 2.8 bar pressure for sterilization. Once cooled enough to touch, add 100 µL of 100 mg/mL ampicillin solution to the LB agar and gently stir. Pour 25 mL of LB agar into the Petri plates. Store the plates at 4 °C after the LB agar solidifies.
1× LB Broth, 30 mL
Dissolve 0.75 g of Luria Bertani broth powder in RO water to a final volume of 30 mL in a 100 mL conical flask. Autoclave at 121 °C for 15 min at 2.8 bar pressure for sterilization.
1× LB Broth, 1,000 mL
Dissolve 25 g of Luria Bertani broth powder in RO water to a final volume of 1,000 mL in a 2 L conical flask. Autoclave at 121 °C for 15 min at 2.8 bar pressure for sterilization.
15 × 12.5 mm test tubes
Autoclave 20–25 clean test tubes at 121 °C for 15 min at 2.8 bar pressure for sterilization.
Stock solutions
100 mg/mL Ampicillin
Dissolve 0.5 g of ampicillin powder in ultrapure water to a final volume of 5 mL. Sterilize the solution using a 0.22 µm syringe filter.
1 mg/mL BSA
Dissolve 1 mg of BSA powder in ultrapure water to a final volume of 1 mL. Use this solution to prepare standards (0.1–1.0 mg/mL) in ultrapure water for protein concentration estimation using Bradford reagent.
2 M MgCl2
Dissolve 1.90 g of MgCl2 powder in ultrapure water to a final volume of 10 mL. Filter the solution using a 0.22 µm syringe filter.
1 M unbuffered Tris
Dissolve 1.21 g of unbuffered Tris-base powder in ultrapure water to a final volume of 10 mL. Filter the solution using a 0.22 µm syringe filter.
100 mM ADP
Dissolve 23.5 mg of ADP powder in 100 µL of ultrapure water. Adjust the pH to 7–7.5 with 1 M unbuffered Tris using a pH strip. Make up the volume to 250 µL.
100 mM AMP-PNP
Dissolve 5 mg of ADP powder in 30 µL of ultrapure water. Adjust the pH to 7–7.5 with 1 M unbuffered Tris using a pH strip. Make up the volume to 100 µL.
20% (w/v) Arabinose
Dissolve 10 g of arabinose in RO water to a final volume of 50 mL. Filter the solution using a 0.22 µm syringe filter.
5 M NaCl
Dissolve 146.1 g of NaCl powder in RO water to a final volume of 500 mL. Filter the solution using a vacuum filter assembly with a 0.22 µm membrane filter.
3 M KCl
Dissolve 111.8 g of KCl powder in RO water to a final volume of 500 mL. Filter the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane).
2 M Tris, pH 8
Dissolve 60.7 g of Tris-base in 120 mL of RO water. Adjust the pH to 8 using concentrated hydrochloric acid using a pH meter. Make up the volume to 250 mL. Filter the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane).
Note: Follow the required lab safety procedures while handling hydrochloric acid.
2 M Imidazole
Dissolve 68.07 g of imidazole powder in 500 mL of RO water. Filter the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane).
30% Acrylamide solution
Dissolve 146 g of acrylamide and 3.9 g of bis-acrylamide powders in 500 mL of RO water. Filter the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane).
10% Ammonium persulphate
Dissolve 5 g of ammonium persulphate powder in RO water to a final volume of 50 mL. Filter the solution using a 0.22 µm syringe filter.
10% SDS
Dissolve 5 g of SDS powder in RO water to a final volume of 50 mL.
1.5 M Tris pH 6.8
Dissolve 45.5 g of Tris-base in 120 mL of RO water. Adjust the pH to 6.8 using concentrated hydrochloric acid and a pH meter. Make up the volume to 250 mL. Filter the buffer using a 0.22 µm syringe filter.
1.5 M Tris, pH 8.8
Dissolve 45.5 g of Tris-base in 120 mL of RO water. Adjust the pH to 8.8 using concentrated hydrochloric acid and a pH meter. Make up the volume to 250 mL. Filter the buffer using a 0.22 µm syringe filter.
1 M sodium phosphate monobasic monohydrate
Dissolve 1.37 g of sodium phosphate monobasic monohydrate powder in ultrapure water to a final volume of 10 mL. Filter the solution using a 0.22 µm syringe filter.
1 M sodium phosphate dibasic
Dissolve 1.42 g of sodium phosphate dibasic powder in ultrapure water to a final volume of 10 mL. Filter the solution using a 0.22 µm syringe filter.
1 M sodium phosphate buffer, pH 7.8
Titrate sodium phosphate monobasic monohydrate using 1 M sodium phosphate dibasic to obtain pH 7.8. Use HiIndicator pH Paper, pH range 6.5–9.0 for adjusting the pH.
40% PEG 3350
Dissolve 20 g of PEG 3350 powder in ultrapure water to a final volume of 50 mL. Filter the solution using a 0.22 µm syringe filter. Use this stock to make 16% PEG 3350.
20% Glycerol
Mix 2 mL of glycerol with 8 mL of ultrapure water. Filter the solution using a 0.22 µm syringe filter.
50× SYPRO Orange
Mix 1 µL of 5,000× SYPRO Orange in 9 µL of DMSO to make 500× SYPRO orange. Dilute 500× SYPRO orange to 50× by mixing 2 µL of 500× SYPRO orange in 18 µL in ultrapure water.
Buffers and working stock solutions
10× TGS Buffer
Dissolve 30 g of Tris-base, 144 g of glycine, and 10 g of SDS in 1,000 mL of RO water. Use it to make 1× TGS buffer for running the SDS-PAGE gels.
Coomassie staining solution
Dissolve 2.5 g of Coomassie Brilliant Blue (R-250) in 500 mL of ethanol by stirring. Add 100 mL of glacial acetic acid and 400 mL of RO water to make the final volume of 1,000 mL.
De-staining solution
Mix 100 mL of glacial acetic acid and 200 mL of ethanol with 700 mL of RO water.
12% SDS-PAGE resolving gel (5 mL)
Mix the following solutions and pour onto the 0.75 mm clamped gel setup.
RO water (1.6 mL)
30% acrylamide solution (2 mL)
Tris-HCl, pH 8.8 (1.5 mL)
10% SDS (50 µL)
10% ammonium persulphate (50 µL)
TEMED (3 µL)
Add 100 µL of 100% isopropanol to even out the layer of resolving gel. Once solidified, carefully tilt the gel setup to one side and use tissue paper to remove isopropanol.
5% SDS-PAGE stacking gel (1 mL)
Mix the following solutions and pour over the solidified resolving gel.
RO water (680 µL)
30% acrylamide solution (170 µL)
Tris-HCl, pH 6.8 (130 µL)
10% SDS (10 µL)
10% ammonium persulphate (10 µL)
TEMED (1 µL)
After pouring the stacking solution, place a 10- or 15-well 0.75 mm comb.
Lysis buffer (Na)
200 mM NaCl (10 mL from 5 M NaCl)
50 mM Tris-HCl pH 8 (6.25 mL from 2 M Tris-HCl pH 8)
10% glycerol (12.5 mL from 100% glycerol)
Make up the volume to 250 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C for purification.
Buffer A (Na)
Used as binding buffer in Ni-NTA chromatography.
200 mM NaCl (20 mL from 5 M NaCl)
50 mM Tris-HCl pH 8 (12.5 mL from 2 M Tris-HCl pH 8)
Make up the volume to 500 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C for purification.
Buffer B (Na)
Used as elution buffer in Ni-NTA chromatography.
200 mM NaCl (20 mL from 5 M NaCl)
50 mM Tris-HCl pH 8 (12.5 mL from 2 M Tris-HCl pH8)
500 mM imidazole (125 mL from 2 M imidazole)
Make up the volume to 500 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C before using for purification.
Storage buffer
Used for buffer exchange and protein storage.
50 mM NaCl (1 mL from 5 M NaCl)
50 mM Tris-HCl pH 8 (2.5 mL from 2 M Tris-HCl pH 8)
Make up the volume to 100 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C for purification.
Lysis buffer (K)
300 mM KCl (25 mL from 3 M KCl)
50 mM Tris-HCl pH 8 (6.25 mL from 2 M Tris-HCl pH 8)
10% glycerol (12.5 mL from 100% glycerol)
Make up the volume to 250 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C for purification.
Buffer A (K)
Used as binding buffer in Ni-NTA chromatography and buffer for size exclusion chromatography.
300 mM KCl (100 mL from 3 M KCl)
50 mM Tris-HCl pH 8 (25 mL from 2 M Tris-HCl pH 8)
Make up the volume to 1000 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C for purification.
Buffer B (K)
Used as elution buffer in Ni-NTA chromatography.
300 mM KCl (50 mL from 3 M KCl)
50 mM Tris-HCl pH 8 (12.5 mL from 2 M Tris-HCl pH 8)
500 mM imidazole (125 mL from 2 M imidazole)
Make up the volume to 500 mL with RO water. Filter and degas the solution using a vacuum filter assembly with DuraporeTM membrane filter (0.22 µm, hydrophilic PVDF, 47 mm membrane). Pre-chill the buffer at 4 °C before using for purification.
Crystallization condition for X-ray fluorescence scanning
0.15 M sodium phosphate buffer, pH 7.8 (150 µL from 1 M sodium phosphate buffer, pH 7.8)
16% PEG 3350 (400 µL from 40% PEG 3350)
Make up the volume to 1000 µL with ultrapure water. Sterilize using a 0.22 µm syringe filter.
Equipment
Shaker incubator (Thermo Scientific, model: MaxQTM 6000)
Eppendorf® ThermoMixer® F2.0 (EP5387000013)
Sonicator (Sonics VibraCell)
E-GelTM imager system (Fisher Scientific, catalog number: 4466611)
pH meter (Thermo Scientific, model: ORION STAR A111)
Vacuum filter assembly (Millipore, model: XX1014720)
Centrifuge (Avanti J26S-XP Non-IVD)
Tabletop ultracentrifuge (Beckman Coulter Optima MAX-XP)
High-speed centrifuge (Eppendorf, model: 5430 R)
Benchtop centrifuge (Eppendorf, model: 5810 R)
Centrifuge (Eppendorf MiniSpin®)
ÄKTA PrimeTM Plus FPLC system
Bio-Rad NGCTM chromatography system QuestTM 10
SDS-PAGE apparatus (Bio-Rad, catalog number: 1658002FC)
CLARIOstar® microplate reader
Bio-Rad CFX96 Touch Real-Time PCR Detection System
Software and datasets
GraphPad Prism (for analyzing and plotting thermal shift assay and size exclusion chromatography)
Microsoft Excel (for exporting thermal shift assay results)
ImageJ (for SDS-PAGE gel)
Procedure
Large culture expression
Take 100 ng of the plasmid and add into pre-thawed 50 µL of BL21-AI competent cells under sterile conditions. Incubate for 30 min on ice.
Give a heat shock at 42 °C for 90 s and immediately place the tube on ice for 5 min.
Add 200 µL of LB media into the tube and incubate for 40 min at 37 °C for recovery in shaking condition.
Plate 100 µL of the culture in LB agar plate containing 100 µg/µL of ampicillin antibiotic. Incubate the plate for 10–12 h at 37 °C.
Note: Follow the manufacturer’s protocol when using the commercially available competent BL21-AI cells.
Primary culture:
Next day, inoculate 10–20 colonies in 30 mL of LB media containing 30 µL (100 µg/µL) of ampicillin antibiotic in a 100 mL conical flask.
Grow the culture until OD600 reaches 0.8 at 37 °C in a shaker incubator (180 rpm).
Secondary culture:
Inoculate 10 mL of primary culture into two 2 L conical flasks containing 1 L of LB media.
Add 1,000 µL (100 µg/µL) of ampicillin antibiotic into the media.
Grow the secondary culture until OD600 reaches 0.8–1.0 at 37 °C in a shaker incubator (180 rpm).
Note: It should take 2.5–3 h to reach OD600 of 0.8.
Induction:
Change the shaker incubator temperature to 18 °C.
To induce the secondary cultures, add 10 mL of 20% arabinose to each flask under sterile conditions.
Incubate the culture flasks at 18 °C in the shaker incubator for 10–12 h for protein overexpression.
Cell pelleting:
Pellet down the cells from 2 L culture in JLA 9.1 rotor using an Avanti J26S-XP. Centrifuge at 7,500–8,000 RCF for 20 min at 4 °C.
Discard the supernatant, leaving 10 mL of supernatant for resuspending the pellet.
Transfer the resuspended pellet into 50 mL centrifuge tubes, spin down the cells at 3,200 RCF using a benchtop centrifuge, and discard the supernatant.
Plunge freeze the cell pellet containing the centrifuge tubes in liquid nitrogen and store at -80 °C until further use.
Note: Handle liquid nitrogen with caution. Wear safety glasses and cryo-gloves during use.
Protein purification in NaCl-containing buffer
Cell pellet lysis
Thaw 2 L of culture cell pellet on ice.
Resuspended the cell pellet in 50 mL lysis buffer (Na).
Sonicate the cell suspension using a probe sonicator. Keep the following settings: 1 s ON, 3 s OFF, total time = 3 min, amplitude = 60%. Perform sonication twice with a 5 min resting time between the two cycles.
Cell debris removal
Centrifuge the lysate at 45,000 RCF for 45 min at 4 °C using JA 25.5 rotor in Avanti J26S-XP centrifuge to remove the cell debris.
Transfer the supernatant into a pre-chilled 50 mL centrifuge tube on ice.
Ni-NTA purification
Wash and equilibrate the Akta Prime plus system with Buffer A (Na).
Equilibrate the 5 mL His-Trap column by passing five column volumes of Buffer A (Na).
Load the supernatant at a flow rate of 2.5 mL/min onto the column. The Hexa-His tag at the C-terminus facilitates the binding of the protein to the Ni-NTA beads.
Collect the flowthrough in a 100 mL conical flask.
After loading the supernatant, wash the column with 15 column volumes of Buffer A (Na).
Collect 10 column volumes each of 2% and 5% Buffer B (Na) in 100 mL conical flasks.
Perform fraction collection (5 mL fractions, 5 column volumes) for 10%, 20%, 50%, and 100% Buffer B (Na) in 15 × 12.5 mm autoclaved test tubes.
Observe the UV280 peak on the chromatogram. As the protein elution peak is observed, add 1 mM ADP and 1 mM MgCl2 to the fractions immediately to prevent precipitation.
Run a 12% SDS-PAGE gel for the lysate, supernatant, wash, flowthrough, 2%, 5%, and the fractions along with a protein ladder.
The protein elutes from the last fraction of 20% and all fractions of 50% Buffer B (Na).
Pool the fractions (after analyzing them on the gel) that contain a single band of the protein corresponding to 37 kDa relative to the protein ladder.
Note: Ensure that the fraction has 1 mM ADP and 1 mM MgCl2 added to avoid protein precipitation.
Centrifuge the pooled protein fractions at 45,000 RCF for 30 min at 4 °C using JA 25.5 rotor in Avanti J26S-XP centrifuge to remove any protein precipitants.
Concentration and buffer exchange
Wash and equilibrate a 10 kDa molecular weight cutoff concentrator with Buffer A (Na). Concentrate the pooled fractions in the concentrator at 3,200 RCF in an Eppendorf benchtop centrifuge 5810 R at 4 °C.
As the protein concentrates, observe for any visible precipitation. If precipitation occurs, centrifuge the protein at 21,000 RCF at 4 °C in an Eppendorf high-speed centrifuge.
Add 1 mM ADP and 1 mM MgCl2 to the supernatant to avoid any further precipitation.
Simultaneously, clean the concentrator using Buffer A (Na). Add the supernatant back to the concentrator.
Once the protein volume reaches ~1 mL, dilute it with ~10 mL of storage buffer, 1 mM ADP, and 1 mM MgCl2 for buffer exchange. Concentrate the protein until the concentration reaches ~2 mg/mL. Estimate the protein concentration by performing the Bradford assay as described below.
In the Eppendorf high-speed centrifuge, spin the concentrated protein at 21,000 RCF for 20 min at 4 °C in a 1.5 mL centrifuge tube.
Make 10 µL protein aliquots in 0.2 mL PCR tubes. Plunge freeze the protein aliquots in liquid nitrogen and store them at -80 °C until further use.
Notes:
i. This protein will be used to perform thermal shift assay for optimizing the purification conditions for increasing protein stability.
ii. Wear safety glasses and cryo-gloves while handling liquid nitrogen.
Bradford assay
Make BSA standards from 0.1 to 1.0 mg/mL using 1 mg/mL of BSA stock (see Recipes).
In a clear 96-well microplate, aliquot 5 µL of BSA standards, 5 µL of ultrapure water (blank for the standards), and 5 µL of storage buffer/buffer A (blank for the protein).
Prepare 20 µL of 5×, 10×, and 20× dilutions of purified protein in Buffer A.
Aliquot 5 µL of dilutions into the 96-well microplate in triplicates.
Add 250 µL of Bradford 1× dye reagent to the wells.
Measure the absorbance at 595 nm in a plate reader.
Estimate the protein concentration using Microsoft Excel as follows:
i. Subtract the readings of the standards and the protein dilutions with the respective readings of the blank.
ii. Average the triplicate readings for the protein dilutions.
iii. Plot a standard curve in Microsoft Excel using the scatterplot option.
iv. Format the trendline as “linear” and set the intercept.
v. Calculate the slope and R-squared value.
vi. Using the slope value (m) and absorbance reading (y), calculate the protein concentration (×) using the formula:
y
=
m
x
Note: The Bradford assay kit can also be used for protein concentration estimation. Follow the instructions from the kit for estimating the concentration.
Thermal shift assay
Protein concentration estimation
Thaw the protein aliquots (purified as described above) on ice. Transfer the aliquots to a thick-walled polycarbonate tube.
Ultracentrifuge at 100,000 RCF for 20 min at 4 °C in the OptimaTM Max-XP table-top ultracentrifuge.
Estimate the protein concentration of the supernatant by performing the Bradford assay as discussed in section B.
Performing a thermal shift assay
Note: Optimization experiments for ScMreB5 included testing various buffering agents (Tris, HEPES), pH ranges (5–10), salts (NaCl, KCl), and various additives (CaCl2, MgCl2, ADP, AMP-PNP, GDP, GTP, EDTA).
The protocol outlined below is a generalized protocol that was also used during ScMreB5 purification optimization. Different conditions can be tested in a similar manner. The table below is an example of a reaction setup for comparing NaCl and KCl salts at a concentration of 500 mM.
Prepare the reaction given in Table 1 in a 0.5 mL centrifuge tube on ice.
Table 1. Thermal shift assay reaction composition
Components (stock concentration) Reaction (µL)
NaCl condition KCl condition
Protein (~ 50 µM) 1.25 µL 1.25 µL
Tris-HCl, pH 8 (100 mM) 12.5 µL 12.5 µL
Salt: NaCl (5 M) 2.5 µL
Salt: KCl (3 M) 4.2 µL
Ultrapure water 6.25 µL 4.55 µL
SYPRO Orange (50×) 2.5 µL 2.5 µL
Total reaction volume 25 µL 25 µL
Replace KCl/NaCl with any other salt condition (with different concentrations) to be tested from the above table.
Transfer the reaction to a MultiplateTM 96-well PCR plate and seal the reaction well with Microseal 'B' PCR plate sealing film.
Spin the reaction in the Eppendorf centrifuge 5810 R at 2,500 RCF for 1 at 4 °C min to bring the reaction mix to the bottom of the well.
Place the plate in a Bio-Rad CFX96 Real-Time System.
Set up the protocol as follows:
i. Open CFX Maestro software and create a new Protocol by going to File > New tab.
ii. In the Protocol Editor window, select Before in the Insert Step tab and click on Insert Melt Curve.
iii. Remove the step 2–4 by clicking on Delete Step. Only step 1 is required for the assay.
iv. Set the temperature range to 4–90 °C with a rise of 0.4 °C every 20 s in the step 1.
v. Set the sample volume to 25 µL.
vi. Save the protocol by clicking OK.
vii. Select the Next option and click on Create New.
viii. The Plate Editor window will open. Select the wells to be analyzed by dragging the cursor across the wells.
ix. In the Scan Mode tab, select FRET (excitation: 470 nm; emission: 569 nm).
x. In the Sample Type menu, choose unknown option.
xi. Under Target Names tab, tick the Load Box.
xii. Click on OK when finished.
Use the Run tab to initiate the reaction. (It should take around 1 h 20 min with the above settings.)
After run completion, export the Melt Curve Derivative Results as an Excel file and proceed for analysis.
Open GraphPad Prism and paste the readings in an XY format. The readings of the conditions will be in (-(dRFU)/dT) format.
The first derivative of the florescence (-(dRFU)/dT) is plotted against the temperature. The dip in the curve determines the Tm of the protein for a given condition.
A representative image of an expected melting curve for the above reaction is provided in Figure 1. The original data for 300 mM KCl buffer, used for purifying ScMreB5, has been provided in the original publication as Figure S1, C and D [9].
Figure 1. Representative thermal shift assay for purification condition optimization. Melting curve for ScMreB5 showing Tm for 500 mM NaCl (red) and 500 mM KCl (blue). The buffer used in both conditions is 50 mM Tris, pH 8. The first derivative of the fluorescence (-(dF)/dT) is plotted against the temperature. The dip in the curve determines the Tm of the protein in the given condition. RFU is the relative florescence units.
Protein purification in KCl-containing buffer
After condition optimization, the following changes were made to the protocol in Section B “Protein Purification in NaCl containing buffer”:
A. Lysis buffer (Na) is replaced with lysis buffer (K).
B. Buffer A (Na) is replaced with Buffer A (K).
C. Buffer B (Na) is replaced with Buffer B (K).
D. ADP and MgCl2 were not added at any stage during purification.
E. Purified protein is stored in Buffer A (K)
Cell pellet lysis (as described previously)
Cell debris removal (as described previously)
Ni-NTA purification (as described previously) (Figure 2)
Figure 2. Ni–NTA affinity chromatography of ScMreB5 purified in the optimized buffer condition containing KCl. Representative 12% SDS-PAGE gel of the purified protein after Ni-NTA chromatography; 5 µL of sample mixed with 5 µL of 2× Laemmli buffer is loaded in the gel. Purest fractions (within the dotted rectangular box) are pooled and concentrated to ~1 mL for size exclusion chromatography. Lys: lysate, Sup: supernatant, FT: flowthrough, W: wash. Percentage values denote Buffer B (K) percentage going through the Ni-NTA column along with Buffer A (K).
Size exclusion chromatography
Concentrate the protein to 800 µL in 10 kDa molecular weight cut-off concentrators.
Pre-equilibrate the Superdex 200 column with two column volumes of Buffer A (K).
Inject the concentrated protein into the system using a 1 mL loop.
Elute the protein in 0.5 mL fractions across one column volume of Buffer A (K). The protein will elute at around 14.5 mL, corresponding to the monomeric size of the protein (Figure 3A).
Load the samples from the monomeric protein fractions corresponding to the protein on a 12% SDS-PAGE gel (Figure 3B).
Pool the fractions containing the pure protein after confirming from the gel.
Figure 3. Size-exclusion chromatography of ScMreB5 purified in the optimized buffer condition. (A) Elution profile of the protein eluted during size exclusion chromatography from Superdex-200. Protein starts eluting from ~14.3 mL. The y-axis in the graph represents the milli absorbance unit (mAU). (B) Representative 12% SDS-PAGE gel of the fractions containing monomeric protein from the size exclusion chromatography. “Input (Ni-NTA)” is the concentrated protein from Ni-NTA chromatography; 5 µL of the sample mixed with 5 µL of 2× Laemmli buffer is loaded in the gel. Purest fractions are concentrated and stored at -80 °C.
Concentrate the protein to ~500 µL in a pre-equilibrated 10 kDa molecular weight cut-off concentrator at 3,200 RCF in Eppendorf benchtop centrifuge 5810 R at 4 °C.
Estimate the protein concentration using Bradford assay. The overall yield of the protein will be around 10 mg/mL from a 2 L culture pellet.
In the Eppendorf high-speed centrifuge, spin the concentrated protein at 21,000 RCF for 20 min at 4 °C in a 1.5 mL centrifuge tube.
Prepare 10 µL protein aliquots in 0.2 mL PCR tubes. Plunge freeze the protein aliquots in liquid nitrogen and store at -80 °C until further use.
Running 12% SDS-PAGE Gel
Run SDS-PAGE gel after affinity and size exclusion chromatography as follows:
Mix 10 µL of sample with 10 µL of 2× Laemmli buffer. Heat the sample for 5–10 min at 95 °C in a thermomixer.
Give a quick spin to the sample in a MiniSpin® centrifuge.
Place 12% SDS-PAGE gel in the gel running tank containing 1× TGS.
Load 10 µL of the mix into 12% SDS-PAGE gel along with the protein ladder.
Run the gel at 200 V for 40 min or until the dye front runs out from the gel.
Stain the gel with the staining solution for 5 min.
Destain the gel using the destaining solution for 15 min.
Determining the presence of potassium ion
Crystallization in the absence of K+ in the condition:
Estimate protein concentration using the Bradford assay as described in section B.
Set up crystallization for ScMreB5-AMPPNP as given in the Table 2:
Table 2. Crystallization conditions for ScMreB5 – AMPPNP
Protein ScMreB5 – AMPPNP
Method Sitting drop Vapour diffusion
Plate type 48 well
Temperature (K) 291
Protein concentration 4 mg/mL
Buffer composition of protein 300 mM KCl, 50 mM Tris, pH 8, 2 mM AMPPNP, 2 mM MgCl2
Composition of reservoir solution 0.15 M sodium-phosphate buffer and 16% PEG 3350, pH 7.8
Volume and ratio of drop (µL: µL) 1:1
Volume of the reservoir (mL) 0.1 mL
Add 100 µL of reservoir solution to the reservoir of a 48-well sitting drop crystallization plate.
Prepare a 20 µL reaction containing Buffer A (K), 2 mM AMP-PNP, 2 mM MgCl2, and 4 mg/mL of ScMreB5.
Aliquot 1 µL of reservoir solution into the well and add 1 µL of protein reaction. Mix the reservoir solution with the protein reaction mix.
Prepare 10 such wells for crystallization.
Incubate the plate at 18 °C undisturbed.
After three days, check for needle-like crystals under the microscope.
Pick thick individual needles or thin bundles of needle-shaped crystals using 0.05 or 0.1 mm cryo loops.
Wash out the remaining potassium ion (coming from the protein-containing buffer A) from the crystal by successfully passing the loop through 20% glycerol cryo-protectant drops (made in parent condition). Pass the loop containing crystal at least through three drops.
Freeze the crystal in liquid nitrogen and perform X-ray adsorption spectroscopy at the synchrotron facility.
A peak corresponding to 3.5 keV corresponds to the K+ ion in the scan. The scan plot can be found in the original publication as supplementary Figure S1 A.
Data analysis
Details pertaining to data analysis can be found in the material and methods section of the original research paper [9].
Validation of protocol
This protocol has been validated in the original publication: https://doi.org/10.1083/jcb.202106092
Pande et al. [9]. Filament organization of the bacterial actin MreB is dependent on the nucleotide state. Journal of Cell Biology [(Figure 1 (A–D) and Figure S1 (A–E)].
General notes and troubleshooting
The plasmid used for expressing ScMreB5, pHis17, does not have the lacI repressor gene; therefore, IPTG is not required for expression in BL21-AI cells. This plasmid has been used previously for expressing other MreBs [5,14]. Alternatively, ScMreB5 orthologs can also be cloned pET-derived vectors, having the lacI repressor, as demonstrated for Spiroplasma eriocheris MreB5 [15]. Expression of protein from vectors having the lacI repressor requires the addition of IPTG for expression in BL21-AI, as detailed in the manufacturer’s protocol.
It is important that the entire purification of ScMreB5 be at 4 °C or in ice. For affinity purification, one can use the manual setup in a cold room. It is important to run SDS-PAGE gel after every step of purification to ensure that only fractions that have a single band corresponding to ScMreB5 are taken forward. Before setting up crystallization, it is advisable to perform an ATPase assay to ensure that the protein is catalytically active. The protocol for the ATPase assay is available in the original publication.
This protocol can serve as a reference for purifying other ScMreB5 orthologs. Heterologous expression of MreB5 in E. coli could be toxic for the cells due to the presence of native E. coli MreB. It is recommended to optimize protein expression in various E. coli strains suitable for toxic heterologous proteins. Protein insolubility may be encountered while performing expression check, for which mild detergents can be used in the lysis buffer.
During purification, issues like low protein yield due to protein precipitation may arise. The thermal shift assay can be performed to screen for optimal conditions (salts, buffers, detergents, ligands). The condition with the highest melting temperature can be selected as the optimized condition for further purification trials. Size exclusion chromatography is recommended to assess the polymeric state of the protein, followed by performing crystallography with monomeric protein.
The protocol might be applicable for standardizing other MreBs as well as other proteins that often get stabilized upon the addition of a ligand, most often either the substrate or the product in the case of enzymes. Ligand-induced stability helps us to obtain sufficient protein for carrying out the stability assays using thermal shift. This could lead us to identify other stabilizing additives, which can be used for purifying the protein in the absence of the initial ligand. The presence of excess stabilizing ligand, such as the substrate of the product, precludes us from performing many biochemical experiments such as ligand affinity studies as well as enzyme kinetic assays. Hence, the identification of conditions that stabilize the protein without the excess substrate/ligand is highly beneficial.
Acknowledgments
We thank Saket R. Bagde, Nivedita Mitra and Shrikant Harne for their contributions to the study of ScMreB5 characterization. We thank the macromolecular crystallography facility at IISER Pune and synchrotron facilities at European Synchrotron Radiation Facility (ESRF), Grenoble, Diamond Light Source (MX22637) and Department of Biotechnology India for facilitating data collection at ID29, ESRF. We thank the IISER Pune for the infrastructure to perform experiments. P. Gayathri acknowledges the support from the following funding agencies: Department of Science and Technology INSPIRE Faculty Fellowship (IFA12/LSBM-52), Innovative Young Biotechnologist Award (BT/07/IYBA/2013) and Department of Biotechnology Membrane Structural Biology Program grant (BT/PR28833/BRB/10/1705/2018) and IISER Pune. V. Pande thanks IISER Pune for the fellowship and the Infosys Travel award. Graphical overview created with BioRender.com.
Competing interests
The authors declare that they have no competing interests.
References
Wagstaff, J. and Löwe, J. (2018). Prokaryotic cytoskeletons: protein filaments organizing small cells. Nat Rev Microbiol. 16(4): 187–201.
Kawai, Y., Asai, K. and Errington, J. (2009). Partial functional redundancy of MreB isoforms, MreB, Mbl and MreBH, in cell morphogenesis of Bacillus subtilis. Mol Microbiol. 73(4): 719–731.
Kruse, T., Bork‐Jensen, J. and Gerdes, K. (2004). The morphogenetic MreBCD proteins of Escherichia coli form an essential membrane‐bound complex. Mol Microbiol. 55(1): 78–89.
Errington, J. (2015). Bacterial morphogenesis and the enigmatic MreB helix. Nat Rev Microbiol. 13(4): 241–248.
van den Ent, F., Izoré, T., Bharat, T. A., Johnson, C. M. and Löwe, J. (2014). Bacterial actin MreB forms antiparallel double filaments. eLife. 3: e02634.
Salje, J., van den Ent, F., de Boer, P. and Löwe, J. (2011). Direct Membrane Binding by Bacterial Actin MreB. Mol Cell. 43(3): 478–487.
Ursell, T. S., Nguyen, J., Monds, R. D., Colavin, A., Billings, G., Ouzounov, N., Gitai, Z., Shaevitz, J. W. and Huang, K. C. (2014). Rod-like bacterial shape is maintained by feedback between cell curvature and cytoskeletal localization. Proc Natl Acad Sci USA. 111(11): E1025–E1034.
Harne, S., Duret, S., Pande, V., Bapat, M., Béven, L. and Gayathri, P. (2020). MreB5 Is a Determinant of Rod-to-Helical Transition in the Cell-Wall-less Bacterium Spiroplasma. Curr Biol. 30(23): 4753–4762.e7.
Pande, V., Mitra, N., Bagde, S. R., Srinivasan, R. and Gayathri, P. (2022). Filament organization of the bacterial actin MreB is dependent on the nucleotide state. J Cell Biol. 221(5): e202106092.
Citti, C., Maréchal-Drouard, L., Saillard, C., Weil, J. H. and Bové, J. M. (1992). Spiroplasma citri UGG and UGA tryptophan codons: sequence of the two tryptophanyl-tRNAs and organization of the corresponding genes. J Bacteriol. 174(20): 6471–6478.
van den Ent, F. and Löwe, J. (2006). RF cloning: A restriction-free method for inserting target genes into plasmids. J Biochem Bioph Methods. 67(1): 67–74.
Cimmperman, P. and Matulis, D. (2011). Protein Thermal Denaturation Measurements via a Fluorescent Dye. In: Podjarny, A., Dejaegere, A. P. and Kieffer, B. (Eds.). Biophysical Approaches Determining Ligand Binding to Biomolecular Targets: Detection, Measurement and Modelling. The Royal Society of Chemistry, p.0.
Kozak, S., Lercher, L., Karanth, M. N., Meijers, R., Carlomagno, T. and Boivin, S. (2016). Optimization of protein samples for NMR using thermal shift assays. J Biomol NMR. 64(4): 281–289.
van den Ent, F., Amos, L. A. and Löwe, J. (2001). Prokaryotic origin of the actin cytoskeleton. Nature. 413(6851): 39–44.
Takahashi, D., Fujiwara, I., Sasajima, Y., Narita, A., Imada, K. and Miyata, M. (2022). ATP-dependent polymerization dynamics of bacterial actin proteins involved in Spiroplasma swimming. Open Biol. 12(10): e220083.
Article Information
Publication history
Received: Apr 18, 2024
Accepted: Aug 23, 2024
Available online: Sep 28, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Quantitative Analysis of Fish Morphology Through Landmark and Outline-based Geometric Morphometrics with Free Software
DL Du Luo
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5087 Views: 411
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Abstract
Morphology underpins key biological and evolutionary processes that remain elusive. This is in part due to the limitations in robustly and quantitatively analyzing shapes within and between groups in an unbiased and high-throughput manner. Geometric morphometrics (GM) has emerged as a widely employed technique for studying shape variation in biology and evolution. This study presents a comprehensive workflow for conducting geometric morphometric analysis of fish morphology. The step-by-step manual provides detailed instructions for using popular free software, such as the TPS series, MorphoJ, ImageJ, and R, to carry out generalized Procrustes analysis (GPA), principal component analysis (PCA), discriminant function analysis (DFA), canonical variate analysis (CVA), mean shape analysis, and thin plate spline analysis (TPS). The Momocs package in R is specifically utilized for in-depth analysis of fish outlines. In addition, selected functions from the dplyr package are used to assist in the analysis. The full process of fish outline analysis is covered, including extracting outline coordinates, converting and scaling data, defining landmarks, creating data objects, analyzing outline differences, and visualizing results. In conclusion, the current protocol compiles a detailed method for evaluating fish shape variation based on landmarks and outlines. As the field of GM continues to evolve and related software develops rapidly, the limitations associated with morphological analysis of fish are expected to decrease. Interoperable data formats and analytical methods may facilitate the sharing of morphological data and help resolve related scientific problems. The convenience of this protocol allows for fast and effective morphological analysis. Furthermore, this detailed protocol could be adapted to assess image-based differences across a broader range of species or to analyze morphological data of the same species from different origins.
Key features
• This protocol provides a comprehensive set of commonly used GM-analyzing methods and visualizing skills plus supporting information to help assess the appropriate analysis method
• By incorporating both landmarks and outlines, this protocol facilitates a thorough analysis of two-dimensional shape variation in fish, covering a wide range of morphological features
• The simplified workflow and detailed procedures make it accessible for non-experienced users to successfully complete the analysis while also providing valuable insights for experienced users
Keywords: Morphological variation Landmarking Background removal Outline extraction Digitization Interpretation
Graphical overview
Workflow for conducting geometric morphometrics analysis on fish. The steps include image acquisition as data sources, digitization of fish morphology using landmark-based methods, analysis of shape variation characteristics, and visualization of the results in relation to biological interpretation. Largemouth bass (Micropterus salmoides) is used as an example in the schematic representation.
Background
Morphology has long been recognized as a fundamental trait in the field of biology. The intricate relationship between morphogenetic and evolutionary factors, as well as morphospaces, underscores the need for multivariate methods in biological and ecological research [1]. Geometric morphometrics (GM) has emerged as a widely utilized approach for the quantitative analysis of shape variation, particularly in the domains of biology and anthropology. It has become one of the primary methods for assessing essential morphological variables because it provides a quantitative/unbiased approach and morphological comparison [2]. GM has been instrumental in addressing a diverse array of questions related to morphological variation, including population differences, developmental patterns, responses to environmental factors, evolutionary trends, and functional morphology. Within the realm of morphology, three styles of morphometrics are commonly employed: traditional morphometrics, landmark-based GM, and outline-based GM [3]. In general, GM encompasses the following analytical steps: data acquisition, morphological variation analysis, results visualization, and interpretation.
For novice practitioners, the intricacy of GM often arises from the complexities associated with data construction and transformation, as well as the diverse array of analyzing methods available. Throughout the process, landmarks serve as the foundation for quantifying shape. GM analysis can be conducted using distinct landmark configurations for landmarks and semi-landmarks [4]. Traditionally, there are three types of landmarks [5]. However, as GM has advanced, the classification system for landmarks has evolved, with a more convenient typology being utilized in applied studies [6,7]. In the updated typology, the conventional roster of landmark points can be categorized into six types, intended to supersede the three types outlined in Bookstein [5]. This new classification corresponds to the different operational origins of the points along the curve or curves on which they are situated [6]. The process of landmarking and classifying landmarks may heavily rely on biological interpretation, and the major limitation of these types of techniques is that the labeling-analysis processes are all semi-manual or manual. Nevertheless, landmarking analysis remains the primary technique used in GM to this day. The characteristics of landmarks are described below.
Type I landmarks (anatomical landmarks)
Definition:
Points of clear biological or anatomical significance that can be precisely and consistently identified across all specimens.
These landmarks correspond to specific, discrete, and easily recognizable anatomical features.
Examples:
The tip of the nose.
The corner of the eye.
The junction between bones.
Advantages:
High reliability and repeatability.
Easily comparable across specimens due to clear homology.
Applications:
Frequently used in studies of skeletal morphology and other well-defined anatomical structures.
Type II landmarks (mathematical landmarks)
Definition:
Points defined by geometric properties such as maxima or minima of curvature, or points where certain geometric properties change.
These landmarks may not correspond to specific anatomical features but are identified based on their geometric properties.
Examples:
The point of maximum curvature along a bone.
The deepest point in a notch.
Advantages:
Useful for capturing shape information where anatomical landmarks are not clearly defined.
Can provide additional geometric context to the shape.
Applications:
Often used in conjunction with Type I landmarks to provide a more comprehensive shape analysis.
Type III landmarks (constructed landmarks)
Definition:
Points defined by their relative position or constructed based on other landmarks.
These landmarks are not associated with specific anatomical features but are placed based on their geometric relationship to other landmarks.
Examples:
The midpoint between two anatomical landmarks.
Points evenly spaced along a curve or surface.
Advantages:
Flexible and can be used to outline complex shapes.
Useful in capturing the overall geometry of a structure.
Applications:
Frequently used in semi-landmark analysis to capture the shape of curves and surfaces where fixed landmarks are insufficient.
Procrustes superimposition serves as the foundational step for subsequent analysis in GM [8]. Biologists have grappled with aligning the method of "Cartesian transformations" and "transformation grid" with geometric patterns since its original exposition [9]. Three decades ago, the concept of "morphometric synthesis" emerged, combining Procrustes shape coordinates with thin-plate spline (TPS) renderings for various multivariate statistical comparisons [10]. However, a concluding discussion suggests that the current toolkit of GM, centered on Procrustes shape coordinates and TPS, may be too limited to accommodate the interpretive needs of evolutionary and developmental biology [11]. Common methods used to identify major modes of shape variation and determine group differences include principal component analysis (PCA), TPS, discriminant function analysis (DFA), partial least squares (PLS), and canonical variate analysis (CVA). The interpretation of these methods is based on biological questions or hypotheses, combining patterns of shape variation, key landmarks or curves, and findings with relevant evolutionary or ecological factors.
GM has experienced a surge in applications within evolutionary biology and ecology, particularly with the use of three-dimensional imaging data [12]. However, the majority of studies involving fish morphology are based on two-dimensional data. GM analysis in fisheries primarily focuses on species taxonomy, group diversity, individual development and evolution, and ecomorphological variation [13]. The development of software and the reduction of technical limitations in analysis may enhance fish research. GM has evolved alongside advancements in theory and technology, resulting in a variety of software and analysis methods. There are large pre-existing datasets of fish images that can be analyzed using appropriate GM methods. However, several analysis techniques are often required for a single research project, which can present challenges for novices. Consequently, this protocol aims to compile a comprehensive set of methods for conducting GM analysis on fish using two-dimensional data. Although there are no established standards for performing GM analysis, advancements in technology are crucial for characterizing variations in fish body shape. Furthermore, it is anticipated that this approach will facilitate the sharing of morphological data and help resolve related scientific problems. [14], potentially expediting scientific advancements in the field of fish biology and ecology [15].
Software and datasets
Software:
tpsDig2 Version 2.32 (https://www.sbmorphometrics.org/soft-dataacq.html, accessed January 31, 2023)
tpsUtil Version 1.82 (https://www.sbmorphometrics.org/soft-tps.html, accessed January 31, 2023)
tpsRelw Version 1.75 (https://www.sbmorphometrics.org/soft-utility.html, accessed January 31, 2023)
ImageJ 1.54i (https://imagej.net/ij/download.html, accessed March 13, 2023) [16]
MorphoJ Version 1.08.01 (https://morphometrics.uk/MorphoJ_page.html)
R programs (https://cran.r-project.org)
RStudio (https://posit.co/)
R package of Momocs (https://cran.r-project.org/web/packages/Momocs/index.html)
This protocol is running on Windows 11 (64-bit). Taking into account the compatibility of the operating system, the software tutorial, and the possible requirement of preinstalling a version of Java, please download and install all the necessary software: tpsUtil version 1.82, tpsDig2 version 2.32, tpsRelw version 1.75 [17], MorphoJ version 1.08.01 [18], R version 4.3.2 [19], and RStudio 2023.09.1 [20].
Website
Pixelcut (https://create.pixelcut.ai/background-remover)
Photoroom (https://www.photoroom.com/tools/background-remover)
The two AI-based background-remover tools are used to extract fish by removing the image background.
Digitized images
Four groups of largemouth bass (Micropterus salmoides) images were utilized in this study. Two groups were obtained by photographing fish cultured in farm ponds located in Foshan city, while the other two groups of images were sourced from the internet. When photographing the fish, the digital camera was fixed in position with the lens perpendicular to the ground. The fish was placed horizontally on a solid-colored background directly beneath the camera. If necessary, soft materials were used to adjust the position of the fish to ensure its body axis was horizontal and the head was facing left. The photos were taken in macro mode after focusing and were stored in .jpeg format. The size of the photos depended on the camera's capabilities, with sizes between 2 and 10 MB considered appropriate. The internet images were sourced from Microsoft Bing Images (https://cn.bing.com/images/feed?form=Z9LH, accessed April 4, 2024) and Google Images (https://www.google.com.hk/imghp?hl=en&ogbl, accessed April 4, 2024) using the query terms “largemouth bass,” “Micropterus salmoides,” and “largemouth bass (Micropterus salmoides)”. All images used in the current experiment included fish with sufficient resolution, showing a normal appearance and an integrated outline in left/right lateral views [21]. For self-captured JPEG digital photographs, the file size was greater than 2266 KB. The internet-sourced images, whether in .jpg, .png, or other file formats, were converted to .jpeg (.jpg) format, with a minimum size of 14 KB used in the present research. Finally, four groups of mature largemouth bass images were included in this research: LB-FF-FF (feeding with frozen bait, n = 44), LB-FF-FS (feeding with artificial feed, n = 42), LB-IN-DH (internet-sourced realistic painting, n = 23), and LB-IN-PH (internet-sourced picture, n = 30).
Procedure
The procedure for conducting GM primarily involves the following steps: data collection, landmark placement, digitization, Procrustes superimposition, shape analysis, statistical testing, visualization, interpretation, and biological inference.
Digitization of fish image data through landmarking and file format conversion
Image preparation
Typically, original images with scale in .jpeg format can be used for landmarking. To maintain consistency, the background of the images was first removed using an AI-based online tool, and then the images were used for placing landmarks and extracting outlines.
The two online tools (Pixelcut and Photoroom, accessed April 7, 2024) are both image background removers that offer the free function of downloading background-removed images at standard resolution. The resolution meets the needs of the following analyses. By default, the background is set to be transparent, and the image data quality is determined by the original image.
For Pixelcut: Open the website → Upload image → Download → Download standard quality (1,080 × 720 px).
For Photoroom: Open the website → Start from a photo → Download → Standard resolution (1,280 × 1 280 px).
Taking the fish images from the first group (LB-FF-FF) as example data, the background-removed images are collected and transferred to a designated location (F:\Directory\Images\LB-FF-FF). If the images are not easily distinguishable from one another prior to landmarking, it is best to rename the images.
Landmark placement
Fourteen landmarks (Figure 1) were digitized using tpsDig2 software from each specimen's image [22]. It is recommended that the total number of landmarks should be less than half or a third of the number of individuals [23]. Prior to landmarking, a .tps file should be constructed using tpsUtil. This file serves as a link to manipulate all the image files within a specific group or classification.
Open tpsUtil → Operation → Build tps file from images → Input directory → Input → Click any one image and then click open → Output file → Name the output file with .tps as extensions (LB-FF-FF.tps) → Actions → Setup → Check images with Actions → Select Include path → Create → Close.
Landmarking using tpsDig2 and recording the landmarks with x- and y-coordinates in .tps file (Figure 2).
Open tpsDig2 → File → Input source → File → .tps file (LB-FF-FF.tps) → Open → Digitalize landmarks → Save data to TPS file → (LB-FF-FF.tps) → Save → Overwrite (Supplemental File 1. LB-FF-FF.tps).
Figure 1. Landmarks placement of largemouth bass used in this study. The definition of landmarks depends on the biological interpretation specific to each research scope.
Figure 2. Process of landmark placing using software tpsDig2. A. Input source .tps file. B. Save data to .tps file after finishing landmarking.
Set scale according to measurement methodology.
Scale image one by one
Open tpsDig2 → Image edit tools → Measure → Scale factor → Reference length (1-Centimeters) → Set scale → Draw a line with the same reference length → OK → Back to tpsDig2 window → Save data to TPS file → (LB-FF-FF.tps) → Save → Overwrite.
Scale as a variable for all images
Open tpsUtil → Operation → Add variable → Input file → Input → Click (LB-FF-FF.tps) and open → Output file → Name the output file with .tps as extensions (LB-FF-FF _allscale.tps) → Actions → Setup → Fill Variable keyword (Scale) → Fill Value → Create → Close.
Compile files (.tps)
If the images are not landmarked together with all the required data in one document, the .tps files should first be collected in one folder and then compiled into a single file by appending them.
Open tpsUtil → Operation → Append files → Input file → Input → Click (Group1.tps) and open → Output file → Name the output file with .tps as extensions (Expreiment1-Appendgroupfile.tps) → Actions → Setup → Tick .tps files → Create → Close.
The format of .tps file in GM
In GM, .tps files are commonly used to store landmark data. These files contain both landmark coordinates and additional information about the specimen. The .tps file format was first developed by Fred L. Bookstein in the 1990s. To check the contents of a .tps file, you can open it as a .txt document.
The general format of a TPS file is as follows:
LM= 14
19.00000 170.00000
81.00000 206.00000
616.00000 217.00000
…
774.00000 155.00000
IMAGE= F:\Directory\Images\ LB-FF-FF \IMG_2852.jpg
ID=0
VARIABLES=Scale=0.018155
…
A .tps file consists of two parts: a header (LM = 14) and a data section. The header provides information about the number of landmarks, while the data section contains the actual landmark coordinates. Each line in the data section represents a single landmark with its x-coordinate and y-coordinate. Additionally, other information such as image ID, image directory, specimen ID, and scale can be included at the beginning or end of each specimen's data.
Convert .tps format file to .nts format for use in MorphoJ
Open tpsUtil → Operation → Convert tps/nts coordinates file → Input file → Input → Click (LB-FF-FF_allscale.tps) and open → Output file → Name the output file with .nts as extensions (LB-FF-FF_allscale.nts) → Actions → Create → (Tick “use scale factor” [this will convert the pixel coordinates into standard units of measurements], 2D landmarks and Image name) → Create → Close.
The format of .nts file in GM
The .nts (Numerical Taxonomic System) file format is a legacy format used for storing landmark data, developed by Fred Bookstein in the early 1980s. .nts files can be created and edited using various software programs, including MorphoJ, tpsDig, and tpsRelw.
The general format of a .nts file is as follows:
1 44L 28L 0
IMG_2852.jpg IMG_2854.jpg…IMG_2976.jpg
X1 Y1 X2 Y2…X14 Y14
176.00000 384.00000 223.00000 419.00000 … 854.00000 376.00000
…
The parameters in the .nts file must adhere to specific guidelines. The first code in the parameter line should be "1". The second parameter, 44L, denotes the number of specimens, while the third parameter, 28L, represents the total count of x and y coordinates for each specimen. The fourth code in the parameter line is "0,” indicating the absence of missing data; if missing data exists, this code should be "1." It's important to note that some editing or removing redundant blank spaces may be necessary if a file does not solely consist of a single data matrix.
Shape variation analysis and visualization with MorphoJ
Create project and dataset [24]
File → New Project → Name for the new project (Project-images1) → Dimensionality of the data (2 dimensions) → Object symmetry? (no) → Name for the new dataset (Dataset- LB-FF-FF) → File type (NTSYSpc) → Add documents (LB-FF-FF_allscale.nts) → Create Dataset (Figure 3A) → File → Save Project → Name the Project (Project-LMB1.morphoj)
Figure 3. Creating a project and extracting a classifier in MorphoJ software. A. Creating a project and dataset. B. Extracting the classifier from ID strings by counting positively from left to right.
Extract and add classifiers
Preliminaries → Extract new classifier from ID strings → Name for a new classifier (fish_source) → First character (21) → Last character (28) → Execute (Figure 3B).
Repeat the above steps to add more classifiers and the added classifiers can be edited from: Preliminaries → Edit classifiers (or directly click Graphics).
Outliers check
Preliminaries → Find outliers.
Repeat the above steps to create other datasets
Procrustes fit
MorphoJ uses a full Procrustes fit. For most circumstances, there is very little difference from generalized Procrustes analysis (GPA).
Preliminaries → New Procrustes Fit → Select Align by principal axes → Perform Procrustes Fit.
Export dataset for use in other software
Project Tree → Click Dataset (Dataset-images1.NTS) (Dataset-LB-FF-FF) → Click File in menu → Export Dataset → Select Data types and Classifiers (click with Ctrl or Shift for more than one type) → Name the Dataset as .txt file (Dataset-images1.NTS.txt) → Save.
Principal component analysis (PCA)
Purpose: Reduces the dimensionality of the shape data and identifies the main axes of variation.
Output: Principal components (PCs) that describe the major patterns of shape variation, and scatter plots of specimens in PC space.
Preliminaries → Generate Covariance Matrix → Selected dataset (Dataset-images1.NTS) → Execute → Variation → PCA → Results (without ticking the box “Pooled within-group covariances.”).
Eigenvalues → Variance explained by each PC.
PC scores → Visualization of individuals in the shape space.
Combine datasets
Click on the dataset in the Project Tree → Preliminaries → Combine Datasets → Name for the new dataset → Select the Start dataset and other datasets → Execute.
Discriminant function analysis (DFA)
Purpose: Classifies specimens into predefined groups based on shape.
Output: Discriminant functions, classification accuracy, scatter plots of specimens in discriminant space.
Click on the combined dataset → Comparison menu → Discriminant Function → Name for the discriminant function analysis → Select Dataset and Datatype (combined dataset) → Classifier to be used as grouping criterion (fish_source) → Select Pairs of groups to be included (or tick “Include all pairs of groups”) → Permutation runs:1000 → Execute.
Canonical variate analysis (CVA)
Purpose: Maximizes separation between predefined groups (e.g., species, populations) to find axes that best differentiate the groups.
Output: Canonical variates (CVs), scatter plots showing group separation (Figure 4), classification rates.
Click the combined dataset → Comparison menu → Canonical Variate Analysis → Name the CVA → Select the Dataset and Data type → Select classifier variable to use for grouping (fish_source) → Tick Permutation tests → Number of iterations = 10,000 → Execute.
Figure 4. Scatterplot of the CV scores showing the shape features that best distinguish multiple groups of specimens. The color of points represents different specimen groups.
Visualization of DFA shape difference combined with outline data
Extract fish outline
Open ImageJ → File → Open → Select the background removed image → Open → Image → 8-bt → Image → Adjust → Brightness/Contrast (Figure 5A) → Adjust the minimum and maximum value → Apply → Apply Lookup Table → OK → Process → Binary → Make Binary → Process → Binary → Fill holes → Process → Binary → Outline → Wand (tracing) tool → Click the outline of the fish → File → Save As → Jpeg → Save.
Figure 5. Extracting an outline and creating an outline file for use in MorphoJ software. A. Adjusting brightness and contrast while extracting the outline using ImageJ software. B. Creating the outline file through the outline object tool and saving it as xy coordinates using tpsDig2 software.
Create outline file.
Open tpsDIG2 → File → Input source → File → Select the outline image → Open → Outline object → Right click to save as “Save as XY coords.” (Figure 5B) → File → Save just outlines → Save the outline as .txt files → Copy the coords. to Excel or other table processing software → Insert landmarks before the coodrs. → Insert a column to number the landmarks and coords. (such as “0” and “1,” respectively) → Copy the data back to .txt file to create a file.
Import outline file.
Click the dataset → File → Import Outline File → Name for the outline → Select the outline file → Open → Jump to Graphics of DFA → Shape difference → Right click → Change the type of graph → Warped Outline Drawing (Figure 6).
Figure 6. Visualization of discriminant function analysis (DFA) shape difference. Combining with outline data, the shape differences show general changes from the first to the second group. Lollipop graph and transformation grid graph can also be used to display specific variations.
Outline analysis and visualization with the Momocs R package
The analysis of semi-landmark data follows a procedure similar to that previously described. The outlined data analysis is subjected to the following steps using the Momocs R package [25].
### Starting
setwd("F:/Directory/GM-momocsu/IMGS") # Working directory
library(Momocs)
### Collection of outline coordinates data
## Extract outline coordinates from image files
lf1 <- list.files('F:/Directory/GM-momocsu/IMGS/LB-FF-FF-FIJI', full.names = TRUE)
lf1
coo1 <- import_jpg(lf1) # Data storage directory/Coo1
## Export each outline coordinates as a .csv file
# For coo1: Data storage directory/Coo1
setwd("F:/Directory/GM-momocsu/IMGS/outputdata/coo1_ff")
coo_list <- lapply(1:length(coo1), function(i) {
df <- data.frame(
x = coo1[[i]][, 1],
y = coo1[[i]][, 2],
image = basename(lf1[i])
)
return(df)
})
#
for (i in seq_along(coo_list)) {
filename <- paste0("outline_", i, ".csv")
write.csv(coo_list[[i]], file = filename, row.names = FALSE)
}
#
## Import and combine individual data files
library(dplyr)
imported_files1 <- lapply(paste0("outline_", seq_len(length(lf1)), ".csv"), read.csv)
combined_df1 <- bind_rows(imported_files1)
## Export as a combined .csv file
write.csv(combined_df1, file = "combined_outlines1.csv", row.names = FALSE) # (Supplemental File 2. combined_outlines1.csv).
### Loading data, converting to an Out object and inspecting
setwd("F:/Directory/GM-momocsu/IMGS/outputdata/coo1_ff")
## Read the .csv file
coo1_ff <- read.csv("combined_outlines1.csv")
# Convert the data frame to a list of matrices
coo1_list <- lapply(split(coo1_ff[, c("x", "y")], coo1_ff$image), as.matrix)
## Convert the list of matrices to an Out object
out1 <- Out(coo1_list)
## Plots outlines for inspection
# Panels of outlines (Figure 7)
panel(out1, c(6, 8),names=TRUE, cex.names=0.5)
# Plot one of the Out
coo_plot(out1[3])
Figure 7. Outlines of largemouth bass from one experimental group. The panel of outlines can be used for inspecting the extraction.
## Check if the outline is closed
coo_is_closed(out1)
### Outline data normalization
## Outline smoothing
# Extract coordinates from the Out object
coo_list_out1 <- lapply(out1$coo, function(coo) as.matrix(coo))
# Apply smoothing to each set of coordinates
smoothed_coo_list_out1 <- lapply(coo_list_out1, function(coo) coo_smooth(coo, n = 5))
# Convert the smoothed coordinates back to an Out object
smoothed_out1 <- Out(smoothed_coo_list_out1)
# Check for one of the smoothed outline
coo_plot(smoothed_out1[2], main = "iterations=5")
## Centering and scaling the outlines
# Center each set of coordinates
centered_matrices_out1 <- lapply(coo_list_out1, coo_center)
# Scale each set of coordinates
scaled_matrices_out1 <- lapply(centered_matrices_out1, coo_scale, scale = 1)
# Reconstruct the Out object
centered_scaled_out1 <- Out(scaled_matrices_out1)
# Check for centering and scaling
stack(centered_scaled_out1)
## Sampling pseudo-landmarks and reversing anticlockwise coordinates
# Sampling pseudo-landmarks
out1_resample <- coo_sample(centered_scaled_out1, 1000)
# Test if all shapes are developing consistently clockwise
coo_likely_clockwise(out1_resample)
# If not, the coordinates can be reversed
out1_resample_rev <- out1_resample
# Prepare Coo data for reversing
out1_rev_coo <- out1_resample_rev$coo
# Iterate over each outline
for (i in seq_along(out1_rev_coo)) {
# Extract the coordinates for this outline
coords <- out1_rev_coo[[i]]
# Check if the outline is likely not clockwise
if (!coo_likely_clockwise(coords)) {
# Reverse the coordinates
reversed_coords <- coo_rev(coords)
# Update the outline within the list
out1_rev_coo[[i]] <- reversed_coords
}
}
#
# Check if all outlines are clockwise
out1_rev_coor <- Out(out1_rev_coo)
coo_likely_clockwise(out1_rev_coor) # All TRUE
### Adjust data by defining landmarks, sliding coordinates to create Coe objects
## Define landmarks on the sampled-reversed outlines
out1_rev_coor_ldk <- def_ldk(out1_rev_coor,4)
## Sliding coordinates
out1_rev_coor_ldks <- coo_slide(out1_rev_coor_ldk,ldk = 1)
## Procrustes superimposition
out1_rev_coor_ldks_fgp <- fgProcrustes(out1_rev_coor_ldks)
## Creation Coe objects by Elliptical Fourier transform
out1_rev_coor_ldks_fgp_ef <- efourier(out1_rev_coor_ldks_fgp, 12, norm = FALSE) # In some cases, Elliptical Fourier Transform may address the limitations of traditional methods [26].
### Principal component analysis (PCA) and successive statistics
## PCA for the group out1
out1_rev_coor_ldks_fgp_ef_pca <- PCA(out1_rev_coor_ldks_fgp_ef)
summary(out1_rev_coor_ldks_fgp_ef_pca)
boxplot(out1_rev_coor_ldks_fgp_ef_pca, fac = NULL, nax = 1:12)
scree_plot(out1_rev_coor_ldks_fgp_ef_pca)
## For PCA of combined outlines
# Repeating the previous steps to develop data for other groups
# Combining group outlines
outall_rev_coor_ldks_fgp <-combine(out1_rev_coor_ldks_fgp,
out2_rev_coor_ldks_fgp,
out3_rev_coor_ldks_fgp,
out4_rev_coor_ldks_fgp)
# Creating Coe objects
outall_rev_coor_ldks_fgp_ef <- efourier(outall_rev_coor_ldks_fgp, 12, norm = FALSE)
# PCA and visualization
outall_rev_coor_ldks_fgp_ef_pca <- PCA(outall_rev_coor_ldks_fgp_ef)
summary(outall_rev_coor_ldks_fgp_ef_pca)
boxplot(outall_rev_coor_ldks_fgp_ef_pca, fac = NULL, nax = 1:12)
scree_plot(outall_rev_coor_ldks_fgp_ef_pca) # (Figure 8)
Figure 8. Principal component analysis (PCA) shows the proportion of each component for combined outlines. The number on top of each column of the bar chart represents the accumulated proportion of the 24 components.
### Mean shape analysis and Thin plate spline (TPS) analysis
## Mean shape analysis and visualization
out1_rev_coor_ldk_fgps_ef_mean <- (MSHAPES((out1_rev_coor_ldk_fgps_ef), fac = NULL, FUN = mean, nb.pts = 1000))
# Visualization of mean shape
coo_plot(out1_rev_coor_ldk_fgps_ef_mean)
# Visualization of mean shape and group outlines (Figure 9)
out1_rev_coor_ldks_fgp %>% coo_center %>% stack
coo_draw(out1_rev_coor_ldks_fgp_ef_mean,border='forestgreen',lwd = 3)
# Visualization of mean shapes differences
coo_arrows(out1_rev_coor_ldks_fgp_ef_mean,
out2_rev_coor_ldks_fgp_ef_mean,
length = coo_centsize(out1_c)/2, angle = 20, code = 2)
coo_draw(out2_rev_coor_ldks_fgp_ef_mean, border = "blue", centroid = T,
first.point = F, zoom = 0.7, lwd = 2)
Figure 9. Visualization of the mean shape and stacked outlines from the experimental Group 1. The green line represents the mean shape of the outlines from Group 1, while the grey lines represent the individual outlines from Group 1.
## Thin plate spline analysis (TPS) (Figure 10)
tps_grid(out1_rev_coor_ldks_fgp_ef_mean,
out3_rev_coor_ldks_fgp_ef_mean, over = 1.2,
amp=1, grid.size=20,shp.border = c("red", "blue"),
shp.lwd = c(2, 2),legend = F)
Figure 10. Visualization of the mean shape difference between experimental Group 1 and Group 3 through thin plate spline (TPS) analysis. The red line represents the mean shape of outlines for Group 1, while the blue line represents the mean shape of outlines for Group 3.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Santos et al. [27]. Geometric morphometrics as a tool to identify species in multispecific flatfish landings in the Tropical Southwestern Atlantic. Fish Res. (Figure 2, 3).
Caillon et al. [28]. A morphometric dive into fish diversity. Ecosphere. (Figure 1, 3).
Rabe et al. [29]. Geometric morphometric analysis of an ontogenetic cranial series of the Permian dicynodont Diictodon feliceps. Proc R Soc B. (Figure 2).
General notes and troubleshooting
Based on experience from the current research, the size of an image used for outline extraction should be no smaller than 14 KB.
For research requiring scale, landmarking should be performed for measurements before removing the image background.
If a landmark is misplaced, all subsequent landmarks in that image must be crossed out. The researcher needs to correct the misplacement by re-establishing the landmarks from scratch.
For images of fish without an original scale, real measurements cannot be accomplished. In the methods section for setting the scale for measurement, the scale can be set to “scale=1.”
Based on the current versions of tpsDig2 (Version 2.32) and tpsUtil (Version 1.82), the file path is included in the name of each image during the process of creating the .tps file and placing landmarks. Sometimes, the image name in the .tps file may be presented as its original name. Therefore, prior to landmarking, images from each group should be appropriately named to differentiate them from one another and to provide sufficient information regarding their biological characteristics. For example, the images can be named LB_Foshan_group1_female_mature_bait1_IMG_1001.jpeg. The names can be shortened as necessary by abbreviating words within the name.
The xy coordinates of the outline can also be created using ImageJ software, though the number of coordinates is typically much less than that generated through tpsDig2.
For outline analysis, reversing the anticlockwise coordinates and defining landmarks to create Coe objects are two key steps for samples with significant shape variations among images in a group.
When applying this protocol to GM analyses in other species, landmarking, outline extraction, and the selection of analysis methods should correlate with biological interpretation.
Acknowledgments
The author thanks Dr. Tingwei Zhang for assisting in taking photos of fish. This work was supported by the Central Public-interest Scientific Institution Basal Research Fund, CAFS (2024SJRC9), the Project of Innovation Team of Survey and Assessment of the Pearl River Fishery Resources (2023TD10) and the National Natural Science Foundation of China (31600446).
Competing interests
The author declares no competing interests.
Ethical considerations
All animal study protocols were approved by the Laboratory Animal Ethics Committee of Pearl River Fisheries Research Institute, CAFS (LAEC-PRFRI-20160323).
References
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Dwivedi, A. K. and De, K. (2023). Role of Morphometrics in Fish Diversity Assessment: Status, Challenges and Future Prospects. Natl Acad Sci Lett. 47(2): 123–126. https://doi.org/10.1007/s40009-023-01323-x
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Collyer, M. L., Davis, M. A. and Adams, D. C. (2020). Making Heads or Tails of Combined Landmark Configurations in Geometric Morphometric Data. Evol Biol. 47(3): 193–205. https://doi.org/10.1007/s11692-020-09503-z
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Wärmländer, S. K. T. S., Garvin, H., Guyomarc'h, P., Petaros, A. and Sholts, S. B. (2018). Landmark Typology in Applied Morphometrics Studies: What's the Point?. Anat Rec. 302(7): 1144–1153. https://doi.org/10.1002/ar.24005
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Thompson, D. A. (1917). On growth and form. Abridged and edited by j. T. Bonner, Cambridge University Press, 1961. Cambridge University Press, Cambridge.
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Bookstein, F. L. (2023). Reworking Geometric Morphometrics into a Methodology of Transformation Grids. Evol Biol. 50(3): 275–299. https://doi.org/10.1007/s11692-023-09607-2
Hu, H., Bjarnason, A. and Benson, R. (2021). 3D geometric morphometric protocol - quantifying the morphology of living and extinct vertebrates. Bio-101: e1010664. https://doi.org/10.21769/BioProtoc.1010664
Wang, C., Fang, Z. and Chen, X. J. (2022). Advances in the application of biometrics-based geometric morphometrics in fisheries. Marine Fisheries. 44 (1): 112–128. https://doi.org/10.13233/j.cnki.mar.fish.2022.01.004
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Supplementary information
The following supporting information can be downloaded here:
Supplemental File 1. LB-FF-FF.tps
Supplemental File 2. combined_outlines1.csv
Article Information
Publication history
Received: Jun 12, 2024
Accepted: Aug 25, 2024
Available online: Sep 13, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Environmental science > Marine vertebrates
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5,088 | https://bio-protocol.org/en/bpdetail?id=5088&type=0 | # Bio-Protocol Content
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Chemogenetic Silencing of Neonatal Spontaneous Activity of Projection Neurons in the Dorsal Striatum of Mice
BK Bojana Kokinovic *
MR Maria Ryazantseva *
SM Svetlana Molchanova
(*contributed equally to this work)
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5088 Views: 268
Reviewed by: Oneil Girish BhalalaAllan-Hermann Pool Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Mar 2024
Abstract
Neuroscience incorporates manipulating neuronal circuitry to enhance the understanding of intricate brain functions. An effective strategy to attain this objective entails utilizing viral vectors to induce varied gene expression by delivering transgenes into brain cells. Here, we combine the use of transgenic mice, neonatal transduction with adeno-associated viral constructs harboring inhibitory designer receptor exclusively activated by designer drug (DREADD) gene, and the DREADD agonist clozapine N-oxide (CNO). In this way, a chemogenetic approach is employed to suppress neuronal activity in the region of interest during a critical developmental window, with subsequent investigation into its effects on the neuronal circuitry in adulthood.
Key features
• Comprehensive protocol for newborn viral transduction in the dorsal striatum of mice
• Uses a viral construct encoding inhibitory DREADD under the control of Cre recombinase to attenuate the activity of specific cell types in the brain
Keywords: Neonatal virus transduction hM4Di CNO Activity suppression Synaptic development
Graphical overview
Graphic overview of the procedure for chemogenetic silencing of striatal neurons in developing mouse brain. Newborn pups (first or second day of life) are injected with adeno-associated virus (AAV) coding inhibitory designer receptors exclusively activated by designer drugs (DREADD) to the dorsal striatum. Starting from postnatal day 6, pups are injected with clozapine N-oxide (CNO) subcutaneously twice a day for 9 consecutive days. Analysis of the consequences can be performed at the adult age.
Background
Using neonatal animals in scientific research allows the investigation of developmental processes. This is especially important for studying the maturation of neuronal networks. Time-limited critical periods of brain development are of particular interest due to their possible role in the origin of neuropsychiatric disorders [1]. Therefore, the approaches of time-specific alteration of neuronal activity within different brain areas are needed for this kind of research. Chemogenetic manipulations are widely used for time-limited modulation of neuronal activity in vivo. Here, we designed the protocol to suppress the activity of striatal projection neurons during the second week of life (P6–P14) to study the activity-dependent development of long-range GABAergic projections. To achieve this aim, we performed viral injections to the striatum of newborn (P0–2) mice with minimal surgery and under inhalation anesthesia, which ensures a high survival rate for the pups.
The current protocol for viral injections followed by designer receptors exclusively activated by designer drugs (DREADD)-agonist administration (CNO, clozapine-N-oxide dihydrochloride) was adapted from two previously published protocols [2] and [3]. As demonstrated by [2], inhalation-based anesthesia with isoflurane for early postnatal (P0−2) mice significantly enhances animal welfare. In our protocol, the skin and skull were gently perforated with a needle, which was later used to deliver the viral constructs. Alternatively, the skin and skull can be carefully perforated by a G30 syringe needle, and a glass pipette attached to a Hamilton syringe can be used to deliver the virus instead of a needle. This alternative is preferable when targeting smaller brain areas, such as the amygdala [4]. This needle-based or pipette-based injection through the skin and skull enhanced the survival of operated pups, because of the absence of sutures, which usually triggers infanticide. Although our protocol was designed for injections into the dorsal striatum, it can be easily adapted to another brain area of neonatal mice.
The following stereotaxic coordinates for viral injections in the striatum were determined in reference to the vascular lambda: AP 2.4; ML ±1.2; DV 2.3 and 2.7. The injections were done bilaterally and at two coordinates in the DV axis. The needle was not removed between the first and the second DV coordinate. The amount of the virus injected into each coordinate was 150 nL (left: 150 nL + 150 nL; right: 150 nL + 150 nL). Coordinates were based on those previously published [3]. If the coordinates for the brain area of interest are not known, the “Developing Mouse Brain Atlas” can be used to suggest the coordinates for injections in neonates. The suggested coordinates can then be checked by injecting a tissue dye under terminal anesthesia, followed by brain removal and slicing with the vibratome to verify coordinates with the dye signal. The brain can also be fixed with 4% PFA before checking the coordinates.
The injections of CNO were performed subcutaneously twice a day (9:00 and 18:00). Starting from P6, mice were injected with CNO dissolved in sterile saline solution (0.9% NaCl) or saline alone consecutively for 9 days. The administration method was chosen to prolong the effective time of the drug without sharply reaching the maximal concentration of the drug in the brain. It takes longer for the concentration of small molecules to reach maximal levels in the interstitial brain fluid when they are injected subcutaneously compared to intraperitoneally injected [5,6]. We used the CNO treatment of wild-type animals not expressing hM4Di as a control to verify that CNO itself did not cause any detectable effects. Alternatively, other muscarinic-based DREADD agonists can be used, such as Compound 21, deschloroclozapine, JHU37152, and JHU37160 [7–10]. These more modern alternative agonists differ from CNO in their pharmacological properties and generally demonstrate less off-target effects [7–9]. In this case, an appropriate dosage and an administration protocol should be defined separately. It is advisable to check if the compound was previously used in research related to the experimental objectives. This can aid in the design and refinement of the protocol and help in mitigating off-target effects if any have been previously documented. To assess potential off-target effects, administration of the compound to wild-type animals lacking DREADD expression can be used.
Materials and reagents
Biological materials
Mice: Oprm1-2A-Cre (Oprm1-Cre [11]),
Virus for DREADD: pAAV-hSyn-DIO-HA-hM4D(Gi)-IRES-mCitrine (a gift from Bryan Roth, Addgene viral prep # 50455-AAV8, http://n2t.net/addgene:50455; RRID: Addgene_50455)
Reagents
Clozapine-N-oxide dihydrochloride (HelloBio, catalog number: HB6149)
Sterile 0.9% NaCl (Fresenius Kabi AG, catalog number: 936808)
Isoflurane (1000 mg/g) (Attane vet 1000 mg/g) (Primal Critical Care, catalog number: 170579)
70% EtOH (Anora Industrial, ETAX, catalog number: 1025904)
Hydrogen peroxide 1% (Pharmacare)
Glycerol 99% (Sigma-Aldrich, catalog number: G6279) or sterile PBS (Sigma-Aldrich, catalog number: P2272)
Green dye: Fast green FCF (Sigma-Aldrich, catalog number: F-7262)
Solutions
CNO solution (see Recipes)
Green dye in glycerol solution or PBS (see Recipes)
Recipes
CNO solution
In a sterile 1.5 mL Eppendorf tube, measure the precise quantity of CNO powder and add sterile saline solution. Vortex well until the powder is totally dissolved. Aliquot the solution and store it at -20 °C.
Components Final concentration Quantity
CNO powder 30 mg/mL 15 mg
NaCl 9 g/L 500 μL
Total ~ 0.5 mL
Green dye in glycerol solution or PBS
Use a sterile Eppendorf tube (0.2–1.5 mL), add 200 µL of glycerol (99%) or sterile PBS, add green dye powder for a final concentration of dye of 1%, and vortex well until it becomes green. Store the solution with PBS at 4 °C; the glycerol-based solution can be stored at room temperature.
Components Final concentration Quantity
Glycerol or PBS 99% 200 μL
Green dye powder 1% 2 mg
Total 200 µL
Laboratory supplies
NF33BV-2 33 ga. beveled NanoFil needle (World Precision Instruments, UK, catalog number: NF33BV-2)
NanoFil syringe 10 μL (World Precision Instruments, UK, catalog number: NANOFIL)
200 μL pipette tip for making a mask (SARTORIUS, Safetyspace Tips, model: 790201)
T-style male luer fitting for tubing i.d. 1/8 in. to connect the mask and tubes (i.e., Sigma-Aldrich, catalog number: Z274240)
Scissors (World Precision Instruments, UK, catalog number: 500216)
Tape (Scotch Brand, Scotch® Magic™ 810, catalog number: 810-3PK-BXD)
Autoclavable bags (200 x 300 mm, Ratiolab, catalog number: 7001000)
Ice
Cotton buds
Eppendorf tube 1.5 mL (FisherBrand, catalog number: 05-408-129)
Eppendorf tube 0.2 mL (Nippon Genetics EUROPE GmbH, catalog number: FG-021)
Pipette 0.1–2 μL (Thermo Scientific™, F1-ClipTip™, catalog number: 4641310N)
Sterile filtered pipette tips 0.1–10 μL (SARTORIUS, Safetyspace filter tips, model: 790011F)
Hamilton syringe with male luer lock (25 μL), instrument syringe (Hamilton, model: 1702 TLLX, catalog number: 80222)
30G syringe needles (hypodermic needle, MicrolanceTM 3) (VWR, catalog number: 613-3942)
Blu Tack (Bostik, model: Blu Tack® Handy)
Wipes Kimberly-ClarkTM Kimtech ScienceTM delicate task wipes (Fisher Scientific, Kimberly-ClarkTM, catalog number: 7558)
Gloves
Tunnel for mice handling (https://3rc.org/refined-mouse-handling-overview/operations-and-tunnels), e.g., cardboard tube
PVC tubing 1/8 in. i.d. 1/4 in. o.d. (i.e., OC-TUBING, WPI, US)
Equipment
Stereotaxic frame for mice (StoelTing, US, model: 51615M)
Temperature controller with heating pad (Supertech Instruments UK Ltd., UK, model: TMP-5b and AHP-1)
Micro 4 Pump controller for NanoFil syringe (World Precision Instruments, UK, model: UMP3-4)
Anesthesia unit (Univentor, model: Univentor 400 Anesthesia Unit with tubes, 10 mL gas-tight glass syringe; anesthesia box, model: 8329001)
Air pump (EHEIM GmbH & Co. KG., Germany, model: EHEIM-100)
Dual guide LED light illuminator (Zeiss, Germany, model: ZEISS Cold Light Source)
Stereo microscope on boom stand (Olympus SZ30, model: SZ3060 SZ-STB1)
Procedure
Viral transduction
Disinfect the workstation and surgical tools with 70% EtOH. If the procedure is done in the laboratory hood, a 15–30 min UV light can also be used to disinfect the workstation and instruments before the procedure. The syringe can be sterilized by autoclaving or another method recommended by the manufacturer. Wear gloves and wash them with 70% EtOH.
Use scissors to make a mask for the pup. Cut the 200 μL tip so it can be connected to the anesthesia tube. Cut the other side so the pup's nose can fit in the tube. Place the tip connected to the anesthesia tube on the heating pad and fix it on the surface using tape or blu tack.
Note: We used a 200 μL tip and 1/8 i.d. tubes, but other materials can be used to make the mask, e.g., a 1 mL tip or a plastic Pasteur pipette tip. With these materials, it is also possible to use 1/4 i.d. tubes and fittings to place the mask.
Prepare the viral vector for injection.
Calculate the amount of the viral vector needed for injection, retrieve the proper amount of the aliquots from the -80 °C freezer, and thaw on ice. Always consider the extra amount needed due to possible loss of liquid when filling the syringe.
Note: Any excess amount of the viral solution should be discarded after the procedure since thaw/freezing cycles are not recommended. Keep the virus-containing liquid on ice.
Using a filtered tip, add glycerol to the viral solution (0.4–4 μL of viral aliquot). If you want to see the liquid with the virus, use glycerol or PBS with dye (see Recipes). Always use a new tip for each aliquot and discard the used tip (into an autoclavable bag or Virkon detergent solution, depending on your safety regulations).
Note: Step A3b can be skipped. Some researchers find that adding glycerol can protect the AAV and be used for in vivo injections [12,13]. Nevertheless, the absence of glycerol should not be considered dramatic, as many effective protocols do not include this step. The dye is needed for two reasons: first, the liquid containing viral particles is visible during the injection procedure, and second, the injected liquid can be visible within the brain of the neonatal pups through the scalp [14].
Mount the needle to the Nanofil syringe and attach it to the stereotaxic frame.
Place the blu tack piece on the heating pad or another surface and place the tube with the virus on it. The tube should stay vertical so the syringe can be filled when attached to the frame.
Using the pump, fill the NanoFil syringe with the virus-containing liquid in the amount needed to be injected for all the coordinates for one animal and 100 nL extra to compensate for liquid loss during injection or syringe itself. Use 10 nL/s pump speed for liquid withdrawal.
Carefully move the syringe attached to the frame to the side, so the frame is available to place the animal.
Anesthesia of the pup.
Turn on a heating pad to keep at 34–35 °C. It should be warm before the animal goes to the anesthesia unit.
Adjust the anesthesia unit to provide 4.7%–4.8% isoflurane anesthesia with appropriate air pressure to the unit box. Place a napkin in the box.
Place the pup in the anesthesia unit box and keep it closed for at least 2 min until breathing slows down. Switch the unit to provide isoflurane to the tube with a mask.
Use a small piece of wipe to make a flat pillow to support the animal's head in a horizontal position. Remove the pup and place it on the heating pad. Adjust its nose to the mask to breathe 4.7%–4.8% isoflurane. Tape the body to the heating pad with a piece of tape. When the animal is fixed on the frame, reduce isoflurane to 2%–2.5%. Check if the animal is anesthetized.
Caution: Always follow the animal's breathing; it should not be too slow or stop. Do not push too much with the tape on the thorax and do not squeeze the pup. The tape should only be kept so that the head does not move. If the breathing is fragmented, remove the animal from the mask immediately.
Viral injection.
Wash the head with the cotton bud with 70% EtOH.
Adjust the light so you can see lambda coordinate as an intersection of thick red vessels (see Figure 1).
Figure 1. Schematic image of the setup for the viral injection procedure. A. Setup consists of an anesthetizing unit for isoflurane connected to the switcher, directing the flow to the mask or box. The air pump supplying the unit is not shown. The syringe with a needle is assembled to the injecting pump headstage. The headstage is attached to the stereotaxic frame. A heating pad is placed on the stereotaxic frame surface. The pup is positioned on the heating pad, with its nose in the anesthesia mask. A stereoscope is used to see head blood vessels and lambda. The light source is adjusted to light up the pup’s head. B. The pup is placed on the heating pad. The pup’s nose is in the cut pipette tip (mask) connected to the tube system, which provides isoflurane. The tube system is fixed on the surface of the heating pad with the blu tack. A tape is used to gently fix the head of the pup. The head is placed on the flat pillow made from the wipe. The direction of isoflurane flow is demonstrated by arrows. C. The 0 coordinate from vascular lambda is defined as the very middle of the crossing between three branches of blood vessels.
Place the syringe tip to ML and AP lambda coordinates to find the 0 coordinate; then, find the coordinate needed for your purpose (for dorsal striatum: AP 2.4; ML ±1.2).
Carefully and slowly move the tip down to cross the skin and skull to achieve the correct DV coordinate (for dorsal striatum, DV 2.3 and 2.7). If the tip does not go smoothly through the skull and skin, use a common 30 G syringe needle to make a small hole in the skull before pushing the injecting needle. Start with the lower coordinate.
Note: There should not be bleeding. In case some minor bleeding occurs, use hydroxy peroxide and cotton bud to remove it. Never contact blood with the needle tip, as it will be clogged.
Inject virus liquid in the needed volume (we used 150 nL). Use 5 nL/s speed. (For dorsal striatum: next, move to the upper coordinate DV 2.3 and inject the same volume again with the same speed.) Wait 5 min before moving the needle up to remove it. Remove it slowly and carefully from the head.
Repeat steps A5c–A5e for the next coordinate if needed.
Wash the pup’s head with the cotton bud with hydroxy peroxide and then dry it.
Recovery.
Remove the animal from the mask but keep it on the heating pad until it reacts to the touch and moves.
Place the pup into the dam’s cage.
Subcutaneous CNO injections
Retrieve the vial with CNO from the freezer and thaw it at room temperature.
Separate pups from the dam and transfer them into the empty clean cage.
Note: When handling active pups, it is recommended to use a plastic tunnel or a cardboard tube rather than cupped hands. See https://3rc.org/refined-mouse-handling-overview/operations-and-tunnels.
Gently transfer a single pup from the cage and place it onto the weighing scale.
Caution: Make sure to prevent the pup's escape during the weighing procedure, e.g., by placing a lid on top of the scale.
Place the new needle onto the 25 μL Hamilton syringe and measure the proper volume of CNO according to the weight of the pup, with a final volume of 1 μL per gram of body weight. Control animals receive the injection of sterile saline, 1 μL per gram of body weight.
Administer the injection subcutaneously into the loose skin on the neck of the pup.
Caution: Always use different places on the skin for repeating injections.
Following the injection, maintain the needle within the skin for a few seconds before gently withdrawing it to prevent any potential drug leakage from the injection site.
Return the pup to the cage with the dam.
Repeat the procedure for the remaining pups. Change the needle for each injection.
Data analysis
The effect of the suppression of neuronal activity during the critical developmental period was further investigated by electrophysiological recordings and immunostaining. A detailed description of performed experiments and subsequent analysis can be found in the original research article [15] under the methods section.
Validation of protocol
This protocol has been used and validated in the research article:
Kokinovic et al. [15]. Spontaneous activity of striatal projection neurons supports maturation of striatal inputs to substantia nigra dopaminergic neurons. eLife.
First, we validated the expression of fluorescent protein co-expressed with DREADD. A coronal slice of dorsal striatum with the expression of mCitrine is demonstrated in Figure S2B [15].
It is important to demonstrate the effectiveness of DREADD, which is expected to modify the firing rate of infected neurons. For example, see validation in Figure S2 C of the same publication. In this validation step, we show in vitro the effect of the 10 μM CNO on the firing rate of the striosomal SPN of a 10-day-old mouse injected by DREADD coding AAV at P0. An example trace of membrane potential recording done with whole-cell patch-clamp technique is demonstrated.
The outcome of the procedure for brain development was analyzed by electrophysiology and microscopy in adult mice, with results shown in Figures 6 and 7 of [15].
General notes and troubleshooting
General notes
Always check and follow the safety rules of your research area when handling viruses and GMO mice.
The procedures in this protocol require appropriate training and animal license.
Avoid contamination when more than one virus is used in the same setup by using different syringes for each or replaceable glass pipettes for each virus to avoid contact between the virus and the syringe. Always change filtered tips when adding glycerol or dye to each virus.
If more than one virus needs to be injected, they can be mixed and injected at once. The increase in the volume of injected liquid should be considered.
Troubleshooting
Problem 1: The NanoFil syringe is not injecting.
Possible cause: The tip of your needle is clogged by blood cells or another dried organic liquid.
Solution: Try to unclog the needle with a thin wire, then wash and sterilize it or simply replace it with a new one.
Problem 2: Unspecific signal in the brain tissue during validation.
Possible cause: Contamination of syringe, needle, or dye stock by another virus.
Solution: Clean and sterilize the syringe and needle and make another dye stock. Use only your own syringe, needle, and stocks.
Problem 3: No expression of the virally delivered construct after injection.
Possible cause(s): The needle did not reach the desired coordinates during injection, and all the liquid went off under the skin or into the brain ventricles; the virus prep is spoiled (i.e., it was kept at room temperature for a long time, exposed to UV light, or subjected to freezing/tawing cycles); reasons associated with construct and experiment design, such as promoter/enhancer, Cre-dependence, etc.
Solution(s): Use dye to visualize the virus-containing liquid during the injection procedure and verify that it goes inside the brain. Try another aliquot/prep for the virus injection. Check your experiment design and animal genotype.
Acknowledgments
Funding for this research was provided by the Research Council of Finland (former Academy of Finland; #330298, M.R.; #324548, S.M.) and Jane and Aatos Erkko Foundation (S.M.). The protocol outlined here was adapted from the original research paper [15].
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal experiments were conducted according to the ethical guidelines (in accordance with EU directive 2010/63/EU) provided by the University of Helsinki. Breeding of the animals was done under the University of Helsinki license KEK22-013; procedures were done under the Regional State Administrative Agency of Southern Finland ESAVI/25166/2020.
References
Dehorter, N. and Del Pino, I. (2020). Shifting Developmental Trajectories During Critical Periods of Brain Formation. Front Cell Neurosci. 14: e00283. https://doi.org/10.3389/fncel.2020.00283
Ho, H., Fowle, A., Coetzee, M., Greger, I. H. and Watson, J. F. (2020). An inhalation anaesthesia approach for neonatal mice allowing streamlined stereotactic injection in the brain. J Neurosci Methods. 342: 108824. https://doi.org/10.1016/j.jneumeth.2020.108824
Olivetti, P. R., Lacefield, C. O. and Kellendonk, C. (2020). A Device for Stereotaxic Viral Delivery into the Brains of Neonatal Mice. Biotechniques. 69(4): 307–312. https://doi.org/10.2144/btn-2020-0050
Ryazantseva, M., Englund, J., Shintyapina, A., Huupponen, J., Shteinikov, V., Pitkänen, A., Partanen, J. M. and Lauri, S. E. (2020). Kainate receptors regulate development of glutamatergic synaptic circuitry in the rodent amygdala. eLife. 9: e52798. https://doi.org/10.7554/elife.52798
Durk, M. R., Deshmukh, G., Valle, N., Ding, X., Liederer, B. M. and Liu, X. (2018). Use of Subcutaneous and Intraperitoneal Administration Methods to Facilitate Cassette Dosing in Microdialysis Studies in Rats. Drug Metab Dispos. 46(7): 964–969. https://doi.org/10.1124/dmd.118.080697
Al Shoyaib, A., Archie, S. R. and Karamyan, V. T. (2019). Intraperitoneal Route of Drug Administration: Should it Be Used in Experimental Animal Studies?. Pharm Res. 37(1): e1007/s11095–019–2745–x. https://doi.org/10.1007/s11095-019-2745-x
Thompson, K. J., Khajehali, E., Bradley, S. J., Navarrete, J. S., Huang, X. P., Slocum, S., Jin, J., Liu, J., Xiong, Y., Olsen, R. H. J., et al. (2018). DREADD Agonist 21 Is an Effective Agonist for Muscarinic-Based DREADDsin Vitroandin Vivo. ACS Pharmacol Transl Sci. 1(1): 61–72. https://doi.org/10.1021/acsptsci.8b00012
Bonaventura, J., Eldridge, M. A. G., Hu, F., Gomez, J. L., Sanchez-Soto, M., Abramyan, A. M., Lam, S., Boehm, M. A., Ruiz, C., Farrell, M. R., et al. (2019). High-potency ligands for DREADD imaging and activation in rodents and monkeys. Nat Commun. 10(1): 4627. https://doi.org/10.1038/s41467-019-12236-z
Nagai, Y., Miyakawa, N., Takuwa, H., Hori, Y., Oyama, K., Ji, B., Takahashi, M., Huang, X. P., Slocum, S. T., DiBerto, J. F., et al. (2020). Deschloroclozapine, a potent and selective chemogenetic actuator enables rapid neuronal and behavioral modulations in mice and monkeys. Nat Neurosci. 23(9): 1157–1167. https://doi.org/10.1038/s41593-020-0661-3
Aomine, Y., Oyama, Y., Sakurai, K., Macpherson, T., Ozawa, T. and Hikida, T. (2023). Clozapine N-oxide, compound 21, and JHU37160 do not influence effortful reward-seeking behavior in mice. Psychopharmacology (Berl). 241(1): 89–96. https://doi.org/10.1007/s00213-023-06465-w
Märtin, A., Calvigioni, D., Tzortzi, O., Fuzik, J., Wärnberg, E. and Meletis, K. (2019). A Spatiomolecular Map of the Striatum. Cell Rep. 29(13): 4320–4333. https://doi.org/10.1016/j.celrep.2019.11.096
Chan, A., Maturana, C. J. and Engel, E. A. (2022). Optimized formulation buffer preserves adeno-associated virus-9 infectivity after 4 °C storage and freeze/thawing cycling. J Virol Methods. 309: 114598. https://doi.org/10.1016/j.jviromet.2022.114598
Qi, L., Iskols, M., Shi, D., Reddy, P., Walker, C., Lezgiyeva, K., Voisin, T., Pawlak, M., Kuchroo, V. K., Chiu, I. M., et al. (2024). A mouse DRG genetic toolkit reveals morphological and physiological diversity of somatosensory neuron subtypes. Cell. 187(6): 1508–1526.e16. https://doi.org/10.1016/j.cell.2024.02.006
Oomoto, I., Uwamori, H., Matsubara, C., Odagawa, M., Kobayashi, M., Kobayashi, K., Ota, K. and Murayama, M. (2021). Protocol for cortical-wide field-of-view two-photon imaging with quick neonatal adeno-associated virus injection. STAR Protoc. 2(4): 101007. https://doi.org/10.1016/j.xpro.2021.101007
Kokinovic, B., Seja, P., Donati, A., Ryazantseva, M., de Kerchove d’Exaerde, A., Schiffmann, S. N., Taira, T. and Molchanova, S. M. (2024). Spontaneous activity of striatal projection neurons supports maturation of striatal inputs to substantia nigra dopaminergic neurons. eLife. 13: e96574. https://doi.org/10.7554/elife.96574
Article Information
Publication history
Received: Jun 13, 2024
Accepted: Sep 1, 2024
Available online: Sep 14, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Basic technology > Chemogenetics
Biochemistry > Other compound > Small molecular
Developmental Biology > Cell growth and fate > Neuron
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Development of the Mammary Gland in Mouse: A Whole-Mount Microscopic Analysis
BW Bo Wang
YX Yuchen Xie
ZY Zejian Yang
JZ Jingyue Zhang
HZ Huiwen Zhang
PL Peijun Liu
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5089 Views: 281
Reviewed by: Chiara AmbrogioEmmanuelle Berret Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Sep 2023
Abstract
The mammary gland undergoes functional, developmental, and structural changes that are essential for lactation and reproductive processes. An overview of such unique tissue can offer clearer insights into mammary development and tumorigenesis. Compared to traditional methods, mouse mammary gland whole mount is a pivotal technique that provides three-dimensional structural perspectives on gland morphology and developmental stages, offering an inexpensive and accessible approach. This protocol outlines the tissue isolation of the mouse mammary gland and provides detailed instructions for whole-mount staining and analysis. Mammary gland tissues are carefully dissected from euthanized mice and stained with Carmine Alum to highlight the ductal structures, enabling detailed visualization of the branching patterns and morphological changes. Light microscopy is used to capture a panoramic image of the stained mammary gland, enabling the quantitative analysis of terminal end buds (TEBs) and bifurcated TEBs to further investigate mammary gland remodeling. This method can provide invaluable insights, particularly in the study of mammary gland morphogenesis and tumorigenesis, underscoring its significance in both basic research and clinical applications.
Key features
• Monitor mammary gland development within 2 days using whole-mount staining
Keywords: Mouse Mammary gland Whole-mount Mammary gland development Morphogenesis Terminal end buds Breast cancer Tumorigenesis
Graphical overview
Mouse mammary gland whole-mount analysis
Background
The mammary gland undergoes dynamic developmental stages during the fetus, puberty, and adulthood stages. Developmentally, initial ductal tree formation begins around E10.5, followed by estrogen-induced formation of terminal end buds (TEBs) that drive extensive branching during puberty [1]. The most significant morphogenic event occurs during pregnancy, where lobuloalveolar structures develop at the ends of ducts for milk production [2]. Post-lactation, these structures regress, and mammary glands undergo remodeling. Therefore, the dynamic developmental stages of the mammary gland can be visually observed in three-dimensional structural morphology, which facilitates a better examination of TEBs structure. Presently, classical traditional methods include immunohistochemistry (IHC) and immunofluorescence (IF). For instance, observing the morphology of the mammary gland through hematoxylin and eosin (H&E) staining and specific epithelial cell staining via IHC allows for precise characterization [3]. IF can further label luminal and basal cells to study the functional roles of these cellular subtypes [4]. Mouse mammary gland whole mount, in contrast, is a crucial technique that provides an entire view of duct architecture and developmental processes in mammary gland development and tumorigenesis, without requiring tissue sectioning. It enables the visualization of branching morphogenesis, the formation of TEBs, and the changes in these structures. Therefore, mouse mammary gland whole-mount analysis serves as a powerful tool for advancing our understanding of mammary gland development, pathology, and oncology. This protocol typically includes several key steps: isolation of mouse mammary gland tissue, procedures for whole-mount staining, and subsequent whole-mount analysis. This method provides valuable insights into mammary gland development.
Materials and reagents
Biological materials
8-week-old female mouse
Reagents
Ethanol (Sinopharm, catalog number: 10009218)
Chloroform (Sinopharm, catalog number: 10006818)
Glacial acetic acid (Sinopharm, catalog number: 10000208)
Carmine (Sigma-Aldrich, catalog number: C1022)
Aluminum potassium sulfate dodecahydrate (Solarbio, catalog number: A7520)
Hydrochloric acid (HCl) (Sinopharm, catalog number: 10011018)
Xylene (Sinopharm, catalog number: 10023418)
Permount (Fisher Scientific, catalog number: SP15-500)
Neutral balsam (Solarbio, catalog number: G8590)
Solutions
Carnoy’s fixative (see Recipes)
Carmine alum (see Recipes)
Wash buffer (see Recipes)
Recipes
Carnoy’s fixative (50 mL)
*Note: Prepare fixative just before use.
Reagent Final concentration Quantity or Volume
Ethanol 60% (v/v) 30 mL
Chloroform 30% (v/v) 15 mL
Glacial acetic acid 10% (v/v) 5 mL
Total (optional) n/a 50 mL*
Carmine alum (100 mL)
*Note: Add approximately 90 mL of water and heat the mixture on a hot plate until it boils for 15 min. After cooling the solution, filter it using filter paper. Adjust the volume to 100 mL and store at 4 °C.
Reagent Final concentration Quantity or Volume
Carmine 0.2% (w/v) 0.2 g
Aluminum potassium sulfate
dodecahydrate
0.5% (w/v) 0.5 g
H2O n/a see note*
Total (optional) n/a 100 mL
Wash buffer (100 mL)
*Note: Add water to make up to 100 mL.
Reagent Final concentration Quantity or Volume
Ethanol 70% (v/v) 70 mL
HCl (36%–38%) 2% (v/v) 5.64 mL
H2O n/a see note*
Total (optional) n/a 100 mL
Laboratory supplies
Glass slide (Citotest, catalog number: 80312)
50 mL centrifuge tubes (Corning, catalog number: 430829)
Coverslip (Citotest, catalog number: 80340)
Funnel (Citotest, catalog number: 84112)
Filter paper (Citotest, catalog number: 84501)
Volumetric cylinder (Citotest, catalog number: 84201)
Equipment
Scissors (RWD, catalog number: S12003-09)
Forceps (RWD, catalog number: F12005-10)
Autoclave (Panasonic, model: MLS-3781L-PC)
Hot plate (IKA, model: C-MAG HP 7)
Chemical fume hood (Rista lab, model: RSD-BLGTFG)
Dissecting microscope (OLYMPUS, model: SZX7)
Software and datasets
Prism v9.3 (GraphPad, 11/15/2021)
ImageJ 1.53e (NIH)
Procedure
Isolation of mouse mammary gland tissue
Euthanize an 8-week-old female mouse using CO2 inhalation.
Secure the limbs of the mice to a foam board with pushpins.
Using sterile scissors, make a skin incision on the abdomen and extend it toward the midline and hind limbs without breaching the peritoneum (Figure 1A).
Figure 1. Isolation of mouse mammary gland tissue for whole-mount staining. A. The euthanized mouse is secured on a foam board, and its skin is incised along the midline (as indicated by the red line). B. The incised skin is pinned back to fully expose the fourth pair of mammary glands (as marked within the red circle). C. The entire mammary gland tissue is flattened on a glass slide. D. The glass slide is inserted into a centrifuge tube for staining.
Securely pin the opened skin flat onto the foam board to adequately expose the fourth pair of mammary glands (Figure 1B).
Gently grasp the distal end of the entire fat pads using sterile forceps and slowly remove the mammary glands from the skin using sterile scissors. For detailed steps, refer to the video "Skin incision, removal of the mammary gland and spreading of the tissue onto glass slides" in [5].
Transfer the entire tissue onto a glass slide and gently spread the tissue using forceps (Figure 1C).
Place the glass slide at room temperature (RT) and allow the tissue edges to dry. After confirming that the tissue is securely adhered and not easily detached, transfer it into a 50 mL centrifuge tube for subsequent whole-mount staining (Figure 1D).
Whole-mount staining
Slowly add newly prepared Carnoy’s fixative to a 50 mL centrifuge tube, ensuring complete immersion of the entire mammary gland tissue in the chemical fume hood for 2–4 h or overnight at RT (Figure 2A).
Figure 2. Process of whole-mount staining of mouse mammary gland tissue. A. The entire mammary gland tissue is fixed in Carnoy’s fixative. B. The entire mammary gland tissue is stained in Carmine Alum. C. The stained mammary gland tissue is cleared in xylene. D. The cleared mammary gland tissue is covered in neutral balsam.
Transfer the glass slides to a 50 mL centrifuge tube for washing with 70% ethanol in the chemical fume hood for 15 min at RT. Mammary gland tissue processed in this step can be stored for several months.
Pass the glass slides sequentially through 50% and 30% ethanol and ddH2O in the chemical fume hood for 5 min each at RT.
Transfer the glass slides to Carmine Alum in the chemical fume hood at 4 °C overnight. Adjust staining duration based on the size and thickness of mammary gland tissue. Completed staining reveals a carmine-colored mammary epithelial tree and lymph node (Figure 2B).
Wash the glass slides with wash buffer in the chemical fume hood at RT until the solution becomes clear.
Pass the glass slides sequentially through 70%, 95%, and 100% ethanol in the chemical fume hood for 15 min each at RT.
Transfer glass slides to xylene for tissue clearing (Figure 2C) and mount them with Permount in the chemical fume hood at RT.
Apply Permount or neutral balsam to cover the entire mammary gland tissue in the chemical fume hood (Figure 2D) and store at RT.
Whole-mount analysis
Capture images of the stained entire mammary gland tissue using a dissecting microscope (Figure 3A).
Figure 3. Information from whole-mount analysis of mouse mammary gland tissue. A. The stained entire mammary gland tissue was captured by dissecting microscopy and contains information at different distances from the lymph node (scale bars, 2.5 mm). B–C. Images of TEBs (B) and bifurcated TEBs (C) were enlarged from the panoramic image (scale bars, 100 μm).
Count the number of TEBs (Figure 3B) and bifurcated TEBs (Figure 3C) directly on the images using a counter.
Count the number of ducts and acini at distances of 5, 10, and 15 mm from the lymph node along the direction of the mammary gland (Figure 3A).
Data analysis
The development of the mammary gland can be assessed directly by evaluating the mammary epithelial tree. Within this panoramic image, structures such as lymph nodes, TEBs, mammary ducts, and branches of the fourth pair of mammary glands can be distinguished (Figure 4A).
Figure 4. Examples of whole-mount analysis of mouse mammary gland tissue. A. Examples of whole-mount staining of mouse mammary gland tissue. B. Measurements of 5, 10, and 15 mm lengths (as marked within the red lines) were taken starting from the lymph node by using ImageJ (scale bars, 2.5 mm).
Prism was utilized to plot and statistically analyze the counts of TEBs and bifurcated TEBs within the entire mammary gland tissue, typically represented as "TEBs/gland (n)" and "bifurcated TEBs (n)."
Using ImageJ, the Straight tool was employed to draw scalebars length, and the scale was set using Analyze > Set Scale with the known distance input for the actual measurement of scale bars, ensuring Global was selected. Measurements of 5, 10, and 15 mm lengths were taken starting from the lymph node (Figure 4B). Prism was used to plot and statistically analyze the counts of ducts and acini within the 5, 10, and 15 mm distances from the lymph node, as well as across all distances, typically represented as "average complexity."
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Wang et al. [7]. CRB3 navigates Rab11 trafficking vesicles to promote γTuRC assembly during ciliogenesis. eLife (Figure 1, panel C).
General notes and troubleshooting
General notes
Mouse mammary gland whole mount is suitable for analyzing different stages of mammary gland development in mice, including pre-pubertal (5 weeks old), pubertal (8 weeks old), pregnant, and aging (25–90 weeks of age) stages.
A 50 mL centrifuge tube can be used to stain slides of two mammary gland tissue samples placed back-to-back.
To a better demonstration of mouse mammary glands whole mount from various groups, we attempted to parallelly lay out two whole mammary gland tissues on a single slide for subsequent staining and analysis.
If Permount is not readily available, mammary gland tissue clearing can be achieved using xylene, followed by mounting with neutral balsam.
Due to the thickness of the entire mammary gland tissues, the coverslip may not always sit flat during mounting. Applying pressure can flatten the coverslip and ensure a smooth seal.
Troubleshooting
Problem 1: The stained mammary gland tissues are inadequately stained or appear faintly stained.
Possible cause: Insufficient staining time.
Solution: Increase the staining duration, potentially allowing the mammary gland tissue to stain in Carmine Alum for several days.
Acknowledgments
This work was supported by grants from the National Natural Science Foundation of China (No. 82203772 and 82372931) and the Key Research and Development Program of Shaanxi Province of China (2023-YBSF-317 and 2024SF-YBXM-245). This protocol was adapted from Deng et al [6].
Competing interests
The authors declare no conflicts of interest.
Ethical considerations
This protocol involving mice was verified and approved by the Committee of Institutional Animal Care and Use of Xi'an Jiaotong University (permit number: XJTUAE2018-1801).
References
Watson, C. J. and Khaled, W. T. (2020). Mammary development in the embryo and adult: new insights into the journey of morphogenesis and commitment. Development. 147(22): e169862. https://doi.org/10.1242/dev.169862
Zwick, R. K., Rudolph, M. C., Shook, B. A., Holtrup, B., Roth, E., Lei, V., Van Keymeulen, A., Seewaldt, V., Kwei, S., Wysolmerski, J., et al. (2018). Adipocyte hypertrophy and lipid dynamics underlie mammary gland remodeling after lactation. Nat Commun. 9(1): 3592. https://doi.org/10.1038/s41467-018-05911-0
Nair, S. J., Zhang, X., Chiang, H. C., Jahid, M. J., Wang, Y., Garza, P., April, C., Salathia, N., Banerjee, T., Alenazi, F. S., et al. (2016). Genetic suppression reveals DNA repair-independent antagonism between BRCA1 and COBRA1 in mammary gland development. Nat Commun. 7(1): e1038/ncomms10913. https://doi.org/10.1038/ncomms10913
Patel, S., Sparman, N. Z. R., Arneson, D., Alvarsson, A., Santos, L. C., Duesman, S. J., Centonze, A., Hathaway, E., Ahn, I. S., Diamante, G., et al. (2023). Mammary duct luminal epithelium controls adipocyte thermogenic programme. Nature. 620(7972): 192–199. https://doi.org/10.1038/s41586-023-06361-5
Tolg, C., Cowman, M. and Turley, E. (2018). Mouse Mammary Gland Whole Mount Preparation and Analysis. Bio Protoc. 8(13): e2915. https://doi.org/10.21769/bioprotoc.2915
Deng, C. X. and Xu, X. (2004). Generation and Analysis of Brca1 Conditional Knockout Mice. In: Schönthal, A.H. (Eds) Checkpoint Controls and Cancer. Methods in Molecular Biology™, Humana Press. 280: 185–200. https://doi.org/10.1385/1-59259-788-2:185
Wang, B., Liang, Z., Tan, T., Zhang, M., Jiang, Y., Shang, Y., Gao, X., Song, S., Wang, R., Chen, H., et al. (2023). CRB3 navigates Rab11 trafficking vesicles to promote γTuRC assembly during ciliogenesis. eLife. 12: e4. https://doi.org/10.7554/elife.86689.4
Article Information
Publication history
Received: Jul 1, 2024
Accepted: Sep 1, 2024
Available online: Sep 14, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
Developmental Biology > Morphogenesis > Organogenesis
Cell Biology > Tissue analysis > Histomorphology
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5,090 | https://bio-protocol.org/en/bpdetail?id=5090&type=0 | # Bio-Protocol Content
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Peer-reviewed
Detecting Native Protein–Protein Interactions by APEX2 Proximity Labeling in Drosophila Tissues
JW Jhen-Wei Wu
CW Chueh-Wen Wang
WH Wei Yang Hong
AJ Anna C. C. Jang
YC Yu-Chiuan Chang
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5090 Views: 442
Reviewed by: Hong-Wen Tang Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Jul 2022
Abstract
Enzyme-catalyzed proximity labeling is a potent technique for the discernment of subtle molecular interactions and subcellular localization, furnishing contextual insights into the protein of interest within cells. Although ascorbate peroxidase2 (APEX2) has proven effective in this approach when overexpressed, its compatibility with endogenous proteins remains untested. We improved this technique for studying native protein–protein interactions in live Drosophila ovary tissue. Through CRISPR/Cas9 genome editing, APEX2 was fused with the endogenous dysfusion gene. By pre-treating the tissue with Triton X-100 to enhance biotin-phenol penetration, we successfully identified multiple proteins interacting with the native Dysfusion proteins that reside on the inner nuclear membrane. Our protocol offers a comprehensive workflow for delineating the interactome networks of ovarian components in Drosophila, aiding future studies on endogenous protein–protein interactions in various tissues of other animals.
Key features
• Elucidating Protein-protein interactions provides deeper insights into the regulation of gene expression in molecular network and complex signaling pathways, advancing protein engineering and drug design
• This protocol utilizes CRISPR/Cas9 knock-in technology to tag endogenous proteins with the APEX2 to label nearby proteins within a 20 nm radius in Drosophila melanogaster
• We optimize APEX2-proximity labeling by using Triton X-100 pre-treatment to enhance biotin-phenol penetration into Drosophila ovaries, enabling endogenous proteins enrichment under native conditions
Keywords: Proximity labeling APEX2 Drosophila Oogenesis Pull-down assay CRISPR/Cas9
Graphical overview
Workflow of proximity labeling with APEX2 tagging endogenous proteins in Drosophila ovary
Background
Proximity labeling is a powerful technique applied to mark adjacent proteins, providing valuable insights into the spatial arrangement of a given protein of interest within cells [1]. This approach excels over conventional assays because it can detect subtle or temporary protein–protein interactions (PPI) and pinpoint the precise localization within intact cellular compartments. Ascorbate peroxidase2 (APEX2) is engineered to enhance proximity labeling techniques for uncovering intricate subcellular proteomes in vivo [2]. By utilizing hydrogen peroxide (H2O2), APEX2 catalyzes biotin-phenol (BP) into biotin-phenoxyl radical, a highly reactive and short-lived species (lasting less than 1 ms), which quickly forms covalent bonds with nearby proteins within a 20 nm radius [3]. The resulting biotinylated proteins can be purified with streptavidin beads and then subjected to liquid chromatography-tandem mass spectrometry (LC-MS/MS) for identification [3]. Although APEX2 has shown efficacy in cell culture systems and Drosophila tissues when overexpressed, the compatibility with endogenous proteins tagged with this enzyme has not been tested [2–6]. While ectopic expression facilitates research on PPI, it may hinder embryonic development or lead to misinterpretation due to aberrant cellular behavior; a preferable method for unveiling PPI is to purify proteins expressed under native conditions.
In this protocol, we improved APEX2-based proximity labeling in live Drosophila ovary tissue to study PPI in the natural state. By using CRISPR/Cas9 genome editing, we generated a transgenic fly line that carries the APEX2 gene inserted at the N-terminal of endogenous dysfusion (dysf) locus. To enhance protein biotinylation catalyzed by APEX2, fly ovary tissues were pre-treated with Triton X-100 to facilitate the penetration of BP into tissues. After a sequence of protein enrichment and LC-MS/MS analysis, we identified numerous proteins that were labeled by APEX2-tagged Dysf proteins [7].
Altogether, our protocol provides a proximity-labeling workflow for mapping the interactome networks of Drosophila ovary. The level of detail in this protocol shall empower future researchers to explore endogenous PPI in living tissues of other multicellular organisms.
Materials and reagents
Biological materials
Drosophila melanogaster
Reagents
Schneider's Drosophila medium (Thermo Fisher Scientific, catalog number: 21720024)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, catalog number: A5670701)
100× Penicillin-Streptomycin (Thermo Fisher Scientific, catalog number: 15070063)
Streptavidin-horseradish peroxidase (streptavidin-HRP) (Life Technologies, catalog number: S-911)
cOmpleteTM, EDTA-free protease inhibitor cocktail (Merck, catalog number: 4693132001)
(3aS,4S,6aR)-hexahydro-N-[2-(4-hydroxyphenyl)ethyl]-2-oxo-1H-thieno[3,4-d]imidazole-4-pentanamide (Biotin-phenol, BP) (Iris Biotech, catalog number: LS-3500)
PierceTM Streptavidin magnetic beads (Thermo Fisher Scientific, catalog number: 88816)
Trolox (Merck, catalog number: Al-238813)
Sodium azide (Merck, catalog number: S2002)
Sodium L-ascorbate (Merck, catalog number: A4034)
Hydrogen peroxide solution (H2O2) (Merck, catalog number: 31642)
Phosphate buffered saline (PBS) (Merck, catalog number: P3813)
Bio-Rad protein assay dye reagent concentrate (Bio-Rad, catalog number: 5000006)
Tris base (cyrusbioscience, catalog number: 101-77-86-1)
Glycine (cyrusbioscience, catalog number: 101-56-40-6)
NaCl (cyrusbioscience, catalog number: 101-7647-14-5)
SDS (cyrusbioscience, catalog number: 101-151-21-3)
Triton X-100 (JT Baker®, X198-07)
Bovine serum albumin (BSA) (Bioman, catalog number: ALB001)
Dimethyl sulfoxide (DMSO) (Merck, catalog number: D2650)
Solutions
10× PBS stock
1 M Tris-HCL pH 8.0
5 M NaCl
10% SDS
Dissection medium (see Recipes)
500 mM Biotin-phenol (see Recipes)
100 mM H2O2 (see Recipes)
1 M Sodium ascorbate (see Recipes)
500 mM Trolox (see Recipes)
1 M Sodium azide (see Recipes)
Quencher solution (see Recipes)
25× Protease inhibitor cocktail (see Recipes)
20% Triton X-100 (see Recipes)
RIPA lysis buffer (see Recipes)
0.3% PBST (see Recipes)
Recipes
Dissection medium
Reagent Final concentration Quantity or Volume
Schneider's Drosophila medium 500 mM 445 mL
FBS 10% 50 mL
100× Penicillin-Streptomycin 1× 5 mL
Total n/a 500 mL
500 mM Biotin-phenol
Reagent Final concentration Quantity or Volume
Biotin-phenol 500 mM 100 mg
DMSO n/a Up to 550 μL
Total n/a 550 μL
The stock needs to be shaken by vortexing.
Aliquot the stock solution into small volumes and store at -80 °C to prevent repeated freeze-thaw cycles.
100 mM H2O2
Reagent Final concentration Quantity or Volume
30% H2O2(10M) 100 mM 10 μL
10× PBS 1× 100 μL
H2O n/a 890 μL
Total n/a 1 mL
Do not store this stock.
1 M Sodium ascorbate
Reagent Final concentration Quantity or Volume
Sodium ascorbate 1 M 0.59 g
H2O n/a Up to 5 mL
Total (optional) n/a 5 mL
500 mM Trolox
Reagent Final concentration Quantity or Volume
Trolox 500 mM 12.5145 mg
DMSO n/a Up to 100 μL
Total (optional) n/a 100 μL
Do not store this stock.
1 M Sodium azide
Reagent Final concentration Quantity or Volume
Sodium azide 1 M 0.65 g
H2O n/a Up to 10 mL
Total (optional) n/a 10 mL
Aliquots can be stored at -20 °C or below for several months.
Quencher solution
Reagent Final concentration Quantity or Volume
500 mM Trolox 5 mM 10 μL
1 M Sodium azide 10 mM 10 μL
1 M Sodium ascorbate 10 mM 10 μL
10× PBS
H2O
1×
n/a
100 μL
870 μL
Total (optional) n/a 1 mL
Make this solution immediately before it is to be used to quench the biotinylation reaction.
Do not store this solution.
25× Protease inhibitor cocktail
Reagent Final concentration Quantity or Volume
cOmpleteTM, EDTA-free protease inhibitor cocktail 25× 1 tablet
H2O n/a Up to 2 mL
Total (optional) n/a 2 mL
20% Triton X-100
Reagent Final concentration Quantity or Volume
Triton X-100 20% 10 mL
H2O n/a 40 mL
Total (optional) n/a 50 mL
RIPA lysis buffer
Reagent Final concentration Quantity or Volume
1 M Tris-HCL pH 8.0 50 mM 2.5 mL
5 M NaCl 150 mM 1.5 mL
10% SDS 0.1% 0.5 mL
20% Triton X-100 1% 2.5 mL
25× Protease inhibitor cocktail 1× 2 mL
H2O n/a 41 mL
Total (optional) n/a 50 mL
0.3% PBST
Reagent Final concentration Quantity or Volume
20% Triton X-100 0.3% 0.75 mL
10× PBS 1× 5 mL
H2O n/a 44.25 mL
Total (optional) n/a 50 mL
Equipment
Standard equipment for fly incubation
Micro refrigerated centrifuge (Kubota, model: 3740)
Magnetic stand (G-Biosciences, catalog number: 786-888)
Multi-mixer overhead mixer shaker Mischer Rotator (NanoEnTek, model: SLRM-3)
Ultrasonic processor (ChromTech, model: UP-500)
xMarkTM microplate absorbance spectrophotometer (BIO-RAD, catalog number: 1681150)
Mini Trans-Blot electrophoretic transfer cell (BIO-RAD, catalog number: 1703930)
Mini-PROTEAN® Tetra Cell (BIO-RAD, catalog number: 1658001)
Software and datasets
NCBIprot (20180429)
Procedure
Transgenic fly generation
The N-APEX2-dysf knock-in fly was produced by WellGenetics Company (Taiwan) using CRISPR/Cas9-mediated genome editing through the following steps.
Guide RNA construction: The pDCC6 vector was digested with BbsI and ligated with the annealed guide RNA (ACGTGGCCTAGACAAGGTGACGG). This was followed by transformation, colony PCR selection, and sequencing.
Donor plasmid construction: The APEX2 and 3XFlag sequences were amplified and cloned into the pUC57-Kan (Cassette RFP) vector using a sequence- and ligation-independent method (named Cassette APEX2-3XFlag-RFP). The two homology arms (upstream and downstream) of dysf were amplified from genomic DNA and then cloned into Cassette APEX2-3XFlag-RFP.
Microinjection: The constructs (donor plasmid and CRISPR/Cas9 vector plasmid) were injected into embryos according to standard procedures.
Selection and verification: Transgenic offspring were identified by genomic PCR and sequencing, and then crossed with balancer flies to establish a stable line. These steps are illustrated in Figure 1.
Figure 1. Workflow of generating the dysfJW allele using CRISPR/Cas9-mediated genome editing. A. APEX2, Piggy Bac terminal repeat, 3XFlag, homology arms, DsRed2, and 3XP3-hsp70 promoter were cloned into the pUC57-Kan plasmid as a donor plasmid. B. The guide RNA was cloned into the CRISPR/Cas9 vector plasmid (pDCC6). C. Both plasmids were co-injected into a Drosophila embryo following the standard protocol.
As a result of homology-dependent repair, the fragment of APEX2-3XFlag-PBacDsRed was inserted into the 5' end of dysf coding region after embryo microinjection, resulting in a new null allele, dysfJW. In order to generate an APEX2 knock-in tag in the dysf locus, dysfJW was crossed with BL8285 (w1118; CyO, P{Tub-PBac\T}2/wgSp-1; l(3)**/TM6B, Tb1) to excise the PBacDsRed fragment. After precise excision, one GTTAAA sequence was left as a linker peptide to bridge 3XFlag and dysf, as illustrated in Figure 2.
Figure 2. Generation of APEX2-3XFlag knock-in at the dysf locus. The APEX2-3XFlag-PBacDsRed fragment was inserted into the 5' end of the dysf coding region, creating dysfJW. To generate an APEX2 knock-in tag, dysfJW flies were crossed with BL8285 to excise the PBacDsRed fragment.
APEX2 reaction in Drosophila ovary
Fatten 60 N-APEX2-dysf genotype female flies, from 3 to 5 days old, with wet yeast for 14–16 h at 29 °C. Subsequently, carefully dissect their ovaries with fine-tip forceps in dissection medium [8].
Note: Quickly and gently tear the ovary, ensuring the process is brief to minimize tissue damage.
After dissection, pre-treat ovaries with 1 mL of 0.3% PBST for 15 min and then wash with 1 mL of 1× PBS once.
Note: Freshly prepare 0.3% PBST to enhance the penetration of biotin-phenol.
After pre-treatment, incubate all ovaries in 200 μL of dissection medium supplemented with 500 μM BP for 15 min at 25 °C.
Critical: Prewarm the medium to 25 °C to facilitate the dissolution of BP. Ensure that the solution fully dissolves in the dissection medium.
Prepare a premix by combining 10 μL of freshly prepared 100 mM H2O2 with 790 μL of dissection medium. Then, add this premix into the samples from step B3, achieving a final concentration of 1 mM H2O2, and incubate at room temperature for 1–2 min.
For the negative control group, we recommend using fruit fly ovaries without APEX2 or ovaries untreated with BP or H2O2.
The biotinylation time varies depending on the expression level of the APEX2-tagged target protein and cell types, typically ranging from 30 s to 2 min [4, 5, 7].
Rinse the ovaries three times, each for 1 min, with 1 mL of fresh quencher solution. Subsequently, carefully remove as much supernatant as possible.
Pause point: The ovaries can be stored at -80 °C for several months.
Ovary sample lysis
Homogenize ovary samples using a plastic pestle with 600 μL of RIPA lysis buffer and then sonicate for 10 cycles at 30% amplitude. Each cycle comprises 5 s of sonication followed by a 5 s rest on ice between pulses.
Centrifuge the lysate at 15,000× g for 10 min at 4 °C. Transfer the supernatant to a new tube and ensure to maintain the lysate on ice throughout the entire process.
Quantify the protein content in each clarified whole-cell lysate using the Bio-Rad Protein assay dye reagent concentrate. If needed, dilute the clarified lysate beforehand to ensure that the concentrations fall within the linear range of the assay. Take 50 μL aliquots of streptavidin magnetic beads and wash them twice with 1 mL of RIPA lysis buffer.
Streptavidin-based enrichment
For each sample, take 50 μL aliquots of streptavidin magnetic beads. Subsequently, incubate 550 μL of each lysate sample with 50 μL of streptavidin magnetic beads for 3 h at 4 °C on a rotator set at 10 rpm. To facilitate rotation, an additional 500 μL of RIPA buffer is added to each sample.
Pellet the beads using a magnetic rack and wash each bead sample five times with 500 μL of RIPA lysis buffer per wash to eliminate nonspecific binders. Keep the wash buffers on ice throughout the procedure.
Western blot analysis
To conduct the western blot analysis, start by preparing and boiling the samples in 1× protein loading buffer. Then, cool them on ice and briefly spin to minimize condensation.
Next, load and run the samples on an 8% (w/v) SDS gel, typically loading 15% of the streptavidin-based enrichment for analysis.
After gel electrophoresis, transfer the samples to a PVDF membrane using standard equipment and protocols.
Once transferred, block the membrane with 5% (w/v) BSA in 1× TBST for 2 h at room temperature.
Subsequently, gently rock the membrane in 10 mL of 0.3 μg/mL streptavidin-HRP in 1× TBST at room temperature for 2 h. Then, wash the membrane with 1× TBST four times for 5 min each before proceeding to membrane development.
If the streptavidin affinity pulldown is effective, it should result in the depletion of biotinylated proteins in the flowthrough fraction, while simultaneously enriching them in the bound fraction, as illustrated in Figure 3.
Figure 3. Streptavidin-based enrichment of biotin-tagged proteins. Western blot probed with streptavidin-HRP reveals proteins labeled by APEX2 in the co-treatment of biotin-phenol and H2O2 (right panel). Figure from Wu et al. [7]
Liquid chromatography-tandem mass spectrometry (LC-MS/MS)
Perform electrophoresis on the remaining samples from step E2 using an 8% Tris-glycine mini gel.
Apply Coomassie Blue staining to visualize proteins (Figure 4) and then carefully excise the stained bands from the gel using a new razor blade.
Figure 4. Coomassie blue staining for streptavidin-based enrichment. SDS-PAGE was stained with Coomassie blue to show biotinylated proteins. Color bars indicate sample fractions that were subjected to proteomic analysis. Figure from Wu et al. [7]
Submit the excised samples to a reputable spectrometry facility for in-gel tryptic digestion of proteins followed by liquid chromatography and mass spectrometry analysis, adhering to their established protocols.
Analyze the data generated by mass spectrometry from the samples.
Validation of protocol
This protocol has been employed and validated in the following research article:
• Wu et al. [7]. Spatiotemporal gating of Stat nuclear influx by Drosophila Npas4 in collective cell migration. Sci Adv. (Figures 6A and S5).
Acknowledgments
This project was supported by MOST (Ministry of Science and Technology, R.O.C.) grants to A.C.-C.J., MOST 110-2311-B-006-005 and MOST 107-2311-B-006-002-MY3, and to Y.C.-C., MOST 110-2311-B-110-004-MY3. This work was also supported by NSTC (National Science and Technology Council, R.O.C.) grant 113-2311-B-110-003 to Y.C.-C.
Competing interests
The authors declare no competing interests.
References
Bosch, J. A., Chen, C. and Perrimon, N. (2020). Proximity‐dependent labeling methods for proteomic profiling in living cells: An update. WIREs Dev Biol. 10(1): e392. https://doi.org/10.1002/wdev.392
Hung, V., Udeshi, N. D., Lam, S. S., Loh, K. H., Cox, K. J., Pedram, K., Carr, S. A. and Ting, A. Y. (2016). Spatially resolved proteomic mapping in living cells with the engineered peroxidase APEX2. Nat Protoc. 11(3): 456–475. https://doi.org/10.1038/nprot.2016.018
Rhee, H. W., Zou, P., Udeshi, N. D., Martell, J. D., Mootha, V. K., Carr, S. A. and Ting, A. Y. (2013). Proteomic Mapping of Mitochondria in Living Cells via Spatially Restricted Enzymatic Tagging. Science (1979). 339(6125): 1328–1331. https://doi.org/10.1126/science.1230593
Chen, C. L., Hu, Y., Udeshi, N. D., Lau, T. Y., Wirtz-Peitz, F., He, L., Ting, A. Y., Carr, S. A. and Perrimon, N. (2015). Proteomic mapping in live Drosophila tissues using an engineered ascorbate peroxidase. Proc Natl Acad Sci USA. 112(39): 12093–12098. https://doi.org/10.1073/pnas.1515623112
Mannix, K. M., Starble, R. M., Kaufman, R. S. and Cooley, L. (2019). Proximity labeling reveals novel interactomes in live Drosophila tissue. Development 146(14): e176644. https://doi.org/10.1242/dev.176644
Chakraborty, A., Lin, W. C., Lin, Y. T., Huang, K. J., Wang, P. Y., Chang, I. F., Wang, H. I., Ma, K. T., Wang, C. Y., Huang, X. R., et al. (2020). SNAP29 mediates the assembly of histidine-induced CTP synthase filaments in proximity to the cytokeratin network. J Cell Sci. 133(9): e240200. https://doi.org/10.1242/jcs.240200
Wu, J. W., Wang, C. W., Chen, R. Y., Hung, L. Y., Tsai, Y. C., Chan, Y. T., Chang, Y. C. and Jang, A. C. (2022). Spatiotemporal gating of Stat nuclear influx by Drosophila Npas4 in collective cell migration. Sci Adv. 8(29): eabm2411. https://doi.org/10.1126/sciadv.abm2411
Prasad, M., Jang, A. C., Starz-Gaiano, M., Melani, M. and Montell, D. J. (2007). A protocol for culturing Drosophila melanogaster stage 9 egg chambers for live imaging. Nat Protoc. 2(10): 2467–2473. https://doi.org/10.1038/nprot.2007.363
Article Information
Publication history
Received: May 13, 2024
Accepted: Aug 25, 2024
Available online: Sep 25, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Developmental Biology > Morphogenesis > Motility
Molecular Biology > Protein > Protein-protein interaction
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Metabolic RNA Labeling and Translating Ribosome Affinity Purification for Measurement of Nascent RNA Translation
HI Hirotatsu Imai
AY Akio Yamashita
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5091 Views: 475
Reviewed by: Alessandro DidonnaHemant Kumar PrajapatiWeidong An
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Cited by
Original Research Article:
The authors used this protocol in Nucleic Acids Research Aug 2023
Abstract
Regulation of gene expression in response to various biological processes, including extracellular stimulation and environmental adaptation, requires nascent mRNA synthesis and translation. Simultaneous analysis of the coordinated regulation of dynamic mRNA synthesis and translation using the same experiment remains a major challenge in the field. Here, we describe a step-by-step protocol for the simultaneous measurement of transcription of nascent mRNA and its translation at the gene level during the acute unfolded protein response (UPR) in HEK293 cells by combining 4-thiouridine metabolic mRNA labeling with translational ribosome affinity purification (TRAP) using a monoclonal antibody against evolutionarily conserved ribosomal P-stalk proteins (P-TRAP). Since P-TRAP captures full-length RNAs bound to ribosomes, it is compatible with 3' mRNA-seq, which analyzes the uridine-rich 3' UTRs of polyadenylated RNAs, allowing robust quantification of T>C conversions. Our nascent P-TRAP (nP-TRAP) method, in which P-TRAP is combined with metabolic mRNA labeling, can serve as a simple and powerful tool to analyze the coordinated regulation of transcription and translation of individual genes in cultured cells.
Key features
• Simple and retriable analysis of nascent mRNA synthesis and its translation in cultured cells
• Combination of 4-thiouridine metabolic RNA labeling with translating ribosome affinity purification (TRAP)
• Ribosomal P-stalk-mediated TRAP (P-TRAP) allows single-step and efficient purification of non-tagged ribosomes and translated mRNAs
Keywords: Translating ribosome affinity purification Metabolic RNA labeling Transcription Translation Deep sequencing 4-thiouridine TRAP-seq
Graphical overview
Background
Regulation of gene expression plays a central role in many biological processes. The final output of a protein-coding gene, the amount of protein, is determined by several regulation processes: transcription and degradation of mRNA, translation of mRNA, and degradation of protein.
Metabolic RNA labeling techniques have been developed to analyze the dynamic transcription of nascent mRNA and its degradation. One of these labeling techniques is a thiol (SH)-linked alkylation for the metabolic sequencing of RNA (SLAMseq), in which RNAs are labeled with the uridine analog 4-thiouridine (4sU) [1]. 4sU is incorporated into nascent RNAs by RNA polymerase II and then alkylated during cDNA library preparation, allowing bioinformatic analysis to detect the specific T-to-C (T>C) conversion at the 4sU incorporation site in 4sU-labeled RNA. By focusing on sequence reads containing T>C conversions, the accumulation rate of nascent transcribed RNA after the addition of 4sU (progressive labeling technique) or the degradation rate of pre-4sU-labeled RNA after the addition of an excess amount of uridine (pulse-chase technique) can be analyzed [2,3].
In addition to mRNA transcription and its degradation, mRNA translation significantly influences the level of gene expression [4]. Currently, to analyze the level of translation, three different methods are employed utilizing next-generation sequencing techniques: polysome profiling, translating ribosome affinity purification (TRAP), and ribosome profiling (Ribo-seq). Polysome profiling fractionates full-length ribosome-bound RNAs using sucrose density gradient ultracentrifugation. TRAP also fractionates full-length ribosome-bound RNAs, but it relies on the immunoprecipitation of ribosomes. Ribo-seq fractionates ribosome-protected RNA fragments (RPFs) after sucrose density gradient via ultracentrifugation and RNase digestion. Each method has advantages and disadvantages and is used according to the experimental design and objectives (reviewed in [5]).
The most dominant regulation process that determines the protein level is different for each gene. Therefore, the simultaneous measurement of these regulation processes in a single experiment is advantageous to understand the regulatory mechanism of gene expression, which is still a major challenge in this field. Kawata et al. simultaneously evaluated the synthesis rate of nascent mRNA and the degradation rate of pre-existing mRNA by using two different metabolic labeling analogs, i.e., 5'-bromo-uridine (BrU) and 4sU [6]. Statistical approaches have also been developed to estimate both the synthesis rate of nascent mRNA and the degradation rate of pre-existing mRNA using only 4sU labeling [2,7]. Recently, Schott et al. developed nascent Ribo-seq (nRibo-seq), a combination of 4sU metabolic RNA labeling and Ribo-seq, which allows simultaneous measurement of nascent mRNA synthesis and its translation at the level of bulk or specific RNA groups [8]. Although nRibo-seq is a pioneering approach for measuring mRNA transcription and translation, Ribo-seq deals with the short length of RPFs, making the reliable quantification of 4sU incorporation for individual genes difficult [8].
Here, we describe a step-by-step protocol for the simultaneous measurement of transcription of nascent mRNA and its translation at the gene level in the acute unfolded protein response (UPR), in which transcription and translation are dynamically reprogrammed to reduce unfolded and misfolded proteins in the endoplasmic reticulum and restore protein homeostasis [9] in HEK293 cells by combining 4sU metabolic mRNA labeling with TRAP (originally reported in [10]). The use of a monoclonal antibody against the evolutionarily conserved ribosomal proteins P0, P1, and P2 (P-stalk) is responsible for the highly efficient purification of endogenous translating ribosomes without an affinity tag (e.g., FLAG-tag, His-tag, and SBP-tag), shortening the experimental period (termed as the P-TRAP method). Combining this P-TRAP with 4sU metabolic mRNA labeling and 3' mRNA-Seq (which analyzes the uridine-rich 3' UTR of polyadenylated RNAs) results in robust quantification of T>C conversion and reliable analysis of nascent mRNA transcription as well as its translation at the individual gene level. If the sensitivity of nascent RNA detection is sufficient, subtracting T>C reads from total reads will yield non-T>C reads that are mostly derived from pre-existing RNAs, contributing to further analysis of translation in not only nascent but also pre-existing RNAs. This simple and versatile nascent P-TRAP (nP-TRAP) method could enhance our understanding of the complex regulation of gene expression in eukaryotes.
Materials and reagents
Biological materials
Flp-InTM T-REXTM 293 cell line (Thermo Fisher Scientific, catalog number: R78007)
Reagents
DMEM (high glucose) with L-Glutamine, phenol red and sodium pyruvate (FUJIFILM Wako, catalog number: 043-30085)
Penicillin-streptomycin mixed solution (Nacalai Tesque, catalog number: 09367-34)
Fetal bovine serum (FBS) (Cell Culture Bioscience, catalog number: 171012)
D-PBS(-) without Ca and Mg, liquid (Nacalai Tesque, catalog number: 14249-24)
Nuclease-free water (not DEPC-treated) (Thermo Fisher Scientific, catalog number: AM9937)
HEPES (Merck, catalog number: H3375-500G)
Tris (Nacalai Tesque, catalog number: 035401-25)
HCl (FUJIFILM, catalog number: 080-01066)
Glycine (FUJIFILM, catalog number: 077-00735)
MgCl2 (Nacalai Tesque, catalog number: 20908-65)
NaCl (FUJIFILM Wako, catalog number: 191-01665)
NaH2PO4·H2O (Nacalai Tesque, catalog number: 31719-05)
Na2HPO4·12H2O (FUJIFILM Wako, catalog number: 196-02835)
DTT (Roche, catalog number: 10708984001)
Triton X-100 (Nacalai Tesque, catalog number: 35501-15)
Tween 20 (FUJIFILM, catalog number: 167-11515)
SDS (SERVA, catalog number: 20765.02)
DMSO (FUJIFILM Wako, catalog number: 043-07211)
Glycerol (FUJIFILM Wako, catalog number: 075-00616)
Bromophenol blue (FUJIFILM Wako, catalog number: 021-02911)
BSA (Nacalai Tesque, catalog number: 01860-07)
Protease inhibitor cocktail for use with mammalian cell and tissue extracts (Nacalai Tesque, catalog number: 25955-11)
Phosphatase inhibitor cocktail (EDTA-free) (Nacalai Tesque, catalog number: 07575-51)
4-thiouridine (4sU) (LKT Laboratory, catalog number: T2933)
Thapsigargin (Cayman Chemical, catalog number: 10522)
Cycloheximide (Nacalai Tesque, catalog number: 06741-91)
RNase A (100 mg/mL) (NIPPON GENE, catalog number: 318-06391)
RNasin® Plus ribonuclease inhibitor (Promega, catalog number: N2611)
DynabeadsTM protein G for immunoprecipitation (Thermo Fisher Scientific, catalog number: 10004D)
Anti-ribosomal proteins P0/P1/P2 mAb [9D5] (MBL, catalog number: RN004M)
Mouse IgG2a isotype control (Proteintech, catalog number: 65208-1-Ig)
Antibodies for western blotting:
Anti-ribosomal protein uL3 (GeneTex, catalog number: GTX114725)
Anti-ribosomal protein uS2 (GeneTex, catalog number: GTX114734)
Anti-PABP4 (BETHYL, catalog number: A301-467A), primary antibody
Anti-CBP80 (BETHYL, catalog number: A301-793A), primary antibody
Anti-eIF4A3 [11], primary antibody
Anti-GAPDH mAb-HRP-DirecT (MBL, catalog number: M171-7)
Anti-rabbit IgG, HRP-linked antibody (Cell Signaling Technology, catalog number: 7074S)
Anti-mouse IgG, HRP-linked antibody (Cell Signaling Technology, catalog number: 7076S)
ProClinTM 950 (Merck, catalog number: 46878-U)
e-PAGEL 5%–20% 18-well precast mini gel (ATTO, catalog number: E-R520L)
Pre-stained XL-Ladder Broad (APRO, catalog number: SP-2110)
EzFastBlot Fast western blotting transfer buffer (ATTO, catalog number: AE-1465)
ECL selectTM western blotting detection reagent (Cytiva, catalog number: RPN2235)
ImmunoStar® LD (FUJIFILM, catalog number: 296-69901)
ISOGEN II (NIPPON GENE, catalog number: 311-07361)
2-Propanol (Nacalai Tesque, catalog number: 29113-95)
Ethanol (99.5) (Nacalai Tesque, catalog number: 14713-95)
Iodoacetamide, No-WeighTM format (Thermo Fisher Scientific, catalog number: A39271)
Sodium acetate nuclease and protease tested (Nacalai Tesque, catalog number: 31137-25)
3 M NaOAc pH 5.2 (FUJIFILM Wako, catalog number: 316-90081)
10 N NaOH (Nacalai Tesque, catalog number: 94611-45)
Glycogen solution (20 mg/mL) from Oyster, nuclease tested (Nacalai Tesque, catalog number: 17110-11)
TE buffer pH 8.0 (Nacalai Tesque, catalog number: 06890-54)
RNA 1000 reagent kit for MultiNA (SHIMADZU, catalog number: 292-27913-91)
SYBRTM Green II RNA gel stain, 10,000× concentrate in DMSO (Invitrogen, catalog number: S7568)
RNA 6000 ladder (Invitrogen, catalog number: AM7152)
Formamide (Nacalai Tesque, catalog number: 16229-95)
QuantSeq 3' mRNA-Seq Library Prep Kit FWD for Illumina (Lexogen, catalog number: 192.14)
Note: We used the currently unavailable catalog number 015.24. One may use 191.24 as an alternative.
Qubit® dsDNA HS Assay Kits (molecular probes life technologies, catalog number: Q32854)
Solutions
20× TBS stock solution
1 M Tris-HCl pH 7.5 stock solution
1 M MgCl2 stock solution
5 M NaCl stock solution
1 M DTT stock solution
1% SDS stock solution
Tris-glycine SDS running buffer (see Recipes)
1 M HEPES-NaOH pH 7.5 stock solution (see Recipes)
100 mg/mL cycloheximide (see Recipes)
1% Triton X-100 lysis buffer (see Recipes)
Wash buffer (see Recipes)
SDS buffer (see Recipes)
6× SDS-dye (see Recipes)
TBST (see Recipes)
1% (w/v) BSA/TBST (see Recipes)
200 mM 4-thiouridine (4sU) (see Recipes)
1 mM Thapsigargin (TPG)
1 M NaH2PO4 (see Recipes)
1 M Na2HPO4 (see Recipes)
0.5 M NaPO4 pH 8.0 (see Recipes)
100 mM IAA (see Recipes)
Recipes
Tris-glycine SDS running buffer (2,000 mL)
Reagent Final concentration Quantity or Volume
Tris 25 mM 6 g
Glycine 192 mM 28.8 g
SDS 0.1% (w/v) 2 g
Deionized water n/a Up to 2,000 mL
Note: Store at room temperature (RT).
1 M HEPES-NaOH pH 7.5 (1,000 mL)
Reagent Final concentration Quantity or Volume
HEPES 1 M 238.3 g
10 N NaOH n/a To pH 7.5
Deionized water n/a Up to 1,000 mL
Notes:
Prepare ~800 mL of deionized water in a suitable container. Add 238.3 g of HEPES to the solution. Adjust solution to the desired pH using 10 N NaOH. Add deionized water until the volume is 1,000 mL.
Store at 4 °C.
100 mg/mL cycloheximide (0.1 mL)
Reagent Final concentration Quantity or Volume
Cycloheximide 100 mg/mL 10 mg
DMSO n/a 0.1 mL
Note: Prepare before use.
1% Triton X-100 lysis buffer (10 mL)
Reagent Final concentration Quantity or Volume
1 M HEPES-NaOH pH 7.5 20 mM 0.2 mL
1 M MgCl2 2.5 mM 0.025 mL
5 M NaCl 150 mM 0.3 mL
Triton X-100 1% (v/v) 0.1 mL
1 M DTT 0.5 mM 5 μL
Cycloheximide (100 mg/mL) 100 μg/mL 10 μL
Protease inhibitor cocktail (100×) 1× 0.1 mL
Phosphatase inhibitor cocktail (100×) 1× 0.1 mL
Nuclease-free water n/a Up to 10 mL
Notes:
Store at RT.
Add DTT, cycloheximide, protease inhibitor cocktail, and phosphatase inhibitor cocktail before use.
Wash buffer (50 mL)
Reagent Final concentration Quantity or Volume
1 M HEPES-NaOH pH 7.5 20 mM 1 mL
1 M MgCl2 2.5 mM 0.125 mL
5 M NaCl 150 mM 1.5 mL
Tween 20 0.025% (v/v) 12.5 μL
Nuclease-free water n/a 47.375 mL
Note: Store at RT.
SDS buffer (10 mL)
Reagent Final concentration Quantity or Volume
1 M HEPES-NaOH pH 7.5 20 mM 1 mL
1% SDS 1% 1 mL
Nuclease-free water n/a 8 mL
Note: Store at RT.
6× SDS-dye (10 mL)
Reagent Final concentration Quantity or Volume
1 M Tris-HCl pH 7.5 0.3 M 3 mL
SDS 9% (w/v) 0.9 g
Glycerol 30% (v/v) 3 mL
1 M DTT 0.6 M 6 mL
Bromophenol blue 0.1% (w/v) 10 mg
Nuclease-free water n/a Up to 10 mL
Note: Store at -20 °C.
TBST (2,000 mL)
Reagent Final concentration Quantity or Volume
20× TBS 1× 100 mL
Tween 20 0.1% (v/v) 2 mL
Deionized water n/a Up to 2,000 mL
Note: Store at RT.
1% BSA/TBST (50 mL)
Reagent Final concentration Quantity or Volume
1× TBST 1× 50 mL
BSA 1% (w/v) 0.5 g
Note: Prepare before use.
200 mM 4-thiouridine (0.5 mL)
Reagent Final concentration Quantity or Volume
4-thiouridine (MW 260.27) 200 mM 26 mg
Nuclease-free water n/a 0.5 mL
Note: Store at -20 °C and protected from light.
1 mM Thapsigargin (1.5 mL)
Reagent Final concentration Quantity or Volume
Thapsigargin (MW 650.8) 1 mM 1 mg
DMSO n/a 1.536 mL
Note: Dispense 20 μL each and store at -80 °C.
1 M NaH2PO4 (10 mL)
Reagent Final concentration Quantity or Volume
NaH2PO4·H2O 1 M 1.38 g
Nuclease-free water n/a Up to 10 mL
Note: Store at RT.
1 M Na2HPO4 (10 mL)
Reagent Final concentration Quantity or Volume
Na2HPO4·12H2O 1 M 3.58 g
Nuclease-free water n/a Up to 10 mL
Note: Store at RT.
0.5 M NaPO4 pH 8.0 (10 mL)
Reagent Final concentration Quantity or Volume
1 M NaH2PO4 0.034 M 0.34 mL
1 M Na2HPO4 0.466 M 4.66 mL
Nuclease-free water n/a 5 mL
Note: Store at RT after 0.22 μm filtration.
100 mM IAA (0.5 mL)
Reagent Final concentration Quantity or Volume
Iodoacetamide, No-WeighTM 100 mM 9.3 mg (1 vial)
EtOH n/a 0.5 mL
Note: Prepare before use.
Laboratory supplies
Cell culture 6-well plate (SPL Life science, catalog number: 30006)
Tissue culture dish Φ96 × 21 mm (10 cm) (TPP, catalog number: 93100)
Falcon® 5 mL serological pipette (CORNING, catalog number: 357543)
Falcon® 10 mL serological pipette (CORNING, catalog number: 357551)
FastGeneTM centrifuge tubes 15 mL (NIPPON Genetics, catalog number: FG400)
SuperClear® centrifuge tubes 50 mL (Labcon, catalog number: 3181-345)
Round-bottom micro tube 1.7 mL (BIO-BIK, catalog number: RC-0170)
Siliconized microcentrifuge tube 1.5 mL (WATSON, catalog number: 131-615CH)
FastGeneTM 0.2 mL 8 strips PCR tube and FlatCap (NIPPON Genetics, catalog number: FG-028FC)
Long tip low adsorption 10 μL (BMBio ST, catalog number: BMSSCH0001)
FastGeneTM tip 200 μL (NIPPON Genetics, catalog number: FG-3001)
Long tip low adsorption 1,000 μL (BMBio ST, catalog number: BMSSCH0004)
Siliconized tip 1,000 μL (WATSON, catalog number: 121-814CH)
Immobilon®-P PVDF membrane (Millipore, catalog number: IPVH00010)
Chromatography paper (ATTO, catalog number: CB-20A)
Equipment
PIPETMAN P-10 (Gilson, catalog number: F144802)
PIPETMAN P-20 (Gilson, catalog number: F123600)
PIPETMAN P-200 (Gilson, catalog number: F123601)
PIPETMAN P-1000 (Gilson, catalog number: F123602)
Pipette Mate NEO (NICHIRYO, catalog number: 00-PMNEO)
Bioclean Bench (PHCbi, catalog number: MCV-B161S)
CO2 incubator Prescyto MG-71C (TAITEC, catalog number: 0081460-000)
Inverted phase contrast microscope (OLYMPUS, catalog number: CK40)
Thermo Minder SDminiN (TITEC, catalog number: 0068750-000)
Aspirator (NISSIN, catalog number: NVP-11)
High-speed refrigerated micro centrifuge (TOMY, catalog number: MX-305)
Biomedical cooler, 4 °C (NIHON FREEZER, catalog number: UKS-5410DHC)
Biomedical freezer, -20 °C (PHCbi, catalog number: MDF-U539)
Ultra-low freezer, -80 °C (PHCbi, catalog number: MDF-DU702VHS1)
VORTEX-GENIE 2 Mixer (M&S Instruments, catalog number: SI-0286)
Ice On I (SK BIO International, catalog number: IO-1)
DynaMagTM-Spin Magnet (Invitrogen, catalog number: 12320D)
Miniature rocker shaker (NISSIN, catalog number: NX-12)
Rotary Mixer with Oshiri Penpen A type (NISSIN, catalog number: NRC-20D)
Hybridization rotator (NISSIN, catalog number: SN-06BN)
Dry thermos unit DTU-1CN (TITEC, catalog number: 0075930-000)
MiniSlab, AE-6401 Gel caster included (ATTO, catalog number: WSE-1165W)
HorizeBLOT 2M (ATTO, catalog number: WSE-4025M)
ImageQuant LAS 4000 mini (Cytiva)
NanoDrop 1000 (Thermo Fischer Scientific, catalog number: ND-1000)
Microchip electrophoresis system for DNA/RNA analysis MultiNA (SHIMADZU, catalog number: MCE-202)
Agencourt SPRIPlate 96R super magnet plate (Beckman Coulter, catalog number: A32782)
SpectraMax Paradigm multi-mode microplate reader (Molecular Devices)
Software and datasets
SLAM-DUNK v.0.4.3 (https://github.com/t-neumann/slamdunk) [12]
R v.4.2.1 (https://www.r-project.org/)
Python v.3.7 (https://www.python.org/)
DESeq2 v.1.3.8 (http://www.bioconductor.org/packages/release/bioc/html/DESeq2.html) [13]
deltaTE (https://github.com/SGDDNB/translational_regulation) [14]
ImageQuantTM TL v.1.2 (Cytiva)
Procedure
Before extracting ribosome-bound RNAs for preparing cDNA libraries, we recommend validating that P-TRAP works well on a small scale by western blot as described in Sections A and B. P-TRAP can be applied to other eukaryotic cell lines and animals. We have successfully applied P-TRAP to human A549 cells, mouse B16 cells, mouse MIN6 cells, mouse liver, budding yeast, nematode, and zebrafish.
Immunoprecipitation and detection of translating ribosomes using western blot analysis in HEK293 cells
Seed 2 × 105 HEK293 cells in 2 mL of DMEM supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin into a 6-well plate and incubate overnight in a humidified incubator with 5% CO2 at 37 °C. Use one well per sample.
After 24 h, aspirate the medium from the plate and add 250 μL of freshly prepared ice-cold 1% Triton X-100 lysis buffer (see Recipes).
Note: In this condition, the ER fraction is soluble, but the nuclear fraction is insoluble.
Gently shake the 6-well plate to ensure that the lysis buffer covers the entire surface of each well, transfer to a 1.5 mL tube using a micropipette, and centrifuge at 20,000× g for 10 min at 4 °C using a refrigerated centrifuge with a swing rotor.
Notes:
Work on ice to keep the 6-well plate and tubes cold.As the sample warms up and translation proceeds, the ribosomes will dissociate from the mRNA.
No need to use a cell scraper.
Collect the supernatant in a 1.5 mL tube. Keep 10 μL of the supernatant in another 1.5 mL tube as input fraction for western blotting. Add 2 μL of 6× SDS-dye (see Recipes) to input fraction, incubate at 95 °C for 5 min, and store at -20 °C.
Prepare the following 10× antibody solution with RNase A (Table 1).
Table 1. 10× antibody solution with RNase A
Reagent Concentration (10×) Amount per 1 sample
1% Triton X-100 lysis buffer (freshly prepared) n/a 18.8 μL
1 μg/mL anti-ribosomal proteins P0/P1/P2 mAb [9D5] 0.05 μg/μL 1 μL
RNase A (100 mg/mL) 1 mg/mL 0.2 μL
Note: The Mouse IgG2a isotype control (from Proteintech) can be used as a negative control for the 9D5 antibody.
Add 20 μL of 10× antibody solution with RNase A to 180 μL of the supernatant in a new 1.5 mL tube (the final antibody solution will be 1×) and rotate at 4 °C for 60 min on an NRC-20D rotary mixer with an Oshiri Penpen A type.
Note: The NRC-20D with an Oshiri Penpen A type is a rotary mixer with a tube tapping function. Tapping the tube during stirring prevents the beads from sticking to the bottom of the tube and improves the efficiency of immunoprecipitation. However, it is not essential.
Dispense 7 μL of DynabeadsTM Protein G beads per sample into a new 1.5 mL tube and wash the beads by 200 μL with wash buffer (see Recipes). Insert the tube into the magnetic rack and remove the clear supernatant.
Transfer 200 μL of sample solution to the Dynabeads tube and rotate at 4 °C for 40 min on an NRC-20D rotary mixer with an Oshiri Penpen A type.
Place the tube in a magnetic rack and remove the clear supernatant. Keep 10 μL of the supernatant in another 1.5 mL tube as an unbound fraction for western blotting. Add 2 μL of 6× SDS-dye (see Recipes) to the unbound fraction, incubate at 95 °C for 5 min, and store at -20 °C. This fraction helps us to check the efficiency of immunoprecipitation.
Wash the beads three times with 1 mL of wash buffer (see Recipes). Each time, resuspend the beads in wash buffer using a micropipette with a siliconized pipette tip and transfer to a new 1.5 mL siliconized tube to allow adsorption of non-specific binding proteins to the tip and tube. Place the tube in a magnetic rack and remove the clear supernatant (wash buffer).
Note: The beads should be suspended in the wash buffer and the supernatant removed as rapidly as possible. Furthermore, care should be taken to prevent the beads from drying out during this process.
Add 30 μL of SDS buffer (see Recipes) to the beads and resuspend to elute the immunoprecipitants. Place the tube in a magnetic rack and transfer the clear supernatant to a new 1.5 mL tube as an eluted fraction for western blotting. Add 6 μL of 6× SDS-dye (see Recipes) to the eluted fraction, incubate at 95 °C for 5 min, and store at -20 °C.
Western blotting
Place the e-PAGEL 5%–20% precast gel (18 wells) in the WSE-1165W mini slab filled with Tris-Glycine SDS running buffer (see Recipes) and load 1.5 μL of molecular weight marker and 5 μL of each sample prepared in Section A. Run the gel at 180 V for 50–60 min at RT.
After the gel runs, activate the PVDF membrane by soaking it in EtOH for a few seconds and incubate with 1× EzFastBlot transfer buffer until use.
Transfer proteins from the gel to PVDF membranes under wet transfer conditions at 7 mA/cm2 (e.g., 441 mA for 6 cm × 9 cm PVDF membrane) for 15 min in the WSE-4025M HorizeBLOT at RT.
Block the membrane with 10 mL of 1% BSA/TBST (see Recipes) for at least 30 min at RT with gentle agitation on the NX-12 rocker shaker (speed: 3).
Prepare each of the primary antibody mix (Table 2) in 50 mL tubes. Each primary antibody is diluted in 2 mL of 1% BSA/TBST and stored at 4 °C until use.
Table 2. Primary antibody mix
Catalog numbers Protein Molecular weight Amount in 2 mL of 1% BSA/TBST Dilution
MBL, RN004N P0 36,000 1 μL 1/2,000
GeneTex, GTX114725 uL3 46,000 1 μL 1/2,000
GeneTex, GTX114734 uS2 31,000 1 μL 1/2,000
BETHYL, A301-467A PABP4 70,000 2 μL 1/1,000
BETHYL, A301-793A CBP80 95,000 2 μL 1/1,000
(Okada-Katsuhata et al. [11]) eIF4A3 51,000 0.1 μL 1/20,000
Note: Primary antibodies can be reused. For long-term storage, add 0.2 μL of the ProClinTM 950 to 2 mL of the solution (1/10,000 dilution) and store at 4 °C.
Place the membrane in 50 mL super-seal cap tubes containing each primary antibody solution with the transferred side facing the inside of the tube. Set the 50 mL tube to the SN-06BN hybridization rotator and incubate at 4 °C with rotation overnight.
Note: If necessary, cut the membrane according to the molecular weight of the target protein.
Wash the membranes three times with 10 mL of TBST (see Recipes) for at least 10 min at RT with gentle agitation on the NX-12 rocker shaker (speed: 3).
Prepare each of the secondary antibody mix (Table 3) in 50 mL tubes. Each secondary antibody is diluted in 2 mL of 1% BSA/TBST and stored at 4 °C until use.
Table 3. Secondary antibody mix
Catalog numbers Organisms Amount in 2 mL of 1% BSA/TBST Dilution
CST, 7076S Mouse 0.4 μL 1/5,000
CST, 7074S Rabbit 0.4 μL 1/5,000
Note: Secondary antibodies should be freshly prepared.
Place the membrane in 50 mL super-seal cap tubes containing each secondary antibody solution with the transferred side facing the inside of the tube. Set the 50 mL tube to the SN-06BN hybridization rotator and incubate at RT for at least 30 min with rotation.
Wash the membranes three times with 10 mL of TBST solution for at least 10 min at RT with gentle agitation on the NX-12 rocker shaker (speed: 3).
Turn on and boot the ImageQuant LAS 4000 mini. Then, incubate the membranes with the appropriate HRP western blot substrate solution on plastic wrap. Place the membrane in the LAS 4000 mini chamber and acquire images (Figure 1).
Notes:
HRP substrate solutions vary in signal intensity depending on the product. Select the most appropriate one according to the titer of the antibody. The exposure time is automatically determined by the ImageQuantTM TL software and is typically in the range of 1–60 s. Longer exposure times may be required if the antibody is inactivated.
Invert the membrane back and forth several times (at least two times) to ensure that sufficient substrate solution is spread over the membrane.
Take a picture with bright light to save an image with the position of the molecular weight markers on the membrane. It helps in the identification of the bands of interest.
Figure 1. Typical results of western blot following P-TRAP. Immunoprecipitation of the endogenous ribosome and ribosome-bound RNA with the anti-ribosomal protein P0 antibody (9D5) or isotype control IgG (NC) from the cytosolic lysate of HEK293 cells in the presence or absence of RNase A, followed by western blotting using several primary antibodies (P0, uL3, uS2, PABP4, eIF4A3, and CBP80). The input contained 1% of the lysate used for immunoprecipitation. Asterisks (*) indicate non-specific signals from the antibodies (9D5 or NC). This figure is cited with modifications from the original publication [10].
4sU metabolic RNA labeling and extraction of cytosolic RNA and ribosome-bound RNA from HEK293 cells
This section describes the procedure for the extraction of cytosolic RNA and ribosome-bound RNA from HEK293 cells subjected to thapsigargin (TPG) to induce endoplasmic reticulum (ER) stress and 4sU for metabolic RNA labeling.
Seed 1 × 106 HEK293 cells in 10 mL of DMEM supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin into a 10 cm dish and incubate overnight in a humidified incubator with 5% CO2 at 37 °C. Use one dish per sample.
Note: Although we present the procedure using 10 cm dishes, scaling down is acceptable.
Twenty-four hours after seeding cells, prepare the following 4sU labeling mix (Table 4) for inducing ER stress and 4sU metabolic RNA labeling.
Table 4. 4sU labeling mix
Reagent Final conc. (in 10 cm dish) Amount (per 1 sample)
Serum-free DMEM n/a 180 μL
200 mM 4sU (see Recipes) 200 μM 10 μL
DMSO or 1 mM TPG (see Recipes) 0.1% (v/v) or 1 μM 10 μL
Add labeling solution to the cultured medium (10 mL) and incubate for 3 h in a humidified incubator with 5% CO2 at 37 °C.
Aspirate the medium from the plate and add 1 mL of freshly prepared ice-cold 1% Triton X-100 lysis buffer (see Recipes).
Gently shake the 10 cm dish by hand to ensure that the lysis buffer covers the entire surface of each well, transfer to a 1.5 mL tube using a micropipette, and centrifuge at 20,000× g for 10 min at 4 °C, using a refrigerated centrifuge with a swing rotor.
Notes:
Work on ice to keep the 10 cm dishes and tubes cold.As the sample warms up and translation proceeds, the ribosomes will dissociate from the mRNA.
No need to use a cell scraper.
Collect the supernatant in a 1.5 mL tube.
Prepare the following 10× antibody mix with RNasin® (Table 5).
Table 5. 10× antibody solution with RNasin®
Reagent Concentration (10×) Amount per 1 sample
1% Triton X-100 lysis buffer (freshly prepared) n/a 34 μL
1 μg/mL anti-ribosomal proteins P0/P1/P2 mAb [9D5] 0.2 μg/μL 4 μL
RNasin® Plus ribonuclease inhibitor (40 units/μL) 10 units/μL 12 μL
For cytosolic RNA preparation, transfer 200 μL of the supernatant in a new 1.5 mL tube. Add 500 μL of ISOGEN II supplemented with 1 mM DTT and mix well. Store the prepared sample solution at -20 °C.
For ribosome-bound RNA preparation, transfer 450 μL of the supernatants in a new 1.5 mL tube. Add 50 μL of 10× antibody mix with RNasin to the supernatant (the final antibody solution will be 1×) and rotate at 4 °C for 60 min on an NRC-20D rotary mixer with an Oshiri Penpen A type.
Note: The NRC-20D with an Oshiri Penpen A type is a rotary mixer with a tube tapping function. Tapping the tube during stirring prevents the beads from sticking to the bottom of the tube and improves the efficiency of immunoprecipitation. However, it is not essential.
Dispense 28 μL of DynabeadsTM Protein G beads per sample into a new 1.5 mL tube and wash the beads with 1 mL of wash buffer (see Recipes). Insert the tube into the magnetic rack and remove the clear supernatant.
Transfer all of the sample solution to the Dynabeads tube and rotate at 4 °C for 40 min on an NRC-20D rotary mixer with an Oshiri Penpen A type.
Place the tube in a magnetic rack, remove the clear supernatant, and wash the beads three times with 1 mL of wash buffer (see Recipes). Each time, resuspend the beads in wash buffer using a micropipette with a siliconized pipette tip and transfer to a new 1.5 mL siliconized tube to allow adsorption of non-specific binding proteins to the tip and tube. Place the tube in a magnetic rack and remove the clear supernatant (wash buffer).
Note: The beads should be suspended in the wash buffer and the supernatant removed as rapidly as possible. Furthermore, care should be taken to prevent the beads from drying out during this process.
Add 500 μL of ISOGEN II supplemented with 1 mM DTT to the beads and resuspend to elute the immunoprecipitants. Place the tube in a magnetic rack and transfer the clear supernatant to a new 1.5 mL tube. Add 200 μL of nuclease-free water supplemented with 1 mM DTT to the transferred ISOGEN II solution and mix well. The prepared sample solution is stored at -20 °C.
RNA purification and alkylation
Note: The initial steps of RNA purification and alkylationmust be performedin the dark or protected from (white) light (e.g., by keeping samples covered, wrapping all tubes in aluminum foil, or working under red light).
Incubate the RNA samples (previously stored at -20 °C) at 37 °C for 5 min.
Vortex for 15 s. Stand tubes at RT for 10 min and centrifuge at 20,000× g for 10 min at RT with a swing rotor.
Transfer 500 μL of supernatants to the new 1.5 mL tubes and add 500 μL of 2-propanol supplemented with 1 mM DTT.
Vortex for 15 s and centrifuge at 20,000× g for 15 min at RT with a swing rotor.
Discard the supernatants by decantation, add 500 μL of 70% EtOH supplemented with 1 mM DTT, and centrifuge at 20,000× g for 3 min at RT with a swing rotor.
Repeat step D5 once.
Discard the supernatants by micropipetting.
Add 15 μL of nuclease-free water supplemented with 1 mM DTT to the RNA pellet and resuspend.
Measure RNA concentration by nanodrop.
Note: To assess the purity of RNA, it is recommended to check the ratio of absorbance at 260 and 280 nm (a ratio of ~2.0 is acceptable).
Prepare the following DMSO/NaPO4 solution (Table 6).
Table 6. DMSO/NaPO4 solution
Reagent Concentration Amount per 1 sample
DMSO n/a 25 μL
0.5 M NaPO4 pH 8.0 (see Recipes) 83 mM 5 μL
Notes: NaOP4 may form aggregates when added to DMSO. Prepare extra volume of DMSO/NaPO4 solution in a 1.5 mL tube (e.g., if you have 12 samples, prepare solution for 16 samples), centrifuge the mixed solution, and transfer only the clear supernatant to a new 1.5 mL tube. Use the supernatant for the following steps.
Prepare the following 4sU alkylation mix (Table 7).
Table 7. 4sU alkylation mix
Reagent Concentration Amount per 1 sample
Sample RNA ≤ 100 ng/μL X μL (≤ 5 μg)
Nuclease-free water n/a 15–X μL
IAA (100 mM) (see Recipes) 10 mM 5 μL
DMSO/NaPO4 solution n/a 30 μL
Notes:
The volume of input RNA should be less than 5 μg.
Use only freshly prepared IAA.
Incubate at 50 °C for 15 min with the DTU-1CN Dry Thermal Unit.
Add 1 μL of 1 M DTT, 1 μL of 20 mg/mg glycogen, 5 μL of 3 M NaOAc, and 125 μL of EtOH to each sample.
Note: After this step, exposure to light is possible.
Vortex for 15 s and centrifuge at 20,000× g for 30 min at 4 °C with a swing rotor.
Discard the supernatants by decantation, add 1 mL of 70% EtOH, and centrifuge at 20,000× g for 3 min at 4 °C with a swing rotor.
Discard the supernatants by decantation, add 1 mL of 70% EtOH, and centrifuge at 20,000× g for 3 min at RT with a swing rotor.
Discard the supernatants by micropipetting.
Add 25 μL of nuclease-free water to the pellet and resuspend.
Note: If the input RNA is less than 5 μg, the volume of nuclease-free water can be reduced (e.g., suspend the pellet in 12.5 μL of nuclease-free water if the input RNA is 2.5 μg).
Measure RNA concentration by nanodrop. Typical RNA concentrations are 100–200 ng/μL when using 5 μg of RNA as input.
Note: To assess the purity of RNA, it is recommended to check the ratio of absorbance at 260 nm and 280 nm (a ratio of ~2.0 is acceptable).
Assess the quality of the RNA with the fragment analyzer MultiNA.
Note: Instead of MultiNA, Bioanalyzer, TapeStation, or electrophoresis with denaturing TBE-agarose gel can be used.
Bring separation buffer and marker solution (reagents from the RNA 1000 kit), SYBR Green II stock solution, and RNA 6000 ladder to RT.
Dilute SYBR Green II stock solution 100-fold with TE buffer.
Prepare the required volume of the following MultiNA buffer solution (Table 8) in a 5 mL screw cap tube (included in the RNA 1000 kit).
Note: The required volume depends on the sample number. Prepare two more samples for the blank tube and RNA 6000 ladder.
Table 8. MultiNA buffer solution
Reagent Amount for 1 sample Amount for 20 samples
Separation buffer (a reagent of the RNA 1000 kit) 63.2 μL 1,264 μL
1/100 diluted SYBR Green II 0.8 μL 16 μL
Formamide 16 μL 320 μL
Prepare the following blank (Table 9), RNA 6000 ladder (Table 10), and RNA samples (Table 11) in 0.2 mL PCR tubes for the MultiNA run.
Table 9. Blank
Reagent Amount
RNA marker solution (a reagent of the RNA 1000 kit) 5 μL
RNA storage solution (a reagent of the RNA 1000 kit) 5 μL
Table 10. RNA 6000 ladder
Reagent Amount
RNA marker solution (a reagent of the RNA 1000 kit) 5 μL
RNA storage solution (a reagent of the RNA 1000 kit) 4 μL
RNA 6000 ladder 1 μL
Table 11. RNA sample
Reagent Amount (for 1 sample)
RNA marker solution (a reagent of the RNA 1000 kit) 5 μL
RNA storage solution (a reagent of the RNA 1000 kit) 4 μL
Sample RNA 1 μL
Incubate at 65 °C for 5 min in a thermal cycler and immediately place on ice.
Set the 5 mL tube (MultiNA buffer solution) and the 0.2 mL PCR tubes (blank, RNA 6000 ladder, and RNA samples) on MultiNA and start the run with MultiNA Control Software (Figure 2).
Figure 2. Quality control of RNA samples using a fragment analyzer.
Representative capillary electrophoresis profiles for cytosolic RNAs and immunoprecipitated RNAs from DMSO- or TPG-treated HEK293 cells were analyzed using the MultiNA. Three independent experiments were performed for each condition (#1, #2, and #3). The lower marker (LM) indicates internal standards (25 nt). RNA ladder (Agilent RNA 6000 Pico Kit) was used as a size marker. The MultiNA Viewer software automatically assesses RNA size and concentration. RNA quality is assessed by the ratio of 28S to 18S rRNA.
Sequencing
Prepare the sequencing library using the QuantSeq 3' mRNA-Seq Library Prep Kit FWD for Illumina, according to the manufacturer’s instructions. Use 500 ng of RNA as an input.
Assess the quantity of the libraries with the QubitTM dsDNA Quantification kit.
Prepare the required volume of the following Qubit working solution (Table 12) in a 1.5 mL tube.
Note: The required volume depends upon the sample number. Prepare 2 more samples for standards.
Table 12. Qubit working solution
Reagent Amount for 1 sample Amount for 20 samples
Qubit dsDNS HS Reagent (Component A) 50 μL 1,000 μL
Qubit dsDNS HS Reagent (Component B) 0.25 μL 5 μL
Mix the working solution with standards or samples (Table 13) in a 96-well black plate.
Note: Pipetting of samples should be done gently to avoid bubble formation.
Table 13. Qubit working solution with standards or samples
Reagent Amount for 1 sample
Qubit working solution (Table 12) 50 μL
Qubit dsDNS HS Standard (Component C or D) or cDNA sample 1 μL
Measure fluorescence using the SpectraMax Paradigm multi-mode microplate reader. The excitation wavelength is 485 nm and the emission wavelength is 530 nm. Measure each plate at least three times to check the stability of the fluorescence in each well.
Determine the concentration of each library and pool them in an equimolar ratio in a 1.5 mL tube for multiplex sequencing.
Sequence samples on Illumina HiSeq system. Sequence so that there are at least 1 million counted T>C reads per sample (of course, the more the better).
Notes:
We recommend longer sequencing read lengths (e.g., ≥ 150 bp) to increase the sensitivity for detecting T>C conversions in bioinformatic analysis.
If you run the sequence in pair-end mode, you will get two fastq files, read 1 (FWD) and read 2 (REV). In this case, use only read 1 (FWD) for analysis, because read 2 (REV) will start sequencing with the poly(T) strand on the 3' side and sequence through the homopolymer strand, reducing the quality of the sequence reads.
Support for the Illumina HiSeq system is being phased out. It is recommended to use newer platforms such as the NovaSeq system.
Data analysis
We performed all sequencing data analysis steps under Ubuntu 20.04 LTS. Total and nascent RNAs were quantified using SLAM-DUNK, a pipeline for SLAMseq data analysis.
Make sure that SLAM-DUNK is installed on your Linux system. If not, install the latest version of SLAM-DUNK according to the document (https://t-neumann.github.io/slamdunk/index.html).
Prepare genome reference. For example, human reference FASTA files are prepared as follows:
```bash
Wget http://ftp.ebi.ac.uk/pub/databases/gencode/Gencode_human/latest_release/GRCh38.p13.genome.fa.gz
gunzip GRCh38.p13.genome.fa.gz
```
Prepare a BED file with 3' UTR coordinates. This can be downloaded from the UCSF Table Browser (https://genome.ucsc.edu/cgi-bin/hgTables). Alternatively, you can create a BED file from the annotation file as follows, using the Python script “create_bed.py” provided as a supplemental file.
Note: This Python script requires “pandas” and “gffpandas”. Place the annotation file and “create_bed.py” in the same directory and run it.
```bash
wget https://ftp.ebi.ac.uk/pub/databases/gencode/Gencode_human/release_41/gencode.v41.annotation.gff3.gz
gunzip ge ncode.v41.annotation.gff3.gz
python generate_BED.py
```
Run “slamdunk”. Here, 12 bases from the 5' end were trimmed (--trim-5p 12), and then five or more consecutive adenines from the 3' end were considered the remaining poly(A) tail and removed (--max-polya 4). Up to 100 regions with multiple mapped reads were allowed (--topn 100). For details on detecting T>C conversions, refer to the original paper [12].
```bash
slamdunk map --trim-5p 12 --max-polya 4 --topn 100 --skip-sam --reference --outputDir slamdunk/map
slamdunk filter --outputDir slamdunk/filter slamdunk/map/*.bam
slamdunk snp --outputDir slamdunk/snp --reference slamdunk/filter/*.bam
slamdunk count --reference --bed --outputDir slamdunk/count --snp-directory slamdunk/snp slamdunk/filter/*.bam
```
Note: It is recommended that FASTQ files be trimmed and quality checked with a trimming tool, such as fastp [15], and a quality control tool, such as FastQC [16], before running “slamdunk”. In addition, tools such as MultiQC [17] can be used to summarize various parameters such as mapping quality and T>C conversion rate to obtain reliable results.
Perform differential gene expression analysis using DESeq2 and differential transcription and translation analysis using the deltaTE method for nascent RNA (Figure 3).
Note: For DESeq2 and deltaTE analyses, use the ReadCount (total read count) and TcReadCount (T>C read count) columns of the tcount file (output from “slamdunk count”) as input. To analyze the TcReadCount column with the DESeq2 and deltaTE method, calculate global normalization factors using the ReadCount column.
Figure 3. Differentially expressed and translated nascent RNA analysis using DESeq2 and deltaTE. Result table (first five rows) (A) and MA plot (B) of nascent RNA obtained by P-TRAP-seq from TPG-treated vs. DMSO-treated HEK293 cells. Results from DESeq2 using the ReadCount and TcReadCount columns as inputs in the "slamdunk" output file are plotted. Differentially expressed genes (adjusted p-value < 0.05 and log2 fold change > |1.5|) are highlighted in red (up-regulated) or blue (down-regulated). (C) Fold change of nascent RNAs in cytosolic RNA-seq and P-TRAP-seq data at the level of the individual genes is analyzed by the deltaTE method using the ReadCount and TcReadCount columns as inputs in the "slamdunk" output file (FDR < 0.05 and log2 fold change > |0|). Regulation modes categorized by deltaTE analysis as described [14]. [Translationally forwarded genes (cyan), intensified genes (blue), exclusive genes (red), and buffered genes (purple) are highlighted.] These figures are cited with modifications from the original publication [10].
To analyze the translation of pre-existing RNA, calculate the non-T>C read count by subtracting the T>C read count from the total read count.
Notes:
The non-T>C reads are expected to include some reads that are derived from nascent RNAs but do not contain T>C conversion. If the proportion of non-T>C reads derived from these nascent RNAs is high, the estimation of the proportion of pre-existing RNAs based on the subtracted read counts will be inaccurate. Therefore, it is recommended to evaluate the sensitivity of the detection of T>C reads under each experimental condition. For example, focus on genes (CHAC1, DDIT3, and HERPUD1 in the UPR, etc.) that are known to be significantly transcribed after the addition of TPG, an inducer of ER stress. This is because most of the reads from these genes are likely to be derived from nascent RNAs. By calculating the proportion of reads detected as T>C reads out of all these reads, the sensitivity of the detection of T>C reads is improved (Figure 4).
To analyze the non-T>C read count with the DESeq2 and deltaTE method, calculate global normalization factors using the total read count column.
Figure 4. Evaluation of the detection sensitivity of T>C reads under each experimental condition. A. Schematic representation of nascent RNA synthesis of massively transcribed genes in the UPR and its 4sU incorporation. A certain percentage of RNAs may not be labeled by 4sU even though they are nascently transcribed. The proportion of 4sU labeling in RNA that is transcribed after thapsigargin treatment is used to assess detection sensitivity. B. Raw total read counts and raw T>C read counts of selected genes in cytosolic RNA-seq in response to DMSO (black) or thapsigargin (TPG) (red) treatment. The means and standard deviations of three replicates are shown. The proportion of T>C reads out of the total reads increased by TPG treatment for stress-induced genes (CHAC1, DDIT3, and HERPUD1) but not control genes (ACTB, RPLP0, and EEF2). These figures are cited with modifications from the original publication [10].
For further analysis, output of DESeq2 (e.g., Log2FC) can be used to generate a cumulative curve to analyze the expression of nascent and pre-existing RNAs in specific gene categories (Figure 5).
Figure 5. Categorical analysis of the gene expression of the nascent and pre-existing RNAs. Cumulative distributions of the Log2FC calculated by DESeq2 of specific gene categories, preferentially translated by polysome upon eIF2α phosphorylation [18] (A) and ATF6/XBP1 transcriptional targets [19,20] (B) compared to all genes, together with p-values of Mann–Whitney U test. These figures are cited with modifications from the original publication [10].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Imai et al. [10]. Simultaneous measurement of nascent transcriptome and translatome using 4-thiouridine metabolic RNA labeling and translating ribosome affinity purification. Nucleic Acids Res.
Acknowledgments
This work was supported by JSPS KAKENHI (21K15015 and 24K18056 to H.I., 21K19413 and 24K02229 to A.Y.) and the Japan Science and Technology Agency (JPMJMS2022 to A.Y.). This work was also supported by research grants from the Manufacturing Promotion Division, Department of Commerce, Industry, and Labor, Okinawa Prefectural Government, Takeda Science Foundation, and Bristol Myers Squibb K.K. We thank Ms. Naima Batool for editing a draft of the manuscript. This protocol was described and validated in original paper [10].
Competing interests
The authors declare no competing interests.
References
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Baptista, M. A. P. and Dölken, L. (2018). RNA dynamics revealed by metabolic RNA labeling and biochemical nucleoside conversions. Nat Methods. 15(3): 171–172. https://doi.org/10.1038/nmeth.4608
Rummel, T., Sakellaridi, L. and Erhard, F. (2023). grandR: a comprehensive package for nucleotide conversion RNA-seq data analysis. Nat Commun. 14(1): 3559. https://doi.org/10.1038/s41467-023-39163-4
Schwanhäusser, B., Busse, D., Li, N., Dittmar, G., Schuchhardt, J., Wolf, J., Chen, W. and Selbach, M. (2011). Global quantification of mammalian gene expression control. Nature. 473(7347): 337–342. https://doi.org/10.1038/nature10098
King, H. A. and Gerber, A. P. (2014). Translatome profiling: methods for genome-scale analysis of mRNA translation. Briefings Funct Genomics. 15(1):22–31. https://doi.org/10.1093/bfgp/elu045
Kawata, K., Wakida, H., Yamada, T., Taniue, K., Han, H., Seki, M., Suzuki, Y. and Akimitsu, N. (2020). Metabolic labeling of RNA using multiple ribonucleoside analogs enables the simultaneous evaluation of RNA synthesis and degradation rates. Genome Res. 30(10): 1481–1491. https://doi.org/10.1101/gr.264408.120
Jürges, C., Dölken, L. and Erhard, F. (2018). Dissecting newly transcribed and old RNA using GRAND-SLAM. Bioinformatics 34(13): i218–i226. https://doi.org/10.1093/bioinformatics/bty256
Schott, J., Reitter, S., Lindner, D., Grosser, J., Bruer, M., Shenoy, A., Geiger, T., Mathes, A., Dobreva, G., Stoecklin, G., et al. (2021). Nascent Ribo-Seq measures ribosomal loading time and reveals kinetic impact on ribosome density. Nat Methods. 18(9): 1068–1074. https://doi.org/10.1038/s41592-021-01250-z
Hetz, C., Zhang, K. and Kaufman, R. J. (2020). Mechanisms, regulation and functions of the unfolded protein response. Nat Rev Mol Cell Biol. 21(8): 421–438. https://doi.org/10.1038/s41580-020-0250-z
Imai, H., Utsumi, D., Torihara, H., Takahashi, K., Kuroyanagi, H. and Yamashita, A. (2023). Simultaneous measurement of nascent transcriptome and translatome using 4-thiouridine metabolic RNA labeling and translating ribosome affinity purification. Nucleic Acids Res. 51(14): e76–e76. https://doi.org/10.1093/nar/gkad545
Okada-Katsuhata, Y., Yamashita, A., Kutsuzawa, K., Izumi, N., Hirahara, F. and Ohno, S. (2011). N- and C-terminal Upf1 phosphorylations create binding platforms for SMG-6 and SMG-5:SMG-7 during NMD. Nucleic Acids Res. 40(3): 1251–1266. https://doi.org/10.1093/nar/gkr791
Neumann, T., Herzog, V. A., Muhar, M., von Haeseler, A., Zuber, J., Ameres, S. L. and Rescheneder, P. (2019). Quantification of experimentally induced nucleotide conversions in high-throughput sequencing datasets. BMC Bioinf. 20(1): 258. https://doi.org/10.1186/s12859-019-2849-7
Love, M. I., Huber, W. and Anders, S. (2014). Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15(12): 550. https://doi.org/10.1186/s13059-014-0550-8
Chothani, S., Adami, E., Ouyang, J. F., Viswanathan, S., Hubner, N., Cook, S. A., Schafer, S. and Rackham, O. J. L. (2019). deltaTE: Detection of Translationally Regulated Genes by Integrative Analysis of Ribo‐seq and RNA‐seq Data. Curr Protoc Mol Biol. 129(1): e108. https://doi.org/10.1002/cpmb.108
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Supplementary information
The following supporting information can be downloaded here:
gencode.v41.customized.canonical.bed
generate_bed.py
Article Information
Publication history
Received: Jun 27, 2024
Accepted: Aug 30, 2024
Available online: Sep 25, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
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Molecular Biology > RNA > mRNA translation
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5,092 | https://bio-protocol.org/en/bpdetail?id=5092&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Generation of Zebrafish Maternal Mutants via Oocyte-Specific Knockout System
CZ Chong Zhang *
WW Wenlu Wei *
TL Tong Lu
YZ Yizhuang Zhang
JL Jiaguang Li
JW Jiasheng Wang
AC Aijun Chen
FW Fenfen Wen
MS Ming Shao
(*contributed equally to this work)
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5092 Views: 274
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Original Research Article:
The authors used this protocol in Biology (Basel) Aug 2021
Abstract
Maternal mRNAs and proteins are produced during oogenesis by more than 60% of zebrafish genes. They are indispensable for fertilization and early embryogenesis. Generation and analysis of the maternal mutant is the most direct way to characterize the maternal function of the specific gene. However, due to the lethality of zygotic mutants, the maternal function of most genes in zebrafish remains elusive. Several methods have been developed to circumvent this obstacle, including mRNA rescue, germ-line replacement, oocyte microinjection in situ, mosaic mutation, and bacterial artificial chromosome (BAC)-mediated conditional rescue. Here, we provide an alternative approach to generate zebrafish maternal mutants rapidly and efficiently by introducing four tandem sgRNA expression cassettes into Tg(zpc:zcas9) embryos. This method is more technically feasible and cost- and time-effective than other established methods.
Key features
• This protocol can circumvent the lethality or infertility of the zygotic mutants to obtain maternal mutants of the target gene.
• This protocol is time-saving (one fish generation).
• Using this protocol, double-gene maternal mutants can be obtained in a single generation.
• Stable lines can be established to continuously produce maternal mutant embryos for the gene of interest.
Keywords: Zebrafish Maternal mutant Conditional knockout Oocyte Cas9 sgRNA
Graphical overview
Background
Maternal factors are mRNAs and proteins deposited in the oocyte. They play vital roles in oocyte maturation, fertilization, and blastocyst development. Approximately 66% of zebrafish genes are expressed maternally [1]. As zygotic genome activation (ZGA) starts at around the 1k-cell stage, maternal factors are dominant to function in the developmental events before this time point. In addition, even after zygotic genome activation (ZGA), maternal factors still function in later embryogenesis, including axis formation, germ layers differentiation, morphogenesis, etc. [2–4].
Generation of the corresponding maternal mutant is the most straightforward way to address the maternal function of a gene in zebrafish. The genotype of primary oocyte is the same as that of somatic cells. Hence, viable and fertile female zygotic mutants are indispensable for giving birth to maternal mutants or maternal and zygotic mutants. However, zygotic mutants usually cannot survive to adulthood or spawn because of the crucial and uncompensable functions of the mutated genes [5–7]. This obstacle represents the major bottleneck to studying maternal factors. Several methods have been developed to bypass this technical hurdle. The first is mRNA rescue. Through microinjection of in vitro–synthesized wild-type mRNA, the zygotic mutant can be rescued to live through a critical period, in which the zygotic products contribute to normal development [4]. Considering that the ectopic expression may disrupt normal embryogenesis and transient expression of mRNA cannot support long-term gene function, this method is not applicable to most genes. In 2002, Ciruna et al. developed the germ-line replacement technology. They injected dead end1 morpholino into wild-type embryos to block primordial germ cell development and then transplanted germ cells from zygotic mutants into these primordial germ cell (PGC)-free hosts. This ensured that the germ lines of the host were entirely replaced by donor PGCs. Upon maturation, all the embryos laid by the female chimera were maternal mutants [8]. Transplantation of PGCs is technically demanding and the chimeric embryos tend to develop into males because of the limited number of transplanted PGCs. Wu et al. established a surgery-based method [9]. Following the opening of the female fish abdomen, the fluorescent lineage tracer and the morpholino targeting the specific gene were co-injected into the stage- oocyte. After recovery, the laid fluorescent embryos were the desired maternal morphants [9]. However, the intricate operative skills required for the application of this method are significantly challenging. In 2018, Xing et al. generated mosaic mutants containing homozygous mutant cells through secondary genome editing in heterozygotes [2]. This approach was designed to circumvent the lethality associated with double zygotic mutants of dvl2 and dvl3a. Some of these homozygous mutant cells were incorporated into the germline, eventually resulting in maternal mutants after fertilization. However, this method is inefficient and time-consuming, requiring at least three generations. In addition to the methods mentioned above, the bacterial artificial chromosome-rescue-based knockout (BACK) is another method to bypass the zygotic lethality and obtain maternal mutants [10]. To obtain such mutants, first, the authors generated mutants of the target gene through nuclease-mediated genome editing. Second, they rescued the zygotic mutants by introducing a bacterial artificial chromosome (BAC) containing the target gene expression cassette flanked by loxP sites via Tol2-mediated transgenesis. Third, they crossed these fish with a germline-specific Cre line. This approach ultimately allowed them to obtain maternal mutants of the target gene. However, this strategy is time-consuming and labor-intensive, as it involves gene knock-out, transgenesis, and crossing with the Cre line.
Recently, we have developed a new strategy to generate single or double-gene maternal mutants through transgenic expression of Cas9 and sgRNA in oocytes [11,12]. Through I-Sce I-mediated transgenesis, we introduced an eGFP reporter and multiple tandem sgRNA expression cassettes into Tg(zpc:zcas9) transgenic fish, where Cas9 is specifically expressed in oocytes. This approach enables genome editing during the early stages of oogenesis. As a result, some of the GFP-positive embryos will have their maternal products completely ablated, thereby becoming maternal mutants. Consequently, we can analyze their phenotypes after identifying them through genotyping. Compared to other methods, it has several advantages. First, it is technically feasible. The main techniques are plasmid construction and transgenesis, which are conventional in most zebrafish labs. Second, it is time saving, taking one generation (2–3 months) for either single or double-genes maternal knockout. Third, the efficiency is generally stable despite individual variation. Through generating maternal mutants of nanog, ctnnb2, rbm24a, dvl2, and dvl3a, we find that the average ratio of the maternal mutant for a single gene is approximately 25% and the maximum efficiency can reach 63.3%. For every single founder fish, the ratio of maternal mutants remains stable in each spawning, so that we can obtain maternal mutants repeatedly once we get a founder. Finally, this approach can be utilized to generate double-gene maternal mutants, which takes nearly the same time.
Materials and reagents
Biological materials
Zebrafish lines:
Wild-type AB zebrafish line was obtained from the China zebrafish resource center
Tg(zpc:zcas9) transgenic line was reported by Ming Shao’s lab and is available upon request [13]
Reagents
NaCl (Solarbio, catalog number: S8210)
KCl (Solarbio, catalog number: P9921)
CaCl2·2H2O (Solarbio, catalog number: C8370)
HEPES (Sigma-Aldrich, catalog number: H3375)
2× Taq Master Mix (Dye Plus) (Vazyme, catalog number: P112-01)
AxyPrep PCR Clean-Up Kit (Axygen, catalog number: AP-PCR-250)
T7 RNA polymerase, 5× transcription buffer (Thermo Scientific, catalog number: EP0111)
ATP/CTP/GTP/UTP, 10 mM each (Thermo Scientific, catalog number: R0481ribo); dilute the 100 mM solution with DNase/RNase-free H2O
RiboLock RNase inhibitor (Thermo Scientific, catalog number: EO0381)
TURBO DNase (Invitrogen, catalog number: AM2238)
Ammonium acetate, 5 M, RNase-free (Thermo Scientific, catalog number: AM9070G)
Phenol:chloroform:isoamyl alcohol 25:24:1, pH 5.2 (Thermo Scientific, catalog number: J62336.AE)
Cas9 protein: GenCrispr NLS-Cas9-NLS nuclease (GenScript, catalog number: Z03389-50)
AxyPrep Plasmid Miniprep Kit (Axygen, catalog number: AP-MN-P-250)
Acc65I (Asp718I) (Thermo Scientific, catalog number: ER0901)
Gibson assembly kit: Hieff Clone Plus One Step Cloning Kit (Yeasen, catalog number: 10911ES20)
CutSmart buffer (NEB, catalog number: B60004)
Agarose (BIOWEST, catalog number: 111860)
Glass capillaries (World Precision Instruments, catalog number: TWF-100F-4)
100× penicillin-streptomycin solution (Gibco, catalog number: 15140122)
cDNA synthesis kit (Transgene, catalog number: AT301)
pBackZero-T vector (Takara, catalog number: 3275)
Primers:
Universal primer:
5'-AAAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGC
TATTTCTAGCTCTAAAAC-3'
Dest forward primer: 5'-TTCTTGTTTAAGCTTTTAATCTCAAAAAAC-3'
Dest reverse primer. 5'-GGCTGTTTACATCTGATAGTGG-3'
Ligation forward primer: 5'-gagtcggtgctttttttaaacctggTTCTTGTTTAAGCTTTTAATCTCAAAAAAC-3'
Ligation reverse primer: 5'-ATCCTGCACTGAATGCAC-3'
bmp2b sgRNA forward primer: 5'- taatacgactcactataGGGAGGCTGAGAGCAACCGGgttttagagctagaa-3'
M13 forward: 5'-GTAAAACGACGGCCAGT-3'
The plasmid system is available upon request to the authors Chong Zhang and Ming Shao
TRIzol (Thermo Scientific, catalog number: 15596026)
PBST (Solarbio, catalog number: P1031)
Chloroform (Sinopharm Chemical Reagent, catalog number: 10006862)
Isopropanol (Sinopharm Chemical Reagent, catalog number: 40064360)
Glycogen (Thermo Scientific, catalog number: R0561)
Solutions
Ringer's buffer (see Recipes)
Recipes
Ringer's buffer
Reagent Final concentration Quantity
NaCl 116 mM 6.779 g
KCl 2.9 mM 0.216 g
CaCl2·2H2O 1.8 mM 0.265 g
HEPES 5 mM 1.192 g
H2O n/a 1,000 mL
Total n/a 1,000 mL
Dissolve all ingredients in 900 mL of deionized H2O. Adjust pH to 7.2 using NaOH. After being autoclaved, it can be long-term stored at room temperature.
Equipment
VeritiTM Dx 96-well thermal cycler (Thermo Fisher, catalog number: 4452300)
NanoDrop 2000 spectrophotometer (Thermo Scientific, model: ND-2000)
Block heater (Yiheng, model: TU-100C)
Electrophoresis system (Bio-Rad, catalog number: 1640302)
Centrifuge (Eppendorf, model: 5424R)
Pico-injector (Warner Instrument, model: PLI-100A)
Puller (NARISHIGE, model: PC-100)
Tweezer (WPI, catalog number: 500341)
Microforge (NARISHIGE, model: MF2)
Microinjector (Harvard Apparatus, model: PLI-100A)
Software and datasets
CRISPRScan, http://www.crisprscan.org/, for designing sgRNAs with high on-target activity and minimal off-target effects
Synthego, https://ice.synthego.com/#/, for analyzing sequencing results and assessing CRISPR editing efficiency
Procedure
The summary of this protocol is illustrated in Figure 1.
Figure 1. Main steps of this protocol
Construction of sgRNA expression plasmid
Prediction of sgRNA targeting sites.
For single-gene maternal knockout, we need 3–4 sgRNAs targeting different sites with high genome editing efficiency. To achieve this, we predict at least six different sgRNA targeting sites using the online CRISPRscan software (http://www.crisprscan.org/) [14]. After submitting the gene name, the website will predict several sgRNAs (Figure 2). The CRISPRscan model was designed and tested on zebrafish [15], and the authors recommend the following rules for selecting appropriate sgRNAs:
1) High CRISPRscan score: The CRISPRscan score represents the potential activity of the sgRNA. A score of at least 55 is required, with a score above 70 being recommended.
2) Low CFD (cutting frequency determination) score: The CFD score indicates the potential off-target cutting efficiency, so a lower CFD score is preferable.
In addition to these criteria, we also recommend the following rules for achieving higher knockout efficiency:
1) Avoid overlaps of selected sgRNA-targeting sites: Independent targeting sites increase the probability of successful gene disruption.
2) Select sites close to the start codon ATG: Targeting the front two-thirds of the coding sequence is recommended, as frameshift mutations closer to ATG will more thoroughly disrupt gene function. The Oligos containing the T7 promoter, the predicted sgRNA sequence, and a 16 nt sgRNA scaffold sequence are synthesized for amplification of the sgRNA IVT DNA template.
Figure 2. Example output from CRISPRScan. Zebrafish-Danio rerio, Cas9-NGG, Gene, in vitro T7 promoter, and 4 mismatches should be chosen, and the gene symbol is then submitted to “Get sgRNAs”. This protocol uses Cas9 for genome editing, so we select Cas9-NGG to specify the prediction of sgRNA for Cas9. Allowing for “4 mismatches” enables the sgRNA site to tolerate up to four mismatches in off-target sequences. This criterion helps to identify a broader range of potential target sites for consideration.
Amplification and purification of each sgRNA DNA template.
Prepare the Fill-in PCR mixture on ice to ensure enzyme stability and activity, as shown in Table 1.
Table 1. PCR reaction for amplification of sgRNA DNA template
Reagent Volume
2× Taq Master Mix (Dye Plus) 25 μL
sgRNA primer (10 μm) 2 μL
Universal primer (10 μm) 2 μL
Deionized H2O 21 μL
Total volume 50 μL
Put the mixture into the thermocycler and run the program as shown in Table 2.
Table 2. PCR program for amplification of sgRNA DNA template
Temperature Time Cycle number
94 °C 3 min 1 cycle
94 °C 15 s 45 cycles
50 °C 15 s
72 °C 30 s
72 °C 5 min 1 cycle
Take 2 μL of PCR products and subject them to electrophoresis; the fragment size should be 117 bp.
Purify the rest of the PCR products with the AxyPrep PCR Clean-Up Kit. Follow the manual book and elute the DNA with 20 μL of deionized H2O.
Measure the concentration of the purified DNA template using a NanoDrop 2000 spectrophotometer. Ensure that the DNA concentration is within the optimal range for subsequent steps and store the DNA at -20 °C until use.
Note: To meet the requirement of the following transcription, scale up the PCR mixture to make the final concentration of purified DNA higher than 40 ng/μL.
In vitro transcription of sgRNA.
Thaw frozen reagents, mix thoroughly, and centrifuge briefly. Place the reaction buffer at room temperature and other components on ice.
Set up the in vitro transcription reaction mixture on ice to maintain enzyme activity and prevent degradation, as shown in Table 3.
Table 3. In vitro transcription reaction for synthesis of sgRNA
Reagent Volume
5× Transcription buffer 4 μL
ATP/GTP/CTP/UTP, 10 mM each 4 μL
DNA template 400 ng
RNase inhibitor 0.5 μL
T7 RNA polymerase 1 μL
DNase/RNase-free H2O to 20 μL
Incubate at 37 °C for 2 h.
To remove the DNA template, add 1 μL of TURBO DNase (2 U/μL). Mix, briefly centrifuge, and incubate at 37 °C for 20 min.
Add 10 μL of ammonium acetate (5 M, RNase-free) and vortex to stop the reaction.
sgRNA purification and precipitation.
Add 70 μL of RNase-free H2O and mix thoroughly. Then, add an equal volume of phenol:chloroform:isoamyl alcohol (25:24:1, pH 5.2). Vortex the mixture briefly and centrifuge at 12,000× g for 30 s to ensure proper mixing and pellet any debris. Transfer the aqueous phase to a new tube.
Add an equal volume of chloroform (100 μL). Vortex for 5 s and centrifuge at 12,000× g for 30 s. Transfer the aqueous phase to a new tube.
Add an equal volume of isopropanol (100 μL) and mix well.
Chill the mixture for at least 30 min at -20 °C or 5 min at -80 °C. Maintaining a lower temperature and extending the incubation time can improve sgRNA precipitation efficiency. Centrifuge at 12,000× g for 15 min at 4 °C to pellet sgRNA. Remove the supernatant liquid carefully (we can preserve some liquid to avoid taking sgRNA incidentally).
Rinse the sgRNA pellet with 1 mL of pre-chilled 70% ethanol. Centrifuge at 12,000× g for 15 min at 4 °C and discard the liquid supernatant.
Dissolve sgRNA with 10 μL of RNase-free H2O. Measure the concentration with NanoDrop 2000 spectrophotometer and dilute sgRNA to 100 ng/μL by adding RNase-free H2O. Aliquot the sgRNA to 1 μL per tube and store at -80 °C.
Note: Scale up the transcription mixture to yield more sgRNA.
Test sgRNA efficiency.
Dilute Cas9 protein to 200 ng/μL (add 1 μg/μL stock solution to 4 μL of RNase-free H2O and mix well). It can be stored at -20 °C for at least 30 days.
Mix 1 μL of sgRNA with 1 μL of Cas9 protein by pipetting gently on ice. The final concentrations should be 50 ng/μL for sgRNA and 100 ng/μL for Cas9 protein. Ensure thorough mixing to form sgRNA-Cas9 complexes.
Inject 2 nL mixture into each 1-cell stage embryo.
At 24 h post-fertilization (hpf), collect 10 embryos into a single microcentrifuge tube and carefully remove the medium. Use a pipette to ensure embryos are not damaged during collection. Then, add 100 μL of NaOH (50 mM, 10 μL each embryo) and heat at 95 °C in a metal bath or thermocycler for 15 min. Then, neutralize the lysate with 1/10 volume of Tris pH 8.0 (10 μL, 1 M).
Directly use the embryo lysate as the DNA template for PCR amplification of the fragment containing the sgRNA targeting site. Ensure the lysate is thoroughly mixed before use to avoid uneven template distribution.
Note: The flanking sequence of the sgRNA targeting site should be over 100 bp. After electrophoresis to ensure the proper amplification, we subject PCR products to Sanger sequencing, performing gel extraction when needed.
Analyze the sequencing results using the ICE (Inference of CRISPR Edits) web tool at https://ice.synthego.com/#/. Using this tool, we could quantify the sgRNA efficiency [16] to select the highly efficient ones. Upload the sequencing data and follow the tool's instructions for accurate analysis of editing efficiency The sgRNA efficiency will be normalized with uninjected group sequencing data. Choose the sgRNA with an indel percentage ≥ 30% for the following steps (Figure 3).
Figure 3. Example output of ICE analysis. We consider the value of “indel %” as the efficiency of the sgRNA.
Construction of separate sgRNA expression plasmids (Figure 4).
Figure 4. Schematics illustrating the construction of U6 sgRNA expression vector. All U6 vectors have a U6 promoter and a sgRNA scaffold. Digestion of BsmBI makes U6 vectors expose the overhangs recognizing the annealed sgRNA primer. Digestion of PstI and SalI can disrupt the vector undigested by BsmBI to improve positive cloning.
According to the sgRNA sequence, synthesize sgRNA annealing primer through a commercial company.
Forward primer: TTCGN(19)
Reverse primer: AAAC[N(19)]*
GN(19) is the target sequence of sgRNA, starting as GG. Specifically, sgRNA primer given by CRISPRScan:
5'-TAATACGACTCACTATA[GN(19)]GTTTTAGAGCTAGAA-3'
[N(19)]* = Reverse complement of N(19).
For example, TAATACGACTCACTATAGGCGGGAGTGCGTGCAACACGTTTTAGAGCTAGAAATAGC represents the long primer sequence provided in the table, with the target sequence (protospacer) highlighted in bold. Therefore, the forward primer is ttcGGCGGGAGTGCGTGCAACAC, and the reverse primer is aaacGTGTTGCACGCACTCCCGC. The sequence italicized and in bold is the reverse complement of the forward primer sequence underlined.
Annealing of primers.
i. Set up the reaction as shown in Table 4.
Table 4. Reaction for annealing of sgRNA oligos
Reagent Volume
Forward primer (100 μM) 1 μL
Reverse primer (100 μM) 1 μL
10× NEB buffer 2.1 2 μL
Deionized H2O 16 μL
ii. Put the mixture into a thermocycler and run the program as shown in Table 5.
Table 5. Program for annealing of sgRNA oligos
Temperature Time
95 °C 15 min
50 °C 10 min (ramp rate: 0.1 °C/s)
4 °C 10 min
Ligation to U6 vectors.
i. The number of sgRNAs in the final transgenic plasmid determines the choice of U6 vectors [17] and following pGGDestISceIEG-XsgRNA. We list each combination in Table 6.
Table 6. Combination of U6 vectors and pGGDestISceIEG-XsgRNA for different number of sgRNAs
sgRNA number U6 vectors pGGDestISceIEG-XsgRNA
1 pU6a:sgRNA#1 pGGDestISceIEG-1sgRNA
2 pU6a:sgRNA#1, pU6a:sgRNA#2 pGGDestISceIEG-2sgRNA
3 pU6a:sgRNA#1, pU6a:sgRNA#2, pU6b:sgRNA#3, pGGDestISceIEG-3sgRNA
4 pU6a:sgRNA#1, pU6a:sgRNA#2, pU6b:sgRNA#3, pU6c:sgRNA#4 pGGDestISceIEG-4sgRNA
To achieve high efficiency of gene disruption, four different sgRNAs are recommended. We ligate these four different sgRNAs to pU6a:sgRNA#1, #2, #3, #4, which contain different U6 promoters driving sgRNA expression and sgRNA scaffold. Even if we do not have four different sgRNAs, we can ligate single sgRNA repeatedly into different U6 vectors. Then, we assemble these four U6 vectors containing four different sgRNA oligos into pGGDestISceIEG-4sgRNA.
ii. Ligate sgRNA oligos to corresponding U6 vectors (Figure 4). Set up the ligation reaction as shown in Table 7.
Table 7. Reaction for ligating sgRNA oligos to U6 vectors
Reagent Volume
10× Cutsmart buffer 1 μL
10× T4 ligase buffer 1 μL
T4 ligase 0.3 μL
BsmB 0.3 μL
Pst 0.2 μL
Sal 0.2 μL
H2O 1 μL
Annealed sgRNA primer 1 μL
pU6x:sgRNA#x (20 ng/μL) 5 μL
Total volume 10 μL
iii. Place the mixture into the thermocycler and run the program as shown in Table 8.
Table 8. Program for ligating sgRNA oligos to U6 vectors
Temperature Time Cycle number
37 °C 20 min 6 cycles
16 °C 15 min
37 °C 10 min 1 cycle
55 °C 15 min 1 cycle
80 °C 15 min 1 cycle
iv. Transformation.
1). Thaw DH5α competent cells on ice. Add the ligation mixture to 50–100 μL of DH5α cells and gently mix by flicking the tube. Avoid vortexing to maintain cell viability.
2). Incubate on ice for 30 min.
3). Heat shock at 42 °C for 60 s.
4). Chill on ice for 2 min.
5). Spread the mixture on a solid medium plate containing 100 μg/mL spectinomycin.
6). Culture at 37 °C for more than 16 h.
v. Colony PCR.
1). Pick a single bacterial colony using a sterile pipette tip and rinse it into 10 μL of deionized H2O. Ensure the colony is fully resuspended for subsequent PCR analysis.
2). Set up the PCR reaction as shown in Table 9.
Table 9. PCR reaction for identification of positive colony
Reagent Volume
2× Taq Master Mix (Dye Plus) 5 μL
Colony liquid 2 μL
M13 forward (10 μm) 0.4 μL
Reverse primer (10 μm) 0.4 μL
Deionized H2O 2.2 μL
Total volume 10 μL
3). Place the mixture into the thermocycler and run the program as shown in Table 10.
Table 10. PCR program for identification of positive colony
Temperature Time Cycle number
94 °C 3 min 1 cycle
94 °C 15 s 23 cycles
55 °C 15 s
72 °C 30 s
72 °C 5 min 1 cycle
4). Subject PCR products to electrophoresis. The PCR product size of the positive colony is 482 bp.
vi. Bacteria growth and extraction of plasmids.
1). Add the rest of the colony liquid into 3 mL of LB medium containing 100 μg/mL spectinomycin.
2). Shake at 200 rpm and 37 °C for 14 h.
3). Extract plasmids using the AxyPrep Plasmid Miniprep Kit and measure the concentration.
4). Subject plasmids to Sanger sequencing with M13 forward and align with the U6 vector map.
Construction of pGGDestISceIEG-XsgRNA vector.
According to the number of constructed sgRNA vectors, choose the appropriate pGGDestISceIEG-XsgRNA. Different pGGDestISceIEG-XsgRNA can receive different sets of sgRNA expression cassettes. Figure 5A shows an example of four sgRNA U6 vectors ligation to pGGDestISceIEG-4sgRNA through Golden Gate cloning.
Figure 5. Golden Gate cloning of tandem sgRNA expression vector and Gibson assembly of double-genes sgRNA expression cassettes. A. BsaI digestion leaves specific overhangs, which are designed for tandem ligation into pGGDestISceIEG-4sgRNA. B. The amplified sgRNA expression cassette targeting gene b can be cloned into the vector of gene a through Gibson assembly.
Set up the Golden Gate reaction mixture as shown in Table 11.
Table 11. Reaction for Golden Gate assembly
Reagent Volume
10× Cutsmart buffer 2 μL
10× T4 ligase buffer 2 μL
pU6X:sgRNA#X 100 ng each type
pGGDestISceIEG-XsgRNA 50 ng
T4 DNA ligase 1 μL
Bsa 1 μL
H2O To 20 μL
Place the mixture into the thermocycler and run the following program (Table 12):
Table 12. Program for Golden Gate assembly
Temperature Time Cycle number
37 °C 20 min 3 cycles
16 °C 15 min
80 °C 15 min 1 cycle
Take out 5 μL of mixture to do transformation. The solid LB medium plate is ampicillin-positive.
Perform colony PCR with Dest forward primer and Dest reverse primer.
Bacteria growth and extraction of plasmids: Subject plasmids to Sanger sequencing using Dest forward primer and Dest reverse primer. Then, align the sequence results with that of the predicted vector.
Assembly sgRNA expression cassettes for two genes (Figure 5B).
We have reported that the oocyte-specific conditional knockout strategy can be utilized to obtain double-gene maternal mutants. We just need to insert the sg RNA expression cassette of one gene into another through Gibson assembly. For convenience, we termed the pGGDestISceIEG-XsgRNA of two different genes as pISceI-XsgRNA-gene a and pISceI-XsgRNA-gene b.
Check the sequence of two plasmids to find whether there is an Asp718(Acc65) digesting site. If not, select anyone as the backbone. Here, we assume pISceI-XsgRNA-gene a as the backbone.
Amplify the sgRNA expression sequence of pISceI-XsgRNA-gene b.
i. Set up the PCR reaction as shown in Table 13.
Table 13. PCR reaction for amplification of sgRNA expression sequence
Reagent Volume
5× Phusion HF buffer 10 μL
dNTPs (10 mM) 1 μL
Ligation forward primer 2 μL
Ligation reverse primer 2 μL
pISceI-XsgRNA-2 (0.1 ng/μL) 1 μL
Phusion high-fidelity DNA polymerase 0.5 μL
H2O To 50 μL
ii. Run the following PCR program (Table 14):
Table 14. PCR program for amplification of sgRNA expression sequence
Temperature Time Cycle number
98 °C 30 s 1 cycle
98 °C 10 s 35 cycles
58 °C 15 s
72 °C 1 min
72 °C 5 min 1 cycle
iii. After gel visualization and purification with the AxyPrep PCR Clean-Up Kit, measure the concentration and store it at -20 °C.
Linearize pISceI-4sgRNA-1 with Asp718 following the instruction. Perform electrophoresis and gel visualization to confirm sufficient digestion. Then, purify the linearized pISceI-4sgRNA-1 and measure the concentration.
Perform Gibson assembly of purified PCR products and linearized backbone vector using Hieff Clone Plus One Step Cloning Kit.
Perform transformation, colony PCR, bacteria amplification, and plasmids extraction. Next, subject plasmids to Sanger sequencing to ensure the correct ligation. Store plasmids at -20 °C.
Transgenesis of sgRNA expression vector
Assemble the injection solution on ice as shown in Table 15.
Table 15. Injection mixture for I-SceI-mediated transgenesis
Reagent Volume
pGGDestISceIEG-XsgRNA (10 ng/μL) 7.5 μL
10× CutSmart buffer 0.5 μL
I-Sce 0.5 μL
Total volume 10 μL
Collect embryos from wild-type female mating with Tg(zpczcas9) homozygous male.
Inject the solution immediately after collecting embryos. We puncture into the blastodisc instead of the yolk and inject 2 nL of solution. To ensure efficient transgenesis, only 1-cell stage embryos are used.
After injection, incubate the rest of the injection solution at 37 °C for 30 min to test the I-Sce I activity. Then, subject all solutions to electrophoresis to check whether I-Sce works efficiently (Figure 6). If I-Sce I works well, it should completely cut the plasmids into two fragments at this condition. If we can still find the original band, it means the I-Sce I is not effective. This result can be considered as a quality control for I-Sce I-mediated transgenesis.
Figure 6. Representative electrophoresis result of I-Sce efficiency test. I-Sce I digestion can split the original plasmid into two fragments. No original plasmids should be found when the I-Sce1 works well.
At 24 hpf, pick up the embryos with strong and ubiquitous GFP signals. In general, 200 alive embryos will contain 10–20 efficiently transgenic ones. Raise the selected embryos to adulthood.
Pre-screening of founders
Due to the individual variation in the transgenic expression of Cas9, we need to perform pre-screening of the founders. To facilitate a rapid assessment, we inject bmp2b sgRNA into the embryos of the founder. This is because bmp2b is crucial for early embryo dorsal-ventral patterning, and its mutant phenotype becomes visible as early as 10 hpf. At 12 hpf, the dorsalized embryos should display a long elliptical shape, while the normal embryos are spherical (Figure 7).
Figure 7. Representative images of normal and dorsalized embryos at 12 hpf. Scale bar: 200 μm.
Synthesize bmp2b sgRNA as described above.
Mate Tg(zpc:zcas9;U6X:sgRNAX) female individuals with wild-type males.
Collect embryos and inject 2 nL bmp2b sgRNA (50 ng/μL) per embryo at 1-cell stage.
Observe the embryos at 12 hpf. Quantify the dorsalized embryos (long elliptical shape). If the ratio exceeds 50%, the fish is considered a putative founder.
Aspiration of blastoderm cells for genotyping
Zebrafish show regulative development. Hence, we can isolate some cells for examination of maternal mRNA and the rest of the embryo can still develop, so that we can achieve genotyping and phenotype analysis simultaneously. Maternal products originate from multiple duplicated alleles in the oocyte, including the one transmitted. Hence, mutated transmitted alleles could not represent the elimination of wild-type maternal products of the target gene. To identify the maternal mutants, we will examine the maternal mRNAs through RT-PCR.
Add 1.5% agarose in water and boil it to melt. Pour it into the 90 mm Petri dish and put the mold onto the molten agarose. Bend the solidified agarose and remove the mold gently, then the plate is ready to use.
Pull glass capillaries to make tip-closed needles using the puller. To prepare the needles suitable for cell transplantation, we use two light weights, set at Lv1: 60 and Lv2: 90, and select Step2 procedure. Using tweezers, we cut the capillary tips to achieve an opening diameter of 30–40 μm. It is crucial to ensure a clean and precise cut to facilitate effective cell aspiration. Then, make a spike at the tip of the pipette using a microforge. The spike facilitates easier penetration and reduces damage to embryos. The pipette for cell isolation is now ready to use.
Mate putative founder with wild-type fish and collect embryos. Pick up the GFP-positive embryos at 1-cell stage. At 3 hpf, put embryos onto the agarose plate (Figure 8). Reorient the embryos to make the blastomere point to the tip of the capillary. Use a microinjector to aspirate 20–40 cells from the embryo. Carefully control the pressure to avoid damaging the cells [18]. Then, release these cells into 2 μL of deionized H2O at the opening edge of different tubes. Clean the capillary through suction and ejection of deionized H2O three times before each aspiration to prevent cross-contamination and ensure accurate cell collection. Add 200 μL of TRIzol to wash down the cells. The tube can be put on ice temporarily. Put the embryos into 24-well plates separately. To prevent infection, change the medium to 1/3× Ringer’s solution containing 1× penicillin-streptomycin solution to allow the development of the embryos to proper stages for phenotyping. We normally examine 24 embryos at once, which will usually take approximately 1 h. Aspiration of dozens of cells will not be harmful to embryo development. Importantly, zygotic transcription does not commence extensively during this interval, allowing us to specifically examine maternal transcripts in these cells. The cells in TRIzol can be stored at -20 °C until the aberrant phenotype of the corresponding embryos is observed. Subsequently, the genotypes of cells from embryos with or without developmental defects are examined by RT-PCR. However, if no developmental defects are present, cells from all embryos should be genotyped to confirm the presence of maternal mutant embryos.
Figure 8. Illustration of procedures of phenotyping and genotyping individual embryos [11,12,18]
Phenotype analysis of the embryos (Figure 8)
When the embryos develop to the desired stage, maternal mutants may be directly recognized by live imaging. Otherwise, they can be separately fixed with 4% PFA at 4 overnight for in-depth analysis. Wash the samples with 1 mL of PBST three times (10 min for each time), and then subject them to whole-mount in situ hybridization, immunofluorescent staining, or other chemical staining assays. Follow established protocols for each assay to ensure accurate and reproducible results. Optionally, we can dehydrate the embryos by incubating three times in 1 mL of 100% methanol for 5 min each. The dehydrated embryos can be stored at -20 for an extended period. Before the next staining assay, we need to rehydrate the embryos with 1 mL of 75%, 50%, and 25% methanol in PBST for 5 min each, and rinse them with PBST three times to remove the methanol completely.
Genotyping of maternal mutants
For genotyping, first extract the total RNA of isolated cells of corresponding embryos separately as follows: Add 60 μL of chloroform and vortex for 5–10 s. Centrifuge at 12,000× g for 15 min at 4 and transfer supernatant to a new tube. Add an equivalent volume of isopropanol and 1 μL of glycogen (20 mg/mL, facilitating the precipitation of RNA), then mix it and incubate at -80 for 30 min to precipitate RNA. Centrifuge at 12,000× g for 15 min at 4 . Discard the supernatant and rinse the pellet with pre-chilled 70% ethanol. Centrifuge at 12,000× g for 5 min at 4 . Remove the supernatant and resolve the pellet with 8 μL of deionized H2O (the volume depends on the cDNA synthesis reaction so that the total RNA can be applied for cDNA synthesis). Then, reverse-transcribe the cDNA following the cDNA synthesis kit manual.
Use the cDNA as the template to amplify the coding sequence of the target gene. The PCR mixture is as shown in Table 16.
Table 16. PCR reaction for amplification of the coding sequence of the target gene
Reagent Volume
2× Phanta Max Mixture 25 μL
cDNA 1 μL
Forward primer (10 μm) 2 μL
Reverse primer (10 μm) 2 μL
deionized H2O 20 μL
Total volume 50 μL
The PCR procedure is as shown in Table 17.
Table 17. PCR program for amplification of the coding sequence of the target gene
Temperature Time Cycle number
95 °C 30 s 1 cycle
95 °C 15 s 35 cycles
58 °C 15 s
72 °C 30 s/kb
72 °C 5 min 1 cycle
Take 2 μL of PCR products to perform electrophoresis and purify the rest using a clean-up kit.
Ligate the purified PCR products to the pBackZero-T vector following the kit manual.
Mix the ligation mixture with 100 μL of DH5α competent cell. Incubate on ice for 30 min and heat shock for 60 s at 42 . Immediately after heat shock, chill the cells on ice for 2 min. Then, spread it on the ampicillin-resistant solid LB medium and incubate for 14 h at 37 .
Pick up 40–60 colonies randomly, stab them with autoclaved tips separately, and pipette into LB medium in each test tube. After growth at 37 for 14 h, extract plasmids using commercial kits and subject them to Sanger sequencing. Align the sequencing results with the wild-type target gene sequence. If there is no wild-type sequence, we consider the corresponding embryo as a maternal mutant.
Data analysis
Using this protocol, we have successfully obtained ctnnb2, nanog, and rbm24a maternal mutants [12]. Further, we also constructed dvl2 and dvl3a double-gene maternal mutants [11]. Here, we will show the data from these two articles as examples.
We designed three sgRNAs with high efficacy to target the coding sequence of ctnnb2. The maternal expression of ctnnb2 is vital for initiating maternal Wnt signaling and is indispensable for the development of the dorsal organizer [19,20]. Interference with maternal ctnnb2 expression leads to zebrafish embryos exhibiting a ventralized phenotype. Adhering to this protocol, we screened 10 female zebrafish and identified two that produced embryos positive for GFP. Analysis of these embryos at 1-day post fertilization (dpf) showed that, on average, 25.6% of GFP-positive embryos displayed various degrees of ventralization (Figure 9A and B). These phenotypes were classified into four categories (V1 to V4) according to established criteria [19]. Additionally, the ventralized phenotype was successfully rescued by injecting wild-type ctnnb2-myc mRNA at the one-cell stage (Figure 9B). We then randomly selected four embryos representing the V1–V4 phenotypes and collected cell samples from each to construct a plasmid library. Sequencing of 116 colonies confirmed that they all contained mutations in the ctnnb2 coding sequence (Figure 9C).
Figure 9. Generation of ctnnb2 maternal mutants using this protocol. These data were reproduced from our previous publication [12]. A. Phenotypes of the GFP-positive embryos expressing three sgRNAs targeting ctnnb2. B. Injection of wild-type ctnnb2-myc mRNA at the 1-cell stage could efficiently rescue the ventralized phenotypes. C. Schematics shows the mutation types by examining colonies.
In zebrafish, dvl2 and dvl3a play redundant roles in regulating Wnt/PCP and zygotic Wnt/beta-catenin signaling pathways [2,21]. Mutations in either dvl2 or dvl3a alone do not result in noticeable phenotypes. However, the simultaneous loss of maternal and zygotic expression of both dvl2 and dvl3a leads to anteriorization and impaired convergence and extension cell movements [2]. We selected four highly efficient sgRNAs each for targeting the coding sequences of dvl2 and dvl3a. After constructing the sgRNA expression vectors for dvl2 and dvl3a and performing transgenesis via I-Sce I, we obtained two F0 founder females capable of producing a high percentage of GFP-positive progeny. Among the GFP-positive embryos from these founders, 5.4% exhibited a shortened body axis and expanded dorsal structures at 12 h post-fertilization (hpf). By 30 hpf, these embryos showed pronounced yolk extension defects. Crossbreeding these two female founders with dvl2+/-;dvl3a+/- mutant males resulted in embryos with broad notochords and severely shortened body axes at 12 hpf. At 30 hpf, these embryos presented posterior truncation, closely resembling the phenotypes described for MZdvl2;MZdvl3a mutants (Figure 10A). To genotype the dvl2 and dvl3a maternal mutants, we amplified the coding sequences of both genes. In comparison to wild-type and other GFP-positive control embryos, one double-gene maternal mutant (#1) lacked the wild-type coding sequences for both dvl2 and dvl3a (Figure 10B). Sanger sequencing revealed substantial deletions in the coding regions of both dvl2 and dvl3a (Figure 10C).
Figure 10. Generation of dvl2 and dvl3a maternal mutants using this protocol. These data were reproduced from our previous publication [11]. A. Phenotypes of wild-type, Mdvl2;Mdvl3a and MZdvl2;MZdvl3a embryos at 12 hpf and 30 hpf. B. Gel analysis after amplifying the coding sequence of dvl2 and dvl3a. C. Large deletions happened both in dvl2 and dvl3a coding sequences.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Zhang et al. [11]. A Time-Saving Strategy to Generate Double Maternal Mutants by an Oocyte-Specific Conditional Knockout System in Zebrafish. Biology (Basel) (Figures 2–3).
Zhang et al. [12]. Rapid generation of maternal mutants via oocyte transgenic expression of CRISPR-Cas9 and sgRNAs in zebrafish. Science Advances (Figures 2–4).
General notes and troubleshooting
Problem Solution
No commercial Cas9 protein for sgRNA test. In vitro transcription and purification of Cas9 mRNA and use it to test sgRNA efficiency.
Low maternal mutant frequency. Prescreen Tg(zpc:zcas9) by injecting bmp2b sgRNA into the embryos and select the fish with injected embryos displaying a high ratio of dorsalized phenotypes.
Failure in amplification of the coding sequence. Use nested PCR.
Acknowledgments
We are grateful to Bo Zhang and Wenbiao Chen for providing zcas9 and pU6X:sgRNA#X plasmids. This work was supported by Guangdong Basic and Applied Basic Research Foundation [2023A1515110960], Central People’s Hospital of Zhanjiang Startup Project of Doctor Scientific Research [2020A12], and Zhanjiang Science and Technology Project [2022A01076 and 2022A01079]. This protocol was adapted from our previous work [11,12]. We thank all the authors who conducted the original work.
Competing interests
The authors declare no competing interests.
Ethical considerations
Zebrafish were raised under standard conditions. All experiments were designed and performed following the principles issued by the Ethics Committee for Animal Research of Life Science of Shandong University (permit number SYDWLL-2018-05).
References
White, R. J., Collins, J. E., Sealy, I. M., Wali, N., Dooley, C. M., Digby, Z., Stemple, D. L., Murphy, D. N., Billis, K., Hourlier, T., et al. (2017). A high-resolution mRNA expression time course of embryonic development in zebrafish. eLife. 6: e30860.
Xing, Y. Y., Cheng, X. N., Li, Y. L., Zhang, C., Saquet, A., Liu, Y. Y., Shao, M. and Shi, D. L. (2018). Mutational analysis of dishevelled genes in zebrafish reveals distinct functions in embryonic patterning and gastrulation cell movements. PLos Genet. 14(8): e1007551.
Hino, H., Nakanishi, A., Seki, R., Aoki, T., Yamaha, E., Kawahara, A., Shimizu, T. and Hibi, M. (2018). Roles of maternal wnt8a transcripts in axis formation in zebrafish. Dev Biol. 434(1): 96–107.
Gritsman, K., Zhang, J., Cheng, S., Heckscher, E., Talbot, W. S. and Schier, A. F. (1999). The EGF-CFC Protein One-Eyed Pinhead Is Essential for Nodal Signaling. Cell. 97(1): 121–132.
Shao, M., Lu, T., Zhang, C., Zhang, Y. Z., Kong, S. H. and Shi, D. L. (2020). Rbm24 controls poly(A) tail length and translation efficiency of crystallin mRNAs in the lens via cytoplasmic polyadenylation. Proc Natl Acad Sci USA. 117(13): 7245–7254.
Zhang, C., Huang, R., Ma, X., Chen, J., Han, X., Li, L., Luo, L., Ruan, H. and Huang, H. (2021). The Ribosome Biogenesis Factor Ltv1 Is Essential for Digestive Organ Development and Definitive Hematopoiesis in Zebrafish. Front Cell Dev Biol. 9: e704730.
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Ciruna, B., Weidinger, G., Knaut, H., Thisse, B., Thisse, C., Raz, E. and Schier, A. F. (2002). Production of maternal-zygotic mutant zebrafish by germ-line replacement. Proc Natl Acad Sci USA. 99(23): 14919–14924.
Wu, X., Shen, W., Zhang, B. and Meng, A. (2018). The genetic program of oocytes can be modified in vivo in the zebrafish ovary. J Mol Cell Biol. 10(6): 479–493.
Liu, Y., Zhu, Z., Ho, I. H. T., Shi, Y., Xie, Y., Li, J., Zhang, Y., Chan, M. T. V. and Cheng, C. H. K. (2017). Germline-specific dgcr8 knockout in zebrafish using a BACK approach. Cell Mol Life Sci. 74(13): 2503–2511.
Zhang, C., Li, J., Tarique, I., Zhang, Y., Lu, T., Wang, J., Chen, A., Wen, F., Zhang, Z., Zhang, Y., et al. (2021). A Time-Saving Strategy to Generate Double Maternal Mutants by an Oocyte-Specific Conditional Knockout System in Zebrafish. Biology. 10(8): 777.
Zhang, C., Lu, T., Zhang, Y., Li, J., Tarique, I., Wen, F., Chen, A., Wang, J., Zhang, Z., Zhang, Y., et al. (2021). Rapid generation of maternal mutants via oocyte transgenic expression of CRISPR-Cas9 and sgRNAs in zebrafish. Sci Adv. 7(32): eabg4243.
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Article Information
Publication history
Received: Jun 10, 2024
Accepted: Aug 28, 2024
Available online: Oct 13, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biological Sciences > Biological techniques > CRISPR/Cas9
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Generation and Maintenance of Patient-Derived Endometrial Cancer Organoids
MB Mali Barbi *
DG Divya Gowthaman *
AK Arielle Katcher
MG Megan Gorman
BY Brian Yueh
AN Aaron Nizam
CC Charlie Chung
EA Erdogan Oguzhan Akyildiz
MF Marina Frimer
GG Gary L. Goldberg
SB Semir Beyaz
(*contributed equally to this work)
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5093 Views: 438
Reviewed by: Alessandro DidonnaRakesh Bam Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Frontiers in Endocrinology Apr 2023
Abstract
Endometrial cancer (EC) is the leading cause of gynecologic cancer morbidity and mortality in the U.S. Despite advancements in cancer research, EC death rates are increasing, particularly high-grade endometrial cancers. The development of three-dimensional (3D) patient-derived organoid (PDO) models for EC is crucial, as they provide a more accurate representation of the biological and genetic complexity of a patient’s tumor compared to traditional 2D cell lines. Here, we describe a protocol for cultivating PDO models from normal endometrium and EC across different EC subtypes. These EC PDO models can be expanded across multiple passages and facilitate the exploration of tumor behavior and drug responses, thereby advancing our understanding of the disease and potentially leading to more effective and individualized novel therapeutic strategies.
Key features
• Establishment of patient-derived EC and normal endometrium organoids.
• Accurate replication of the various histologic and molecular subtypes of EC within the organoids.
• Our approach provides a clinically relevant platform for studying EC development, metastasis, progression, and recurrence.
• It offers potential for developing targeted therapeutic interventions tailored to specific EC subtypes.
Keywords: Endometrial cancer Patient-derived organoids Long-term expansion Tumor selectivity Translational research
Background
Endometrial cancer (EC) is the fourth most common cancer and the sixth leading cause of cancer mortality among women in the US. It is one of the few cancers experiencing a rapid rise in both incidence and mortality rates, and by 2040, it is projected to become the third most prevalent cancer and the fourth leading cause of cancer death, surpassing lung and colorectal cancers. Furthermore, EC presents a significant challenge due to its racial disparity, with the largest Black/White difference in 5-year relative survival among all cancers (63% vs. 84%) [1–4]. These concerning trends highlight the urgent need for research into the molecular and cellular basis of EC to advance our understanding and develop more effective therapeutic strategies.
Immortalized 2D cell lines and mouse models have been commonly used for EC research; however, they do not provide reliable models. 2D cell culture systems lack the critical 3D structure inherent in primary endometrial tissue, affecting glandular cell function influenced by their microenvironment and orientation. Similarly, mouse models do not accurately represent the biology of the human endometrium, highlighting the need for reliable biological models that mimic human EC biology [5,6].
Patient-derived organoids (PDOs) are 3D structures that precisely replicate the microenvironment and cellular composition of endometrial tissue, providing a clinically relevant model for studying specific histologic and molecular subtypes of EC [7–11]. This is particularly valuable for the rarer and more aggressive EC subtypes that disproportionately affect minority populations, especially African American women [2,12–14]. Additionally, the establishment of PDOs addresses the limitation of tissue samples by providing a sustainable cellular resource for ongoing research [15]. PDOs have been shown to maintain their phenotype and genotype through multiple passages, making them reliable models that preserve biological integrity over extended periods in culture [16,17]. However, there remains a need to enhance organoid culture techniques, as organoid cultures may exhibit heterogeneity and variable complexity in cellular composition, as well as poorly controlled morphogenesis during the self-assembly process [18].
Here, we present an optimized method for establishing, expanding, and banking endometrial organoids derived from patient tissue samples of normal endometrium and endometrial cancers.
Materials and reagents
Biological materials
Fresh tissue endometrial specimen in RPMI-1640 medium (Thermo Fisher Scientific, catalog number: 11879020)
Reagents
Advanced DMEM/F12 (Thermo Fisher Scientific, catalog number: 12634028)
Matrigel® matrix (Corning, catalog number: 356234)
RPMI-1640 medium (Sigma-Aldrich, catalog number: R73-88)
GlutaMAX (Thermo Fisher Scientific, catalog number: 35050061)
HEPES (Sigma-Aldrich, catalog number: S7067)
R-Spondin conditioned media (homemade) [19], filter through a 0.2 μm mesh prior to use
N2 supplement (Life Technologies, catalog number: 17502048)
B27 supplement minus vitamin A (Thermo Fisher Scientific, catalog number: 12587010)
Chemically defined lipid concentrate (Thermo Fisher Scientific, catalog number: 11905031)
Recombinant human Noggin (PeproTech, catalog number: 120-10c)
Primocin (InvivoGen, catalog number: ant-pm-1)
Nicotinamide (Sigma-Aldrich, catalog number: N0636)
N-Acetyl-L-cysteine (NAC) (Sigma-Aldrich, catalog number: A9165-5G)
Y-27632 dihydrochloride (Tocris, catalog number: 1254)
A83-01 (Sigma-Aldrich, catalog number: SML0788)
Recombinant human EGF (PeproTech, catalog number: AF-100-15)
Recombinant human FGF-α (PeproTech, catalog number: 100-26)
Recombinant human FGF-4 (PeproTech, catalog number: 100-31)
Recombinant human HGF (PeproTech, catalog number: 100-39)
Recombinant human IGF-1 (PeproTech, catalog number: 100-11)
Recombinant human FGF-10 (PeproTech, catalog number: 100-26)
Recombinant human FGF-β (PeproTech, catalog number: 100-18B)
SB202190 (p38i) (Sigma-Aldrich, catalog number: S7067)
17-β Estradiol, water soluble (Sigma-Aldrich, catalog number: E4389)
Collagenase from Clostridium histolyticum (Sigma-Aldrich, catalog number: C9407)
Cell recovery solution (Corning, catalog number: 354253)
RecoveryTM cell culture freezing medium (Thermo Fisher Scientific, catalog number: 12648010)
TrypLETM express enzyme (1×), no phenol red (Thermo Fisher Scientific, catalog number: 12604103)
Phosphate buffered saline (PBS) (Thermo Fisher Scientific, catalog number: 10010023)
ACK buffer lysis (Sigma-Aldrich, catalog number: A1049201)
Trizol reagent (Fisher Scientific, catalog number: 15-596-018)
0.4% trypan blue solution (Thermo Fisher, catalog number: 15250061)
Solutions
Human tumor endometrial organoid media (see Recipes)
Human normal endometrium organoid media (see Recipes)
Recipes
Human tumor endometrial organoid media (50 mL)
Reagent Final concentration Quantity or Volume
Advanced DMEM/F12 1× 30 mL
GlutaMAX 1% 500 μL
HEPES 12.5 mM 500 μL
R-Spondin conditioned media 10% 5 mL
B27 2% 1 mL
N2 1% 500 μL
Lipid concentrate 1% 500 μL
Noggin (NOG) 100 ng/mL 5 μL
Primocin 100 μg /mL 100 μL
Nicotinamide 2 mM 250 μL
NAC 1.25 mM 102 μL
Y-27632 dihydrochloride 10 μM 5 μL
A83-01 0.25 μM 0.25 μL
hEGF 50 ng/mL 2.5 μL
hFGF-α 25 ng/mL 2.5 μL
hFGF-4 50 ng/mL 5 μL
hHGF 20 ng/mL 4 μL
hIGF-1 40 ng/mL 2 μL
SB202190 0.1 μM 0.25 μL
17-β Estradiol 10 nM 5 μL
Total (optional) n/a 50 mL
Store at 4 °C for a maximum of 10 days.
Human normal endometrium organoid media
Reagent Final concentration Quantity or Volume
Advanced DMEM/F12 1× 30 mL
GlutaMAX 1% 500 μL
HEPES 12.5 mM 500 μL
R-Spondin conditioned media 15% 7.5 mL
B27 2% 1 mL
N2 1% 500 μL
Lipid concentrate 1% 500 μL
Noggin (NOG) 100 μg/mL 5 μL
Primocin 100 μg /mL 100 μL
Nicotinamide 2 mM 250 μL
NAC 1.25 mM 102 μL
Y-27632 dihydrochloride 10 μM 5 μL
A83-01 0.25 μM 0.25 μL
hEGF 50 ng/mL 2.5 μL
hFGF-10 10 ng/mL 1 μL
FGF-β 2 ng/mL 2 μL
SB202190 10 μM 25 μL
17-β Estradiol 1 nM 0.5 μL
Total (optional) n/a 50 mL
Store at 4 °C for a maximum of 10 days.
Laboratory supplies
Falcon 15 mL tube (Corning, catalog number: 352097)
Falcon 50 mL tube (Corning, catalog number: 352098)
Eppendorf 1.5 mL tube (Eppendorf, catalog number: 22364116)
Eppendorf 5.0 mL tubes (Eppendorf, catalog number: 0030119487)
CRYOVIAL® internal thread with silicone washer seal (Simport, catalog number: T311-1)
Mini-strainer 40 μm mesh (Corning, catalog number: CLS431750)
Mini-strainer 100 μm mesh (Corning, catalog number: CLS431752)
Falcon® 6-well cell culture plate (Corning, catalog number: 353046)
PYREX® 100 × 10 mm Petri dish (Corning, catalog number: 3160-100)
Pipette tips, 10, 20, 200, 1000 μL
Cell lifter (Corning, catalog number: 3008)
Forceps (Roboz Surgical Instrument, catalog number: RS-5136)
Scissors (Roboz Surgical Instrument, catalog number: RS-5910)
Equipment
Sterile tissue/cell culture hood (Labgard Class II TYPE A2)
Humidified CO2 incubator (Binder Incubator, model: CB150)
Water Bath (Sheldon Manufacturing Shel Lab H2O Bath Series)
Refrigerated centrifuge (Eppendorf, model: Centrifuge 5810R)
Light microscope (Nikon, model: Microscope Eclipse Ts2)
Portable Pipet-Aid (Thermo Scientific, model: S1 Pipet Fillers)
Cell drop automated cell counter (DeNovix, catalog number: CellDrop BF)
Fridge and freezers (4 °C, -20 °C, and -80 °C)
Software and datasets
NIS-Elements D Imaging Software
Images were acquired using the NIS-ELEMENT D_V5 software (Nikon) with a Nikon Eclipse TS2 microscope.
Procedure
Fresh tumor and normal endometrial tissue specimens were obtained from patients undergoing hysterectomy for EC. Institutional Review Board approval was obtained (study IRB #18-0897), and all patients provided informed consent prior to specimen collection. The hysterectomy specimens were transported from Northwell Health LIJ Medical Center to Cold Spring Harbor Laboratory shortly after surgery or the following morning. If tissue could not be delivered or digested the same day as the surgery, it was stored overnight at 4 °C.
Generation of organoids from endometrial tumor and normal tissue
Pipette 4.5 mL of RPMI media from the sample and transfer to a 15 mL Falcon tube.
Remove the rest of the media from the original specimen tube and wash the tissue once with 5 mL of cold 1× PBS.
Note: If the sample is bloody, add 1 mL of ACK buffer lysis to the plate and keep on ice for 10–15 min. Spin and then remove supernatant. Proceed to step A3.
Mince 5 g of tissue with shears in a Petri dish to obtain 1 mm fragments.
Transfer the tissue to a 5 mL centrifuge tube with 4.5 mL of RPMI, 500 μL of 1 mg/mL Collagenase IV, and 10 μM Y-27632 dihydrochloride.
Incubate for 1–1.5 h at 37 °C. Vigorously shake the Falcon tube approximately 10 times every 15–30 min.
Evaluate digestion for dissociated fragments under light microscopy intermittently.
Allow any remaining tissue pieces to settle to the bottom of the tube. Then, collect the supernatant and transfer to a new Falcon tube. Alternatively, filter through a 100 μm mesh. Spin down at 300× g for 5 min at 4 °C.
Remove the supernatant and resuspend the pellet in a mixed solution of 1 mL of TrypLE and 10 μM Y-27632 dihydrochloride.
Incubate for a maximum of 15 min at 37 °C. Every 5 min, pipette up and down before placing it back in the incubator at 37 °C, confirming that the larger pieces of tissue are being digested. Aim for small clumps of organoids, not single cells.
Dilute contents 1:1 with advanced DMEM to stop the reaction. Pipette up and down to mix and then spin down at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend in a separate centrifuge tube in a 70:30 Matrigel and media mixture (900 μL of Matrigel and 300 μL of media for a total of 12,00 μL of volume).
Plate densely (100 cells/μL) in a 6-well plate and allow 5–10 min for the dome to polymerize in the CO2 incubator. Plate four domes (50 µL each) in each well.
Add 3 mL of culture media to each well and place in a 30 °C incubator.
On passage 0, day 1:
Check back the next day to assess organoid growth and health. The human normal crypts should have formed oval-shaped organoids. Organoids from the tumor isolation can vary greatly in size but check to see that they are growing and not dark.
On passage 0, day 2:
Freeze some normal human organoids before they turn dark (refer to Section C). If possible, take two tubes of Trizol for RNA isolation as well. Also, take two tubes of Trizol for RNA isolation of the human tumor organoids. Split (see Section B) some of the normal human organoids to see how they respond to passaging. This can be a trial run with only a few wells or more if you want to try various techniques.
On passage 0, day 3:
Freeze some human tumor organoids. Test their response to passaging on this day. Check on the normal human organoids that were split the day before and split most or all of the remaining ones this day.
On passage 0, day 4:
Check back on the tumor organoids. Split most or all of the remaining tumor organoids, if needed. Check to see if less dense normal organoids need to be combined into one well. Do not forget to exchange media with new media (2.7 mL).
Continue to assess confluency, dissociation requirements, and consolidation as your P1 organoids grow. From this point, the normal and tumor organoids can differ greatly in their needs and must be considered at different time points due to their growth rate (Figure 1).
Figure 1. Patient-derived organoids from normal endometrium and cancerous tissues. A. Organoids from normal endometrium, FIGO1 (low grade), FIGO2 (intermediate grade), FIGO3 (high grade), serous cancer, and carcinosarcoma demonstrating that all histologic subtypes can be cultured and maintained through several passages. B. Serous cancer organoids demonstrating survival and maintenance over several passages in culture at passage 1 (P1), passage 7 (P7), and passage 14(P14). Scale bars: 100 μm (10×) and 200 μm (4×).
Passaging of organoids
Aspirate media from desired wells and scrape off the Matrigel domes using a cell lifter.
Tilt the plate and collect all the Matrigel and organoids into the corner of the well closest to you.
Use Corning’s cell recovery solution (CRS) to collect the dome (minimum 3× volume of Matrigel domes collected) and then transfer to a new Eppendorf tube. For example, for 200 μL of Matrigel, add 600 μL of minimum CRS (for 3 wells, 1.2 mL).
Keep the tube on ice for 1 h or until organoids visibly settle. Evaluate intermittently under light microscopy for dissociation into single cells.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend the pellet with TrypLE + 10 μM Y-27632 (≥ 500 μL of TrypLE for a 25 μL cell pellet). Lightly pipette up and down to a maximum of 10 times to break up the pellet.
Incubate at 37 °C for 10 min.
Mix the solution to break apart organoids and assess for level of dissociation under light microscopy.
Repeat steps B7 and B8 as needed until organoids are broken down to the desired level, checking every 5–10 min based on the rate of digestion. Neutralize the TrypLE with an equal volume of media.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend with 75%/25% Matrigel and culture medium. (For 3 wells, 450:150 μL ratio; for 6 wells, 900:300 μL ratio.)
Plate and wait 10 min for the dome to polymerize at 37 °C.
Add culture medium to each well (2.7 mL/well in a 6-well plate).
Freezing organoids
Aspirate media from desired wells and scrape off the Matrigel domes using a cell lifter.
Tilt the plate and collect all the Matrigel and organoids into the corner of the well closest to you.
Use Corning’s cell recovery solution (CRS) or matrix melting solution (MMS) to collect the dome (minimum 3× volume of Matrigel domes collected), then transfer to a new Eppendorf tube.
Keep the tube on ice for 1 h.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and suspend in 500 μL of freezing media.
Thawing frozen organoids
Remove cryovials from liquid nitrogen storage.
Warm in a 37 °C water bath for 1 min.
Add culture medium 1:1 (500 μL) and transfer to Eppendorf tube.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend with 75%/25% Matrigel and culture medium. (For 3 wells, 450:150; for 6 wells, 900:300.)
Plate and wait 10 min for the dome to polymerize at 37 °C.
Add culture medium to each well (2.7 mL/well in a 6-well plate).
RNA isolation (low yield 50 cells/μL)
Aspirate media from desired wells.
Wash wells gently with 5 mL of 1× PBS and then remove PBS.
Add 400 μL of Trizol reagent directly to the well.
Break up Matrigel using a cell lifter and pipette up and down to mix, doing so until the solution is clear (all organoids/cells have been lysed).
Transfer to a labeled Eppendorf tube.
Vortex the tube for ~30 s.
Immediately freeze in -80°C.
RNA isolation (normal yield 100 cells/μL)
Aspirate media from desired wells and scrape off the Matrigel domes using a cell lifter.
Tilt the plate and collect all the Matrigel and organoids into the corner of the well closest to you.
Use Corning’s cell recovery solution (CRS) or matrix melting solution (MMS) to collect the dome (minimum 3× volume of Matrigel domes collected) without mixing.
Transfer the CRS with organoids to an Eppendorf tube and pipette up and down 10 times gently.
Keep on ice for 45 min.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and resuspend the pellet with 400 μL of Trizol.
Pipette up and down until the solution is clear (all organoids/cells have been lysed).
Vortex the tube for 30 s.
Immediately freeze at -80 °C.
Quantification of organoids
Aspirate media from desired wells and scrape off the Matrigel domes using a cell lifter.
Tilt the plate and collect all the Matrigel and organoids into the corner of the well closest to you.
Use Corning’s cell recovery solution (CRS) or matrix melting solution (MMS) to collect the dome (minimum 3× volume of Matrigel domes collected), then transfer to a new Eppendorf tube.
Keep the tube on ice for 1 h.
Spin down at 300× g for 5 min at 4 °C.
Remove supernatant and suspend in 500 μL of culture media.
Add 1 μL of 0.4% trypan blue solution to 1 μL of suspension and vortex.
Place 1 μL of the mixture into a cell drop automated cell counter to determine quantification.
Validation of protocol
This protocol or parts of it has been used and validated in the following research articles:
Chung, et al. [20]. Abstract 2511: Autologous patient-derived organoid-immune cell co-culture platform for therapeutic discovery in high-grade endometrial cancer. Cancer Res.
Nizam, et al. [21]. Abstract LB236: Utilizing endometrial tumor organoids to model cancer immunomodulation. Cancer Res.
General notes and troubleshooting
General notes
Passaging should be done before organoids exhibit signs of stress or poor health. These signs are a noticeable accumulation of dead cells surrounding the organoid, loss of three-dimensional morphology, and darker color.
Collecting the dome can be done with cold 1× PBS or CRS. CRS may be helpful if you do not aim to dissociate with trypsin but just want to replate while getting rid of the Matrigel. It can also help if you notice that your organoids have not been breaking up as well as expected. To facilitate dissociation, organoids can be incubated on ice in CRS for ~20 min. When using 1× PBS or CRS, any volume from a few hundred microliters to 1 mL can be used at a time to wash the dome off.
Always orient Eppendorf tubes in the centrifuge correctly to help you predict where the pellet will form.
Organoids/cells should be pelleted properly before the supernatant is removed.
Organoid dissociation involves mechanical shearing, enzymatic, or non-enzymatic digestion. Mechanical shearing can be either with pipetting or mixing with a vortex. Enzymatic digestion utilizes trypsin or Trypl Express (TrypLE) to cleave proteins that hold the cells together. Non-enzymatic digestion via EDTA or gentle cell dissociation reagent helps break the organoids apart via calcium chelation. Normally, we will dissociate with trypsin for a few min to “loosen” up the organoid, before pipetting up and down to completely break it up. In some situations, organoids can only be mechanically sheared and then replated. This should also be considered if the organoids are very unhealthy and may not survive trypsinization. Mechanical shearing on its own also makes it extremely hard to get small organoids or single cells and is a benefit when you want to have grown organoids in only a few days. For dissociation, we use TrypLE (gentler) or Pan Trypsin (harsher). Cells will begin to lyse if left in trypsin or TrypLE for too long. Ideally, the duration of trypsinization is long enough that organoids are loose and will easily separate with a bit of pipetting. Another option is to use gentle cell dissociation reagent (GCDR) for non-enzymatic dissociation.
Troubleshooting
Collapsing domes: It is beneficial to first prewarm the plates while you are passaging, with an emphasis on 24 and 48 wells. The dome may easily collapse when plating on a room-temperature plate. Although most protocols state that a 70/30 Matrigel media ratio is best, this can vary based on your goals. For maintenance, you can go as low as 50/50, especially if the wells are large (6 wells). On the other hand, if plating in a 48-well plate, you could do 80/20 to help make sure the dome does not collapse. Pure Matrigel or a pre-made mixture can be added if working with exceedingly small amounts of cells.
Carry-over of unhealthy organoids: Healthy organoids will be bright and clear, but it is important to note when some organoids are dying but others are healthy. This can happen with carry-over from a previous passage and should be taken into consideration when deciding the next time to split. This can also occur when organoids compete for resources, i.e., they are plated densely or grow too large and take up too much room.
Acknowledgments
We thank Cold Spring Harbor Laboratory (CSHL) Cancer Center Organoid Shared Resource supported by NCI Cancer Center Support grant 5P30CA045508, Northwell Health Biospecimen Repository and Northwell Health Department of Obstetrics & Gynecology research staff for their assistance in recruiting and consenting patients and obtaining specimens for this protocol. S. B. acknowledges funding from The Mark Foundation for Cancer Research, Chan-Zuckerberg Initiative, The Oliver S. and Jennie R. Donaldson Charitable Trust and The CSHL and Northwell Health Affiliation.
Competing interests
No conflicts of interest or financial disclosures.
Ethical considerations
Informed consent was obtained from all human patients and protocol #18-0897 was approved by the Northwell IRB committee.
References
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Desmond, D., Arter, Z., Berenberg, J. L., Killeen, J. L., Bunch, K. and Merritt, M. A. (2023). Racial and ethnic differences in tumor characteristics among endometrial cancer patients in an equal-access healthcare population. Cancer Causes Control. 34(11): 1017–1025. https://doi.org/10.1007/s10552-023-01716-9
Park, A. B., Darcy, K. M., Tian, C., Casablanca, Y., Schinkel, J. K., Enewold, L., McGlynn, K. A., Shriver, C. D. and Zhu, K. (2021). Racial disparities in survival among women with endometrial cancer in an equal access system. Gynecol Oncol. 163(1): 125–129. https://doi.org/10.1016/j.ygyno.2021.07.022
Gellersen, B., Brosens, I. and Brosens, J. (2007). Decidualization of the Human Endometrium: Mechanisms, Functions, and Clinical Perspectives. Semin Reprod Med. 25(6): 445–453. https://doi.org/10.1055/s-2007-991042
Liu, T., Shi, F., Ying, Y., Chen, Q., Tang, Z. and Lin, H. (2020). Mouse model of menstruation: An indispensable tool to investigate the mechanisms of menstruation and gynaecological diseases (Review). Mol Med Rep. 22(6): 4463–4474. https://doi.org/10.3892/mmr.2020.11567
Lancaster, M. A. and Knoblich, J. A. (2014). Organogenesis in a dish: Modeling development and disease using organoid technologies. Science (1979). 345(6194): e1247125. https://doi.org/10.1126/science.1247125
Clevers H. (2016). Modeling Development and Disease with Organoids. Cell. 165(7):1586–1597. https://doi.org/10.1016/j.cell.2016.05.082
Kim, J., Koo, B. K. and Knoblich, J. A. (2020). Human organoids: model systems for human biology and medicine. Nat Rev Mol Cell Biol. 21(10): 571–584. https://doi.org/10.1038/s41580-020-0259-3
Boretto, M., Maenhoudt, N., Luo, X., Hennes, A., Boeckx, B., Bui, B., Heremans, R., Perneel, L., Kobayashi, H., Van Zundert, I., et al. (2019). Patient-derived organoids from endometrial disease capture clinical heterogeneity and are amenable to drug screening. Nat Cell Biol. 21(8): 1041–1051. https://doi.org/10.1038/s41556-019-0360-z
Hibaoui, Y. and Feki, A. (2020). Organoid Models of Human Endometrial Development and Disease. Front Cell Dev Biol. 8: e00084. https://doi.org/10.3389/fcell.2020.00084
Amant, F., Moerman, P., Neven, P., Timmerman, D., Van Limbergen, E, and Vergote, I. (2005) Endometrial cancer. Lancet. 366(9484): 491–505. https://doi.org/10.1016/s0140-6736(05)67063-8
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Abel, M. K., Liao, C. I., Chan, C., Lee, D., Rohatgi, A., Darcy, K. M., Tian, C., Mann, A. K., Maxwell, G. L., Kapp, D. S., et al. (2021). Racial disparities in high-risk uterine cancer histologic subtypes: A United States Cancer Statistics study. Gynecol Oncol. 161(2): 470–476. https://doi.org/10.1016/j.ygyno.2021.02.037
Fitzgerald, H. C., Dhakal, P., Behura, S. K., Schust, D. J. and Spencer, T. E. (2019). Self-renewing endometrial epithelial organoids of the human uterus. Proc Natl Acad Sci USA. 116(46): 23132–23142. https://doi.org/10.1073/pnas.1915389116
Turco, M. Y., Gardner, L., Hughes, J., Cindrova-Davies, T., Gomez, M. J., Farrell, L., Hollinshead, M., Marsh, S. G. E., Brosens, J. J., Critchley, H. O., et al. (2017). Long-term, hormone-responsive organoid cultures of human endometrium in a chemically defined medium. Nat Cell Biol. 19(5): 568–577. https://doi.org/10.1038/ncb3516
Katcher, A., Yueh, B., Ozler, K., Nizam, A., Kredentser, A., Chung, C., Frimer, M., Goldberg, G. L. and Beyaz, S. (2023). Establishing patient-derived organoids from human endometrial cancer and normal endometrium. Front Endocrinol (Lausanne). 14: e1059228. https://doi.org/10.3389/fendo.2023.1059228
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Nizam, A., Chung, C., Goldberg, G. L. and Beyaz, S. (2021). Abstract LB236: Utilizing endometrial tumor organoids to model cancer immunomodulation. Cancer Res. 81: LB236–LB236. https://doi.org/10.1158/1538-7445.am2021-lb236
Article Information
Publication history
Received: Jul 1, 2024
Accepted: Aug 25, 2024
Available online: Sep 29, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Host Receptor Pili for Cryo-EM Single-Particle Reconstruction
RM Ran Meng
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5094 Views: 238
Reviewed by: Alba BlesaCuncai GuoSrajan Kapoor
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Original Research Article:
The authors used this protocol in Nature Communications Mar 2024
Abstract
Single-stranded RNA bacteriophages (ssRNA phages) infect their hosts by binding to the host receptor pili. Purification of pili usually involves mechanical shearing of pili from cells followed by precipitation. However, previous methods often result in low efficiency or unstable results due to pili retraction. This protocol presents an optimized method for purifying receptor type IV pili from Acinetobacter genomospecies 16 (A. gp16), incorporating enhancements in shearing and collection steps to achieve high yields. We found that repeated passage through syringe needles increases yield, and temperature control is crucial during purification. Additionally, the CsCl density gradient was optimized specifically for this specific strain. The purified type IV pili are suitable for cryogenic electron microscopy (cryo-EM) and various biochemical experiments.
Key features
• Pili purification for single-particle cryo-electron microscopy (Cryo-EM) analysis
• This protocol builds upon the F-pili purification method developed by Costa et al. [1] extending its application to the Acinetobacter genomosp. 16.
• It is optimized for higher and more stable pili yields, as well as increased reproducibility.
• The method is tested on various bacterial species and can be adapted to purify different types of pili.
Keywords: Host receptor Pili purification Single particle cryo-EM CsCl step gradient
Graphical overview
Workflow overview
Background
Acinetobacter, a Gram-negative bacterium, is an aerobic, non-flagellated, and widely distributed coccobacillus [2]. Acinetobacter baumannii, Acinetobacter calcoaceticus, Acinetobacter pittii, and Acinetobacter nosocomialis are opportunistic pathogens known for causing hospital-acquired infections. Bacteria can invade tissue and evade immune response through various mechanisms, including the production of enzymes. Adherence to the host is facilitated by structures such as pili [3–5]. Each year, U.S. hospitals report around 12,000 infections caused by multidrug-resistant Acinetobacter species. The protein secretion processes in Acinetobacter, which are critical targets for vaccine development, are not well understood. Known protein secretion systems in Acinetobacter include the Type II secretion system (T2SS), Type VI secretion system (T6SS), Type IV pilus secretion system, autotransporters, and outer membrane vesicles (OMVs) [6].
In this protocol, we focus on the type IV pili purification from Acinetobacter genomosp. 16 (A. gp16, NCBI: txid70347), which is a target host receptor of single-stranded RNA phage AP205 (ssRNA phage AP205, a virus that can infect Acinetobacter) [7]. Given the crucial role retractile pili play in the virulence of many bacteria, if infection-dependent pili detachment is a common feature of ssRNA phages, these simple viruses could potentially be used to inhibit retractile pili deployment and thereby decrease the virulence of numerous significant bacterial infections [7]. The purified pili can not only be utilized for Cryo-EM structural studies but also for biochemical or biophysical experiments to test protein binding affinity.
Type IV pili are extracellular helical appendages, typically with a diameter of 6–9 nm [8], which are composed of thousands of noncovalently linked pilin subunits. The major pilin of Acinetobacter type IV pili is the PilA monomer, which is composed of an N-termini hydrophobic α-helix, an α-β loop for connection, and the C-termini soluble β-sheet. The mature pilins assemble into type IV pilus by type IV pilus secretion system. Pilins bury the N-terminus helix inside, forming the pilus’s central axis, with the C-terminus headgroup (β-sheet) exposed outside [9].
Several protocols have been established for the purification of different types of pili [1,10,11]. While these methods share commonalities, specific steps can vary depending on the bacterial strain. We tested various pili purification techniques and found that the method developed by Costa et al. [1] for Escherichia coli (E. coli) F-pili provided higher efficiency and yield than other methods. First, we optimized and highlighted critical steps within this protocol to improve and stabilize yield, such as using a syringe to facilitate pili detachment from cells and controlling temperature to prevent pili retraction. Second, we focused on optimizing the procedure for Acinetobacter pili purification, allowing for the collection and separation of two different types of pili simultaneously. There are two types of pili outside the cell boundary: type IV pili and FilA filament. Lastly, this paper outlines a four-section process and provides two options (one for hyper-piliated strains, the other for low-yield strains), making it suitable for pili from other bacteria species as well (Graphical overview). We found that some steps could be simplified when bacterial pili yield is high. For instance, Acinetobacter pili have a relatively higher yield compared to F-pili, which have only one or two pili per cell. Microscope negative staining can be used to visually assess the number of pili per cell, determining whether the strain is hyper-piliated or has a lower yield. Based on this assessment, one can decide whether to follow the method in Sections B and C or Sections B2 and C2. Sections B and C provide a quicker method for obtaining pili from Acinetobacter and other high-yield species. Additionally, alternative steps in Sections B2 and C2, which mainly optimize the method from Costa et al. [1] are provided for strains with lower pili yields (e.g., E. coli F-pili).
In summary, this protocol has been optimized based on previous pili purification methods to enhance yield and stability, particularly for Acinetobacter pili. This method can also be generalized for the purification of pili in other species.
Materials and reagents
Biological materials
Acinetobacter genomosp. 16 (A. gp16, NCBI: txid70347)
Reagents
Difco LB broth, Miller (Miller Broth) 500 g (BD Bioscience, catalog number: 244620)
Tris (Bio-Rad, catalog number: 1610719)
Sodium chloride (NaCl) (crystalline/certified ACS) (Fisher Scientific, catalog number: 7647-14-5)
Sodium citrate dihydrate (granular/certified) (Fisher Scientific, catalog number: 6132-04-3)
(Optional) Polyethylene glycol 6000 (PEG6000) (Thermo Fisher Scientific, catalog number: 25322-68-3)
Cesium chloride (Thermo Fisher Scientific, catalog number: 7647-17-8)
Solutions
SSC buffer (see Recipes)
Dialysis buffer (see Recipes)
5% PEG6000 (see Recipes)
CsCl step gradient buffer (see Recipes)
Recipes
Note: All in sterile water.
SSC buffer
Reagent (room temperature) Final concentration (pH = 7.2)
Sodium citrate 15 mM
NaCl 150 mM
The buffer requires autoclaving.
Dialysis buffer
Reagent (room temperature) Final concentration (pH = 8.0)
Tris-HCl 50 mM
NaCl 150 mM
The buffer requires autoclaving.
5% PEG6000
Dissolve PEG6000 into dialysis buffer at pH = 8.0. The final volume should be 1 L of dialysis buffer with 50 g of PEG6000. The buffer requires autoclaving.
CsCl step gradient buffer
Reagent (room temperature) Final concentration (pH = 8.0)
CsCl
The buffer needs different densities to help form the gradient in the centrifuge tube.
Preparing density at 1.0 g/cm3, 1.1 g/cm3, 1.2 g/cm3, 1.3g/cm3, 1.4 g/cm3
Dialysis buffer
Laboratory supplies
Slide-A-LyzerTM dialysis cassettes, 20 K MWCO (Thermo Fisher, catalog number: 66003)
Sterile syringes for single use, 5 mL and 3 mL (Fisher Scientific, catalog numbers: 14-955-457, 14-955-458)
Disposable hypodermic needles 25 G, 23 G, and 18 G (Fisher Scientific, catalog numbers: 26406, 26408, 26420)
BD General Use and PrecisionGlide hypodermic needles (Fisher Scientific, catalog number: 305127)
FisherbrandTM Petri dishes with clear lids (Fisher Scientific, catalog number: 0875713)
FalconTM round-bottom polypropylene test tubes with cap (Fisher Scientific, catalog number: 14-959-11B)
PierceTM Protein Concentrators PES, 30 K MWCO (different sizes) (Thermo Fisher Scientific, catalog number: 88502, 88522, 88531)
Equipment
SW 41 Ti swinging-bucket rotor (Beckman, catalog number: 331362)
13.2 mL, open-top thin-wall ultra-clear tube, 14 × 89 mm, 50 Pk (Beckman, catalog number: 344059)
Procedure
Host bacteria culture
Day 1 afternoon
Recover A. gp16 (NCBI: txid70347) from -80 °C storage and place on ice.
Using a sterile inoculation loop, obtain a small amount of bacteria from the thawed stock.
Streak the loop across the surface of an LB agar plate using the quadrant streak method to gradually dilute the bacterial concentration.
Incubate the plate at 30 °C overnight to allow colony formation. After incubation, select well-isolated single colonies for further analysis or experimentation (Figure 1A).
Figure 1. Host bacteria culture, collection, and pili shearing. (A) Pipeline for host bacteria inoculation and culture. (B) Pipeline for pili collection. L-shaped spreaders are used to gently scrape cells from the surface of the plate. The collected bacterial cell suspension is then passed through a syringe to further remove the pili, followed by vortexing.
Day 2 afternoon
Prepare 100 LB agar plates without antibiotics (for use on Day 3).
Pick a single colony and inoculate it into 3 mL of LB medium for overnight shaking at 30 °C.
Day 3
In the morning (7 AM), take 100 µL from the 3 mL overnight culture and inoculate into 30 mL of LB medium. Incubate this A. gp16 cell culture with shaking at 200 rpm at 30 °C until the culture reaches OD600 = 0.1.
Note: For different strains, the target OD600 may vary. For example, for E. coli, OD600 = 0.6 is typical.
Add 100 µL of culture to each of the 100 prepared agar plates and spread evenly using a cell spreader.
Option 1: 100 plates. Most are for large-scale purification.
Option 2: 30 plates. For A. gp16 type IV pili, a smaller scale can also yield satisfactory results.
Option 3: Six large (25 cm × 25 cm) LB medium plates. This requires increasing the inoculum to 1 mL per plate. These larger plates reduce preparation time and simplify the procedure.
Incubate the plates at 30 °C for 30 min to allow the surface to dry slightly.
For overnight incubation, the plates should be inverted (bottom up, lid down) to prevent accumulated humidity from dripping onto the surface, which could affect pili formation. (Overnight incubation typically takes about 16 h. Deviating from this time frame may impact pili yield.)
(Option 1)
Host bacteria collection
Day 4
Pre-chill the dialysis buffer (1 L) at 4 °C.
Take out the overnight plates from 30 °C and immediately place them on ice. Process five plates at a time to prevent pili retraction at room temperature.
Collect the bacterial cells from the plates using 300 µL of dialysis buffer with an L-shaped spreader, then collect the liquid from the plate surface using a 1 mL pipette into a 50 mL Falcon tube.
Note: For 25 cm × 25 cm plates, use approximately 8 mL of dialysis buffer (Figure 1B).
Pili shearing and precipitation
Day 4
Pass the bacterial suspension through needles three times to shear the pili. Begin with the 18 G needle and then pass through the 25 G needle twice.
Note: Thinner needles help shear the pili but may clog.
Repeat the shearing process iteratively for all plates, processing 5–10 plates at a time. After shearing, vortex the liquid for 30 s (Figure 1B).
Centrifuge the cell solution at 3,470× g for 30 min at 4 °C to remove the cells.
Note: If the solution remains unclear, repeat the centrifugation. Higher speeds (4,000–8,000× g) can also be used as long as the cells are adequately spun down.
Collect the pili from the supernatant using a pipette. If starting with 30 plates, concentrate the pili solution to 2–3 mL using centrifugation. For larger volume purification (100 plates), concentrate the pili solution to 8–10 mL.
(Option 2)
For high-yield pili, such as A. gp16, this method is unnecessary. Sections B2 and C2 below can be used as an alternative method for pili purification of other low-yield species. It involves more steps and is relatively complicated. Refer to the general notes section for detailed explanations.
B2. Host bacteria collection (generalized method for all other kinds of pili purification)
Day 4
Pre-chill the SSC buffer at 4 °C.
Place the overnight plates on ice, processing five plates at a time to prevent pili retraction at room temperature.
Collect the bacterial cells from the plates using 500 µL of SSC buffer and an L-shaped spreader. Then, collect the liquid from the plate surface using a 1 mL pipette. For 25 cm × 25 cm plates, typically use 8 mL of SSC buffer. Slightly more buffer can be used in this step.
C2. Pili shearing and precipitation (generalized method for all other kinds of pili purification)
Day 4
Pass the bacterial suspension through needles three times to shear the pili. Begin with the 18 G needle first and then pass through the 25 G needle twice.
Note: Thinner needles help shear the pili but may clog.
Repeat the shearing process iteratively for all plates, processing 5–10 plates at a time, and vortex the liquid for 30 s.
After collecting all the plates, adjust the total volume of the solution to 1 L by adding pre-chilled SSC buffer. Incubate the liquid suspension at 4 °C for 2 h and stir gently.
Centrifuge the cell solution at 8,000× g for 30 min at 4 °C to remove any remaining cells. Centrifuge twice to make sure cells are removed, and then collect the supernatant using a pipette.
As the pili are present in large volumes, precipitate the supernatant by adding 5% PEG6000 (diluted from 50% PEG) and 500 mM NaCl. After 4 h of incubation at 4 °C, collect the precipitate by centrifuging the suspension at 15,000× g for 40–50 min, twice.
At this stage, resuspend the purified pili pellet in 2 mL of 50 mM Tris-HCl, 200 mM NaCl, pH 8.0 (F pilus).
Dialysis the 2 mL pili samples using dialysis buffer overnight.
Pili purification and dialysis
Days 5 and 6
CsCl step gradient setup: The purification steps need CsCl step gradients of 1.1, 1.2, and 1.3 g/cm3 (4 mL, 3 mL, and 3 mL, respectively). If the band is formed high in the tube, consider adding 1.4 g/cm3 for 0.5 mL and reducing 1.3 g/cm3 to 2.5 mL.
When loading the CsCl gradient, ensure the needle reaches the bottom of the centrifuge tube. Different density layers are loaded onto the tube one by one. Begin by loading the low-density gradient, followed by the middle-density gradient, and finally the highest-density gradient. The needle should always reach the bottom of the centrifuge tube. Gently depress the syringe plunger to allow the lighter gradients to rise smoothly. Finally, load around 1.5 mL of the pili sample onto the top of the gradient solution (Figure 2A).
Figure 2. CsCl gradient preparation, sample loading, and sample collection. (A) CsCl gradient preparation. When loading the CsCl gradient, ensure the needle reaches the bottom of the centrifuge tube. Begin by loading the low-density gradient, followed by the middle-density gradient, and finally the highest-density gradient. The needle should always reach the bottom of the centrifuge tube. Gently depress the syringe plunger to allow the lighter gradients to rise smoothly. The pili sample is loaded onto the top of the gradient solution in the final step. (B) When extracting the bands, remove the bottom band first to avoid disturbing the gradients above it.
Centrifuge the sample at 19,200× g for 19 h at 4 °C using SW 41 Ti swinging-bucket rotor. Ensure proper tube balance before starting centrifugation.
Note: For A. gp16, there will be two bands formed within the tube—the top band represents curved, thin pili, while the bottom band is composed of straight, thicker pili, which is the type IV pili. This is verified by the ssRNA phage as an antibody as well as mass spec. For F-pili, only one band forms in the tube, as strain MC4100 cells produce only one type of pili.
Use a syringe with a 23 G needle to extract the pili band from the tube.
Note: We prefer the 23 G needle because it is thin enough to precisely extract a single band while being sufficiently rigid to puncture the side of the centrifuge tube (Figure 2B).
Carefully remove the pili band and extensively dialyze it against the dialysis buffer using a dialysis cassette.
Perform initial dialysis for 4 h in 1 L of buffer, followed by overnight dialysis in 2 L of fresh buffer, and finally another 4 h of dialysis in 2 L of buffer.
Confirm the presence of pili using SDS-PAGE and verify their identity using LC-ESI MS/MS. Two types of pili are present in A. gp16. The presence and purity of type IV pili are confirmed by negative-stain electron microscopy.
Note: We used 4%–20% acrylamide gradient gels.
Validation of protocol
The purification method described in this protocol has been validated in two cryo-EM structure papers published in Nature Communications [7,12]. In Figure 3A, we present bands formed during two different pili purification batches. The “Pili Only” tube (right) illustrates typical results after type IV pili purification, where Layer 1 contains curved, thin FilA pili and Layer 2 contains straight, thick type IV pili. Cryo-EM analysis (Figure 3C and 3D) confirmed the purity of these layers. In A. gp16 cells, there are more curved, thin FilA pili than the straight, thick type IV pili outside cell boundaries. This is why in the final image, curved, thin pili showed relatively higher concentration.
In Figure 3A, the “Pili-Phage” tube (left) demonstrates an experiment where phage was premixed with the pili solution before the CsCl step. The phage with RNA of higher density appears in Layer 3 as a reference. The image shown in Figure 3B shows the phage and host receptor adsorption of purified pili from Layer 1 and Layer 2. (Purified AP205 phage was added into a mixture of pili from layer 1 and layer 2.) The red arrow points to straight, thick type IV pili with phage, while the blue arrow indicates curved, thin pili, which do not act as host receptors and show no phage adsorption. The scale bar represents a length of 500 A.
For mass spec and cryo-EM results for layers 1 and 2 please see the published paper’s supplementary section [7].
Figure 3. Validation of purified samples using EM. (A) Sample bands formed during two different purifications. The “Pili Only” tube (right) shows results after type IV pili purification, with the top band (Layer 1) containing curved, thin FilA pili, and the bottom band (Layer 2) containing straight, thicker type IV pili. The “Pili-Phage” tube (left) demonstrates an experiment where phage was premixed with the pili solution. The phage, containing RNA of higher density, appears in Layer 3 as a reference. (B) Cryo-EM image of purified pili from Layer 1 and Layer 2, along with additional purified AP205 phage. The red arrow points to straight, thick type IV pili bound with phage, while the blue arrow indicates curved, thin FilA pili, which are not host receptors and thus show no phage adsorption. The scale bar represents a length of 500 A. (C) Cryo-EM image of Layer 1 curved, thin pili. (D) Cryo-EM image of Layer 2 straight, thick pili.
General notes and troubleshooting
General notes
In Section D, the concentrated pili solution (approximately 1.5 mL) is layered on top of a CsCl gradient and centrifuged to separate into bands. For A. gp16, which yields relatively high amounts of pili, material collected from 15 plates and concentrated to 1.5 mL typically forms a visible band after CsCl gradient centrifugation. Processing pili from 100 plates yields about 9 mL of highly concentrated solution distributed into six CsCl tubes.
For low-yield strains, such as E. coli F-pili, the method described in Sections B2 and C2 should be used. This method is more complex but more broadly applicable to various pili purifications. It involves adding an SSC buffer, which aids in pili detachment. PEG6000 is used to precipitate and pellet the pili, which are then resuspended in 2 mL of dialysis buffer, resulting in a higher concentration per milliliter (containing pili from 100 plates in 2 mL). Refer to Sections B2 and C2 for details. Please note the bacterial culture temperature in Section A should be optimized if a different strain is used.
When preparing the CsCl gradient, start with 1.0, 1.2, and 1.4 g/cm3 gradients. Mix 1.0 and 1.2 to obtain a 1.1 g/cm3. Mix 1.2 and 1.4 to obtain a 1.3 g/cm3. It is better to check the density of the gradient by measuring the weight and dividing by the volume.
Different bacterial species may have multiple types of pili on the cell surface, so species other than A. gp16 might produce a different number of bands. For example, there is only one band formed for E. coli F-pili. Each layer of the band can be checked by negative stain electron microscopy (EM) to confirm its content.
We generally use 1.0, 1.1, 1.2, and 1.3 g/cm3 CsCl gradients for pili purification. If the pili band appears high in the tube, 0.5 mL of 1.4 g/cm3 CsCl can be added to slightly increase the overall density.
The CsCl gradient centrifugation should be balanced properly before loading on the high-speed centrifuge. The volume of solution in the centrifuge tube needs to be lower than the 13.2 maximum line but higher than the minimum.
Troubleshooting
If no band forms at the end, it indicates either that the strain has a low yield, or a critical step was not correctly performed.
To increase reproducibility and avoid sample loss, consider the following tips:
Pre-check the strain: Use negative staining under a microscope to assess the number of pili outside the cell boundary.
Bacteria culture: When culturing bacteria on LB plates, invert the Petri dish to prevent moisture accumulation and dripping, which can limit pili yield.
Two critical steps: Ensure precise control of temperature to avoid pili retraction. Ensure the proper use of the syringe at least three times to remove pili from cells.
Centrifugation tips: When using a centrifuge and condenser to concentrate samples, resuspend the concentrated solution with a pipette between spins to prevent pili from sticking to the concentrator membrane.
Increase sample concentration: If the band is not visible, increase the concentration of the samples loaded into the CsCl gradient centrifugation tube. Typically, 1–1.5 mL of sample is loaded, which can contain pili from 15–20 plates or more. For F-pili, pili from 50 plates may be necessary to form a visible band.
Acknowledgments
This protocol is based on a published research paper: Meng, R., Xing, Z., Chang, J. Y., Yu, Z., Thongchol, J., Xiao, W., Wang, Y., Chamakura, K., Zeng, Z., Wang, F., et al. (2024). Structural basis of Acinetobacter type IV pili targeting by an RNA virus. Nat Commun. 15(1): 2746 [7].
Competing interests
There is no competing interest.
References
Costa, T. R., Ilangovan, A., Ukleja, M., Redzej, A., Santini, J. M., Smith, T. K., Egelman, E. H. and Waksman, G. (2016). Structure of the Bacterial Sex F Pilus Reveals an Assembly of a Stoichiometric Protein-Phospholipid Complex. Cell. 166(6): 1436–1444.e10.
Ronish, L. A., Lillehoj, E., Fields, J. K., Sundberg, E. J. and Piepenbrink, K. H. (2019). The structure of PilA from Acinetobacter baumannii AB5075 suggests a mechanism for functional specialization in Acinetobacter type IV pili. J Biol Chem. 294(1): 218–230.
Harding, C. M., Kinsella, R. L., Palmer, L. D., Skaar, E. P. and Feldman, M. F. (2016). Medically Relevant Acinetobacter Species Require a Type II Secretion System and Specific Membrane-Associated Chaperones for the Export of Multiple Substrates and Full Virulence. PLoS Pathog. 12(1): e1005391.
Jones, A., Morgan, D., Walsh, A., Turton, J., Livermore, D., Pitt, T., Green, A., Gill, M. and Mortiboy, D. (2006). Importation of multidrug-resistant Acinetobacter spp infections with casualties from Iraq. Lancet Infect Dis. 6(6): 317–318.
Dijkshoorn, L., Nemec, A. and Seifert, H. (2007). An increasing threat in hospitals: multidrug-resistant Acinetobacter baumannii. Nat Rev Microbiol. 5(12): 939–951.
Costa, T. R. D., Felisberto-Rodrigues, C., Meir, A., Prevost, M. S., Redzej, A., Trokter, M. and Waksman, G. (2015). Secretion systems in Gram-negative bacteria: structural and mechanistic insights. Nat Rev Microbiol. 13(6): 343–359.
Meng, R., Xing, Z., Chang, J. Y., Yu, Z., Thongchol, J., Xiao, W., Wang, Y., Chamakura, K., Zeng, Z., Wang, F., et al. (2024). Structural basis of Acinetobacter type IV pili targeting by an RNA virus. Nat Commun. 15(1): 2746.
Strom, M. S. and Lory, S. (1993). Structure-function and biogenesis of the type IV pili. Annu Rev Microbiol. 47(1): 565–596.
Mattick, J. S. (2002). Type IV Pili and Twitching Motility. Annu Rev Microbiol. 56(1): 289–314.
Craig, L. and Altindal, T. (2019). Purification of Type IV Pili and Pilin Subunits. Methods Mol Biol.: 97–110.
Date, T., Inuzuka, M. and Tomoeda, M. (1977). Purification and characterization of F pili from Escherichia coli. Biochemistry. 16(25): 5579–5585.
Meng, R., Jiang, M., Cui, Z., Chang, J. Y., Yang, K., Jakana, J., Yu, X., Wang, Z., Hu, B., Zhang, J., et al. (2019). Structural basis for the adsorption of a single-stranded RNA bacteriophage. Nat Commun. 10(1): 3130.
Article Information
Publication history
Received: Jun 22, 2024
Accepted: Aug 27, 2024
Available online: Sep 25, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Microbiology > Microbial biochemistry > Protein
Biophysics > Electron cryotomography
Biochemistry > Protein > Self-assembly
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Chromogranin B Purification for Condensate Formation and Client Partitioning Assays In Vitro
AP Anup Parchure
JB Julia Von Blume
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5095 Views: 286
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Sep 2022
Abstract
Chromogranin B and other members of the granin protein family form condensates that recruit clients like proinsulin. The condensation in the lumen of trans-Golgi network (TGN) is critical for the biogenesis of secretory granules. Here, we describe a protocol to purify the tagged version of chromogranin B close to its native form at the TGN, which can then be utilized for microscopy-based assays to monitor condensate formation in vitro and client partitioning depending on the material properties of chromogranin B assemblies.
Key features
• First instance of purification of full-length and tagged version of members of the chromogranin family of proteins.
• Allows purification of proteins with post-translational modifications that are acquired en route in the secretory pathway, thus closely resembling their native form at the TGN.
Keywords: Phase separation Chromogranin B Protein purification Client-partitioning
Graphical overview
Background
Members of the chromogranin (CG) family of proteins are predominantly expressed in specialized secretory cells including the pancreatic beta cells and the cells of the neuroendocrine system. They play a critical role in driving secretory granule (SG) biogenesis [1–3]. Expression of either chromogranin A (CGA) and chromogranin B (CGB) or other granin proteins in non-specialized cells results in the formation of ectopic SG-like structures [2,4–6]. Experiments involving extracts or purified CGA, CGB, and secretogranin 2 (SCG2) from the adrenal or pituitary glands demonstrated aggregation of these proteins at an acidic pH (pH 5.2–5.5) and in presence of millimolar concentrations of calcium [6–10]. Since the pH used in previous studies is lower than that seen at the trans-Golgi network (TGN) and the calcium concentrations were much higher [10,11], the physiological significance of the aggregation in cargo sorting has remained elusive. Moreover, molecular mechanisms underlying higher-order assemblies of CGB and the impact of material properties of these assemblies on cargo sorting require further investigation.
We demonstrated that CGB forms condensates in milieu of the TGN, driven predominantly by the mildly acidic compartment of the TGN and independent of the requirement of divalent cations [12]. For this purpose, we developed a protocol using the PiggyBac transposon-based inducible mammalian expression system [13] to purify the superfolder GFP-tagged versions of CGB from the secreted medium. Moreover, by making different assemblies (condensates vs. aggregates) of CGB in the presence of calcium and zinc, respectively, we further demonstrate that the material properties of CGB assemblies drive client partitioning.
Materials and reagents
Materials
Plasmid for cloning human CGB gene (OriGene, RC201744L1)
RINS1 plasmid (Addgene, plasmid #107290)
2× Gibson master mix (NEB, catalog number: E2611)
Nhe1-HF (NEB, catalog number: R3131)
Not1-HF (NEB, catalog number: R3189)
Agarose (American Bio, catalog number: AB00972)
SYBR Safe DNA gel stain (Invitrogen, catalog number: S33102)
TAE Buffer (Thermo Fisher Scientific, catalog number: B49)
PB-T-PAF, PB-RN, and PBase [13]
Water, molecular biology grade (Sigma, catalog number: W4502)
Qiagen Gel Extraction kit (Qiagen, catalog number: 28704)
Mini prep kit (Takara Bio, catalog number: 740588.50)
LB ampicillin plates (Recombinant Technologies)
HEK293 cells (ATCC, catalog number: CRL-1573)
Lipofectamine 2000 (Thermo Fisher Scientific, catalog number: 11668027)
DMEM high glucose (Gibco, catalog number: 11965092)
Fetal bovine serum (FBS) (Gibco, catalog number:16000044)
Penicillin-Streptomycin-Glutamine (100×) (Gibco, catalog number: 10378016)
OptiMEM (Gibco, catalog number: 51985034)
Puromycin dihydrochloride (Thermo Fisher Scientific, catalog number: J67236.XF)
G418 disulfate (Thermo Fisher Scientific, catalog number: J63871.AB)
cOmpleteTM His-Tag purification resin (Millipore Sigma, catalog number: 5893682001)
A23187 (Millipore Sigma, catalog number: C7522)
Doxycycline monohydrate (LKT Labs, catalog number: D5898)
Aprotinin (Millipore Sigma, catalog number: A6106)
Imidazole (Spectrum, catalog number: IM105)
0.45 µm cellulose acetate filtration unit (Corning, catalog number: 430768)
Amicon® Ultra centrifugal filter (Millipore Sigma, catalog number: UFC5030)
Amicon® Ultra-4 centrifugal filter (Millipore Sigma, catalog number: UFC800324)
AmiconTM Ultra-15 centrifugal filter units (Millipore Sigma, catalog number: UFC903024)
NaOH (Sigma, catalog number: 221465-500G)
Glycerol (Fisher Scientific, catalog number: 02-002-937)
5 M NaCl solution (American Bio, catalog number: AB13198-01000)
Tris, 1 M Solution, pH 7.4 (American Bio, catalog number: AB14044-01000)
Sodium phosphate, dibasic, anhydrous (Na2HPO4) (J. T. Baker, catalog number: 3828-01)
Sodium phosphate, monobasic, monohydrate (NaH2PO4.H2O) (J. T. Baker, catalog number: 3818-01)
Cy3-labeled lysozyme (Lyz) (Nanocs, catalog number: LS1-S3-1)
Calcium chloride solution (Millipore Sigma, catalog number: 21115)
Zinc chloride (Sigma, catalog number: Z-4875)
1× DPBS (Gibco, catalog number: 14190250)
Chromatography columns (Bio-Rad, catalog number: 732-1010)
Glass-bottom imaging dishes (Cellvis, catalog number: D35-14-1.5-N)
250 mL filter system (Corning, catalog number: 430768)
PCR tubes (Thomas Scientific, catalog number: 1149K07)
1.5 mL microcentrifuge tubes (USA Scientific, catalog number: 1415-2500)
List of primers used for cloning of sfGFP and His tagged CGB using Gibson assembly (Table 1)
Table 1. List of primers
Primer name Sequence
CGB_sfGFP_6XHis F1 FP ggcggccatcacaagtttgtacagctagcatgcagccaacgctgcttctcagcctc
CGB_sfGFP_6XHis F1 RP caccggtggcgaccggtggatccaagcccctttggctgaatttctcagctatcttctgtagttcc
CGB_sfGFP_6XHis F2 FP ggaactacagaagatagctgagaaattcagccaaaggggcttggatccaccggtcgccaccgg
CGB_sfGFP_6XHis F2 RP ccagcacactggatcagttatctatgcggccgctcattagctgcccttgtacagctcgtccatgcc
Solutions
10× Na-P stock (see Recipes)
His-binding buffer (see Recipes)
Wash buffer (see Recipes)
Elution buffer (see Recipes)
Protein storage buffer (see Recipes)
Phase separation assay buffer (see Recipes)
NaOH solution for washing column (see Recipes)
Recipes
10× Na-P stock (500 mM, 500 mL)
33.1 g of Na2HPO4
2.4 g of NaH2PO4
pH 8.0
Store at 4°C
His-binding buffer (500 mL)
500 mM NaCl (14.61 g), 50 mM Na-P (50 mL stock) pH 6.8 (fresh)
Wash buffer
His-binding buffer with 10 mM imidazole: 150 mL of His-binding buffer + 0.1 g of imidazole (fresh).
Elution buffer
His-binding buffer with 250 mM imidazole: 10 mL of His-binding buffer + 0.17 g of imidazole (fresh).
Protein storage buffer (50 mL)
200 mM Tris-HCl (1 M Tris-HCl 10 mL), 500 mM NaCl (5 M NaCl 5 mL), 10% glycerol (5 mL), pH 6.8.
Store at 4 °C for a week if purifying more than one protein within a week.
Phase separation assay buffer (40 mL)
25 mM Tris-HCl (1 M Tris-HCl 1 mL), 150 mM NaCl (5 M NaCl 1.2 mL), 2.5% glycerol (1 mL), pH 6.1 (fresh).
NaOH solution for washing column
50 mL of 0.5 N NaOH (1 g)
Store at room temperature.
Equipment
Thermal cycler (Eppendorf, model: nexus eco)
Centrifuge (Eppendorf, model: 5425)
Centrifuge (Eppendorf, model: 5910 R)
Centrifuge (Thermo Scientific, model: Sorvall Legend Micro 21R)
S-4x Universal rotor
Zeiss LSM 880 Confocal with Airyscan (63×; 1.4 N.A. oil objective)
Nanodrop (Thermo Scientific, model: NANODROP ONEC)
Procedure
Cloning to generate the PB-T-PAF-CGB_sfGFP_6XHis construct
The construct for expression of sfGFP and 6× His-tagged human CGB protein under a doxycycline-inducible promoter was generated by Gibson assembly. The coding region of the human CGB gene was PCR amplified using the expression plasmid from OriGene (100 ng for a 50 µL PCR reaction). sfGFP (containing the monomerizing mutation) was amplified by PCR amplification from the RINS1 plasmid (100 ng for a 50 µL PCR reaction). PB-T-PAF vector was digested using the restriction enzymes Nhe1-HF and Not1-HF (20 units each per reaction). PCR primers were designed to ensure overlap with the vector and individual fragments. The DNA sequence encoding the 6× His-tag was also incorporated in the reverse primer while amplifying the sfGFP fragment.
Run the PCR fragments on a 1% agarose gel containing SYBR Safe DNA gel stain using TAE buffer. Excise bands of appropriate sizes and purify by gel extraction using the Qiagen gel extraction kit.
Digest the PB-T-PAF vector.
Mix cut vector and individual fragments in 1:3 molar ratios in the Gibson master mix. Make up the total reaction volume to 20 µL using molecular biology–grade water and incubate in a thermal cycler at 50 °C for 1 h.
Transform 2 µL of the Gibson reaction into competent Omni max 2 E. coli.
Extract DNA from individual colonies from Luria agar containing ampicillin plates using the mini prep kit. Confirm positive clones by sequencing individual clones.
Generating stable cell lines for protein expression
Culture and maintain HEK293 cells in DMEM high glucose supplemented with 10% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin (complete medium) in 5% carbon dioxide at 37 °C. One day before transfection, seed approximately 1 million cells in a 6-well dish in the medium mentioned above. Cell culture and transfection are done in a cell culture hood.
The next day, transfect HEK293 cells using PB-T-PAF-CGB_sfGFP_6XHis, PB-RN, and PBase (8:1:1; total DNA: 1.5 µg) and Lipofectamine 2000.
For transfection, incubate DNA in a sterile 1.5 mL microcentrifuge tube in 100 µL of OptiMEM. In a separate tube, add 3 µL of Lipofectamine 2000 to 100 µL of OptiMEM. After 5 min, mix the contents of the two tubes and incubate for 20–30 min.
In the meantime, add fresh growth medium to cells and the transfection mixture in a dropwise fashion (steps B1–4; Figure 1).
Forty-eight hours post-transfection, initiate selection using a combination of 10 µg/mL puromycin dihydrochloride and 500 µg/mL G418 disulfate.
Maintain cells under antibiotic selection for at least a week, and then expand them and freeze them at -80 °C in freezing medium (complete medium containing 10% DMSO). Store them until further use in liquid nitrogen storage.
Figure 1. Scheme for transfecting HEK293 cells for generating stable lines expressing CGB-sfGFP under doxycycline-inducible promoter. Forty-eight hours after transfection, cells are subjected to antibiotic selection.
Protein expression and purification from cell culture supernatant
Harvesting cell culture supernatant
Thaw stable HEK293 cells expressing CGB-sfGFP-6X His under doxycycline-inducible promoter and expand them in at least ten 15 cm dishes. For proteins that, in pilot assays, were secreted to a lesser extent in the supernatant, we have gone up to fifteen 15 cm dishes. Additionally, when a lower percentage of cells showed us fluorescent protein expression after doxycycline induction (less than 50%; see below in notes and troubleshooting), we used 15 dishes.
When the cells are confluent, wash two times in 1× PBS and then incubate in 20 mL of serum-free DMEM containing 1 µg/mL doxycycline, 1µg/mL aprotinin, and 1µg/mL A23187 for approximately 16–20 h. Steps C1a and C1b were done in a cell culture hood.
Figure 2. Scheme for collecting medium from HEK293 prior to protein purification
The next day, collect and pool the medium from all the plates by centrifuging at 2,000 rpm (859× g) for 10 min at 4 °C. Filter using 0.45 µm filter and set aside to load on the column as described subsequently (Steps C1a–c; Figure 2).
Protein purification using Ni-NTA columns
Cut the column at the base using a scissor and then fill with PBS with 2–3 mL of slurry containing Ni-NTA beads (His tagged purification resin) and tightly pack between 1–1.5 cm of bed volume. Allow 50 mL of PBS to flow through.
Equilibrate the column with 100 mL of His-binding buffer.
Load the supernatant containing protein and allow it to run via gravity flow. Maintain a flow rate of one drop per 4–5 s. In our case, we could do this by simply adjusting the height of the reservoir from which the supernatant was loaded onto the column via the tubing.
After all the supernatant has passed through the column, wash the column in 150 mL of wash buffer.
Elute the protein in elution buffer (10 mL). Collect 1 mL fractions in Eppendorf tubes. Because of the GFP tag, visually green fractions are collected and pooled together.
To get rid of imidazole, buffer exchange the protein with protein storage buffer three times using 50 mL AmiconTM Ultra-15 centrifugal filter units with a 30 kDa cutoff by centrifugation at 4,500 rpm (4,347× g) for 15 min. Before adding the proteins, equilibrate the filter using storage buffer. In the first run, 2–3 mL of protein solution is topped with protein storage buffer to fill the tube (12 mL total volume). Concentrate the protein to 1 mL. Add 11 mL of buffer and reconcentrate the protein to 1 mL. Repeat this step two more times (Steps C2a–f; Figure 3).
Figure 3. Scheme for protein purification using Ni-NTA columns. Protein-containing fractions (green) are pooled together and buffer exchanged into protein storage buffer and aliquoted and flash frozen using liquid nitrogen.
Aliquot the protein in 100 µL fractions, flash freeze in liquid nitrogen, and store at -80 °C until further use.
Figure 4. A typical Coomassie-stained SDS gel with purified CGB-sfGFP run on a denaturing gel. The band is seen at a size corresponding to 130 kDa.
Wash the column with 50 mL of 0.5 N NaOH and then wash it with 100 mL of water. Pass 50 mL of 70% ethanol and cap and store the column for reuse at 4 °C in a cold room. We have only used the column a maximum of two times and only in instances when we were purifying the same protein within a month. We have performed protein purification in one day beginning from collection of the medium until storing the purified protein at -80 °C.
Caution:
1) Ensure there are not many dead cells at the time of collecting medium.
2) Ensure there are no air bubbles while packing the column.
Notes and troubleshooting:
1) Protein sticking while concentrating: In our hands, it was better to concentrate the protein at room temperature as it led to minimal sticking of the protein to the membrane during concentration.
2) Cells floating after incubation in serum-free medium: If the cells are alive, it should not be a problem. A solution could be plating the cells on poly-lysine-coated dishes, even though we have not tried this.
3) Cleaning impurities: Assess the quality of the protein by running 1–5 µg of protein on an SDS-PAGE gel followed by Coomassie staining. For CGB-sfGFP, we see a band at 130 kDa (Figure 4). If there are many lower-molecular-weight bands, then that indicates degradation or poor-quality purification, in which case the preparation should be repeated. Purity higher than 90% is good for biochemical experiments. We have not tried extensively with CGB-sfGFP, but we used gel filtration for purification of mCherry-tagged proinsulin using the same protocol with the only exception being that FLAG-tag was used as an affinity tag.
4) The protocol can be widely applied for the purification of mutant forms of chromogranin B and other granin proteins, provided they are secreted from HEK293 cells. Importantly, proteins purified using this method retain the post-translational modifications acquired during their transit through the secretory pathway (Figure 5). After the generation of stable lines, we purified proteins when 50% of the population was fluorescent after doxycycline addition. In some instances, when we felt that the number of fluorescent cells was much lower, we resorted to increasing the number of dishes (starting material) and, in some instances, redoing the transfection and selection, expanding the cell line, and purifying the protein right away before freezing cells.
5) A pilot secretion assay must be performed before large-scale purification. Grow cells to confluency in a 10 cm dish and then induce protein production in serum-free medium as described above. Concentrate the medium and run on SDS-PAGE gels followed by Coomassie staining; see if there is a strong band at the desired size. Alternatively, one can also use Ni-NTA beads to incubate with the concentrated medium. Since the proteins are GFP tagged, beads would turn green post-binding in a cold room for 1 h by rotating it end-to-end. The protein can be eluted after washing the beads with 1× PBS (three times) and boiling in SDS sample buffer. Run samples on SDS-PAGE gels followed by Coomassie staining to monitor the band at the appropriate size.
Figure 5. Purified CGB-sfGFP was subjected to mass spectrometry analysis to detect post-translational modifications at the facility at Yale university. Labeled in red are the sulfation on tyrosine residues. The presence of the post-translational modifications confirms that the purified protein passed through the secretory pathway where it acquires these modifications and suggests that the protein purified is close to the native form at the TGN. This is critical considering that post-translational modifications have been shown to play an important role in governing condensate formation in many different contexts.
Condensate formation and client partitioning assay
Buffer exchanging protein in phase separation buffer at pH 6.1
Thaw the protein solution on ice.
Buffer exchange the protein to phase separation buffer using 0.5 mL Amicon® Ultra centrifugal filter with a 30 kDa cutoff. Repeat the process at least three times. Take 200 µL of protein and fill the tube with 300 µL of buffer. Spin at 12,000 rpm (13,523× g) and concentrate to 100 µL. Repeat the process two more times. In case the initial yields are low, start with a larger volume, bring it down to 100 µL, and then proceed to three buffer exchanges.
Measure the protein concentration using Nanodrop by reading the absorbance at 280 nm. Since CGB-sfGFP forms condensates at pH 6.1, spin down the buffer-exchanged protein solution at full speed (21,100× g) in a tabletop microcentrifuge at 4 °C for 20 min.
Collect the supernatant in a separate Eppendorf tube and store it on ice. Immediately proceed to set up client partitioning assays.
Client partitioning assay using Cy3-tagged Lyz in the presence of calcium and zinc
Buffer exchange Cy3-tagged Lyz in the phase separation buffer by using 0.5 mL Amicon® Ultra-4 centrifugal filter with a 3 kDa cutoff three times.
Dilute calcium chloride and zinc chloride solution with phase separation buffer to a final concentration of 20 mM in two separate tubes. To each tube, add Cy3-tagged Lyz to a final concentration of 1 µM. To each tube, add CGB-sfGFP to a final concentration of 2.5 µM. Gently tap the tubes and incubate at room temperature for 5 min.
Plate the solution from each tube on an imaging dish from Cellvis.
After 5 min, image the samples on a Zeiss 880 confocal microscope using a 63×/1.4 oil objective at room temperature.
Acquire images in both GFP and Cy3 channels for droplets formed using calcium and aggregates formed using zinc (Figure 6). A 488 nm laser was used at 0.2% and a 561 nm laser was used at 5%. The pixel dwell time was 0.67 µs.
Figure 6. Scheme for client partitioning assay form monitoring differential client recruitment to calcium-induced condensates and zinc-induced CGB aggregates
Note: Client partitioning assays can also be set by mixing different client proteins including Cy3-tagged proinsulin by directly mixing the protein with CGB-sfGFP or by inducing phase separation using PEG 8000.
Data analysis for measuring relative client partitioning
Open images acquired in the .czi format on Zeiss 880 in Fiji.
Draw a region of interest (ROI) on the condensate image using the GFP channel (green).
In Fiji, go to Analyze > Tools > ROI manager > Add[t].
Click on measure and you will get intensity values in the results tab. Open the Cy3 channel, click on the ROI in the ROI manager, and again click on measure.
Follow the same procedure for the images of CGB aggregates using zinc (Figure 7).
Copy the values in an Excel sheet and normalize the values (mean intensity) in the Cy3 channel to GFP channel for both calcium and zinc conditions. The normalized values represent the amount of client (Cy3-Lyz) that partitions into condensate or aggregate relative to the amount of CGB-sfGFP.
Do this for multiple ROIs.
Analyze the differences in the relative partitioning from multiple ROIs using a t-test and a two-tailed distribution in Prism.
The data can also be analyzed by quantifying the signals of CGB-sfGFP or Cy3-LyZ from the assemblies; that from the background and absolute partitioning can be quantified, which is the ratio of signals within the condensate/aggregate to that outside. The data can also be represented as the amount of signal in condensate/aggregate to total signal (inside + outside). However, we have only performed the relative partitioning analysis in the manuscript where the client portioning assay was used [12].
Figure 7. Screenshot explaining data analysis in Fiji to measure relative client (Cy3-Lyz; red) partitioning into CGB-sfGFP (green) assemblies generated in the presence of either zinc or calcium. All images have the same size; however, for the purpose of the screenshot, they have been adjusted such that only a part of some of the images is seen. Yellow regions depict ROI. Labeling was done in Adobe Illustrator and Microsoft Power Point.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Parchure et al. [12]. Liquid–liquid phase separation facilitates the biogenesis of secretory storage granules. Journal of Cell Biology (Figure 1B, Figure 5A, B).]
Acknowledgments
Anup Parchure is funded by a Project and Feasibility Award from the Yale Diabetes Research Center (P30 DK04573) and would like to acknowledge the support from R01DK129466 awarded to Jonathan S. Bogan. Julia von Blume is funded by a MIRA grant from NIGMS (1R35GM149293-01) and the NIDDK Innovative Science Accelerator Program (ISAC) Award (DK128851) and would like to acknowledge support from a Project and Feasibility award from Yale Diabetic Research Center (GR112420). We would like to acknowledge support from The Mass Spectrometry (MS) & Proteomics Resource of the W.M. Keck Foundation Biotechnology Resource Laboratory for detection of post-translational modifications on purified CGB-sfGFP. The protocol was adapted from Li et al. [13] PNAS. We used BioRender for making figures for this manuscript. The licenses for all the figures are listed below.
Graphical overview: Created in BioRender. Parchure, A. (2024) BioRender.com/z45l203
Figure 1: Created in BioRender. Parchure, A. (2024) BioRender.com/o81v327
Figure 2: Created in BioRender. Parchure, A. (2024) BioRender.com/a76y056
Figure 3: Created in BioRender. Parchure, A. (2024) BioRender.com/c76z345
Figure 6: Created in BioRender. Parchure, A. (2024) BioRender.com/t30m682
Competing interests
There are no conflicts of interest or competing interest.
References
Bearrows, S. C., Bauchle, C. J., Becker, M., Haldeman, J. M., Swaminathan, S. and Stephens, S. B. (2019). Chromogranin B regulates early-stage insulin granule trafficking from the Golgi in pancreatic islet β-cells. J Cell Sci. 132(13): e231373.
Kim, T., Tao-Cheng, J. H., Eiden, L. E. and Loh, Y. (2001). Chromogranin A, an “On/Off” Switch Controlling Dense-Core Secretory Granule Biogenesis. Cell. 106(4): 499–509.
Tooze, S. A. and Huttner, W. B. (1990). Cell-free protein sorting to the regulated and constitutive secretory pathways. Cell. 60(5): 837–847.
Rustom, A., Bajohrs, M., Kaether, C., Keller, P., Toomre, D., Corbeil, D. and Gerdes, H. (2002). Selective Delivery of Secretory Cargo in Golgi‐Derived Carriers of Nonepithelial Cells. Traffic. 3(4): 279–288.
Wacker, I., Kaether, C., Krömer, A., Migala, A., Almers, W. and Gerdes, H. H. (1997). Microtubule-dependent transport of secretory vesicles visualized in real time with a GFP-tagged secretory protein. J Cell Sci. 110(13): 1453–1463.
Huh, Y. H., Jeon, S. H. and Yoo, S. H. (2003). Chromogranin B-induced Secretory Granule Biogenesis. J Biol Chem. 278(42): 40581–40589.
Colomer, V., Kicska, G. A. and Rindler, M. J. (1996). Secretory Granule Content Proteins and the Luminal Domains of Granule Membrane Proteins Aggregate in Vitro at Mildly Acidic pH. J Biol Chem. 271(1): 48–55.
Gerdes, H. H., Rosa, P., Phillips, E., Baeuerle, P. A., Frank, R., Argos, P. and Huttner, W. B. (1989). The primary structure of human secretogranin II, a widespread tyrosine-sulfated secretory granule protein that exhibits low pH- and calcium-induced aggregation. J Biol Chem. 264(20): 12009–12015.
Chanat, E. and Huttner, W. B. (1991). Milieu-induced, selective aggregation of regulated secretory proteins in the trans-Golgi network. J Cell Biol. 115(6): 1505–1519.
Paroutis, P., Touret, N. and Grinstein, S. (2004). The pH of the Secretory Pathway: Measurement, Determinants, and Regulation. Physiology. 19(4): 207–215.
Pizzo, P., Lissandron, V., Capitanio, P. and Pozzan, T. (2011). Ca2+ signalling in the Golgi apparatus. Cell Calcium. 50(2): 184–192.
Parchure, A., Tian, M., Stalder, D., Boyer, C. K., Bearrows, S. C., Rohli, K. E., Zhang, J., Rivera-Molina, F., Ramazanov, B. R., Mahata, S. K., et al. (2022). Liquid–liquid phase separation facilitates the biogenesis of secretory storage granules. J Cell Biol. 221(12): e202206132.
Li, Z., Michael, I. P., Zhou, D., Nagy, A. and Rini, J. M. (2013). Simple piggyBac transposon-based mammalian cell expression system for inducible protein production. Proc Natl Acad Sci USA. 110(13): 5004–5009.
Article Information
Publication history
Received: Apr 22, 2024
Accepted: Sep 1, 2024
Available online: Sep 25, 2024
Published: Oct 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,096 | https://bio-protocol.org/en/bpdetail?id=5096&type=0 | # Bio-Protocol Content
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Peer-reviewed
Single Cell Isolation from Human Diabetic Fibrovascular Membranes for Single-Cell RNA Sequencing
KS Katia Corano Scheri
TT Thomas Tedeschi
AF Amani A. Fawzi
Published: Vol 14, Iss 20, Oct 20, 2024
DOI: 10.21769/BioProtoc.5096 Views: 308
Reviewed by: Pilar Villacampa AlcubierreYu LiuPreeti Yadav
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Original Research Article:
The authors used this protocol in JCI Insight Nov 2023
Abstract
Single-cell transcriptomic analyses have emerged as very powerful tools to query the gene expression changes at the single-cell level in physiological and pathological conditions. The quality of the analysis is heavily dependent on tissue digestion protocols, with the goal of preserving thousands of single live cells to submit to the subsequent processing steps and analysis. Multiple digestion protocols that use different enzymes to digest the tissues have been described. Harsh digestion can damage certain cell types, but this might be required to digest especially fibrotic tissue as in our experimental condition. In this paper, we summarize a collagenase type I digestion protocol for preparing the single-cell suspension from fibrovascular tissues surgically removed from patients with proliferative diabetic retinopathy (PDR) for single-cell RNA sequencing (scRNA-Seq) analyses. We also provide a detailed description of the data analysis that we implemented in a previously published study.
Key features
• Single-cell suspension from fibrovascular membranes isolated from PDR patients.
• Single-cell RNA sequencing analyses performed using Seurat package in RStudio.
• Trajectory analyses or pseudotime analyses to study the trajectory over (pseudo)time of specific cell types.
• This protocol requires Illumina HiSEQ4000 instrument and knowledge of R and RStudio language for the analyses.
Keywords: scRNA-Seq scRNA-Seq data analysis Sequencing Surgical specimens Fibrovascular membranes
Graphical overview
Fibrovascular membrane isolation and sequencing
Background
Proliferative diabetic retinopathy (PDR) is a late-stage complication of diabetes, responsible for vision loss in diabetic patients. The formation of fibrovascular membranes and scar tissue in the pre-retinal space leads to retinal traction and detachment [1]. Understanding the molecular features of the cells contributing to fibrovascular membrane formation and the cell–cell interactions is crucial for identifying new therapeutic targets that improve treatment approaches and patients’ quality of life.
scRNA-Seq is a powerful technique that allows us to study the molecular profile of each single cell. Since the first paper on scRNA-Seq was published in 2009, studies on this technique have provided insightful information in several fields, followed by exponential growth in the last decade [2–8]. In the context of retinal biology, this approach was fundamental for building a comprehensive transcriptome atlas of fetal and adult retinas and for dissecting the retinal developmental stages [9–11]. scRNA-Seq assays have also revealed the complexity and heterogeneity of the different cell types in the retina and aided in uncovering cell-specific gene expression changes in pathological conditions [12–17]. Single-cell transcriptomics also enables the study of individual pathogenic cell populations in the context of fibrosis at quite high resolution [18–22].
Despite significant improvements, scRNA-Seq is still a challenging technique [23]. Generating high-quality single-cell suspensions from any tissue is critical to preserve their expression profile and ensure meaningful downstream transcriptome data analysis. Several parameters in the dissociation protocol can compromise the viability of the cells and potentially impact the quality of the scRNA-Seq data. Preparing cell suspensions might be difficult for tissue samples, especially for fibrotic tissue, where gentle dissociation might not be sufficient to break the fibers of scar tissue, leading to inefficient cell yield after processing. On the other hand, an excessively harsh digestion might differentially damage cells that are more fragile [23,24].
Papain-based protocols have been shown to successfully dissociate retinal tissue and have been described in several scRNA-Seq studies of retina tissue and organoids [16,23,25]. Another enzyme described for tissue digestion is Collagenase (I and IV), which was described to digest fibrovascular membranes from proliferative diabetic retinopathy patients as well as mouse retinas and other tissues into single-cell suspensions [18,21,26–28]. In our experiments, we used a Collagenase I–based protocol [29].
Another important factor in the scRNA-Seq experiments is the downstream data analyses. Throughout the analysis, it is important to include filtering steps in data analysis processing with the aim of excluding low-quality cells or empty droplets that have very few genes and cell doublets that may exhibit an aberrantly high gene count.
Our protocol is quite efficient in digesting the fibrovascular membranes from PDR patients [29], and we were able to preserve a lot of cell types with a high number of genes detected per cell.
Materials and reagents
Biological materials
Fibrovascular membranes surgically removed from proliferative diabetic retinopathy (PDR) patients
Reagents
Collagenase Type I (Worthington, catalog number: LS004194)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906)
Ethylenediaminetetraacetic acid (EDTA) (Amresco, catalog number: E522)
Illumina TruSeq stranded mRNA (Illumina, catalog number: 20020594)
Chromium Single Cell 3' v3 kit (10X Genomics, catalog number: PN-1000268)
Hank's balanced salt solution (HBSS) (Thermo Fisher Scientific, catalog number: 14025092)
1× Dulbecco’s phosphate-buffered saline (DPBS) with calcium and magnesium (Thermo Fisher Scientific, catalog number: 14040133)
1× Dulbecco’s phosphate-buffered saline (DPBS) without calcium and magnesium (Thermo Fisher Scientific, catalog number: 14190094)
Solutions
Collagenase I solution (see Recipes)
Bovine serum albumin (BSA)-EDTA (see Recipes)
Recipes
Collagenase I solution
Reagent Final concentration Quantity or Volume
Collagenase type I 2 mg/mL 500 µL/sample
PBS with Ca2+ and Mg2+
Bovine serum albumin (BSA)-EDTA
Reagent Final concentration Quantity or Volume
Bovine serum albumin (BSA) 1% 500 µL/sample
EDTA 2 mM
PBS w/o Ca2+ and Mg2+
Laboratory supplies
1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 022431021)
1 mL pipette (USA Scientific, catalog number: 7110-1000)
Laboratory tips (TipOne, catalog number: 1111-2720)
Equipment
Agilent 2100 Bio-analyzer
Illumina HiSEQ4000
Microcentrifuge (Eppendorf, model: 5424R)
Software and datasets
R software (free download from https://cran.r-project.org/bin/windows/base/) (R version 4.3.2, 2023-10-31)
RStudio (free download from https://posit.co/download/rstudio-desktop/) (Version 2023.03.1+446 (2023.03.1+446)
Seurat (version 5.1.0) and Monocle (Version 2.32.0) packages for RStudio
All raw data have been deposited to GEO (accession no. GSE245561) and trajectory inference analysis code was deposited on GitHub (https://github.com/katiacoranoscheri/PDR-trajectory-analysis).
Procedure
Fibrovascular membrane collection
Patients with fibrovascular traction diabetic detachments undergo standard 23-gauge pars plana vitrectomy. Using a combination of microsurgical instruments, dissect the epiretinal fibrovascular membranes and extract large pieces from the eye. These extracted membranes need to be immediately transferred in HBSS on ice.
Tissue digestion, single-cell suspension preparation, and sequencing
After surgery, place the PDR fibrovascular membranes in HBSS and store them on ice.
Prepare a solution of 1× DPBS with calcium and magnesium containing 2 mg/mL Collagenase Type I for tissue digestion (Figure 1).
Pipette 500 μL of the digestion solution to each sample and dissociate them for 30 min at 37 °C, shaking the tubes manually every 5 min.
Gently triturate the samples by pipetting the solution 20 times with a p1000 pipette.
After digestion and trituration, use a 70 μm filter to remove any clumps and debris.
After dissociation, spin down the cell suspension at 400× g for 8 min at 4 °C and carefully discard the supernatant (Figure 1).
Resuspend the cell pellet in 500 μL of 1× DPBS without calcium and magnesium with 1% BSA and 2 mM EDTA and keep the samples on ice until further processing (Figure 1).
Figure 1. Overview of the dissociation procedure for single-cell suspension preparation
Before sequencing, the cell number and viability need to be assessed using an automated cell counter with AO/PI dye.
All our scRNA-Seq experiments were performed at the Functional Genomics Core at the University of Chicago (Chicago, Illinois, USA). Single-cell libraries were generated using a Chromium Single Cell 3' v3 kit (10× Genomics) and RNA quality and quantity were assessed using an Agilent Bio-analyzer. RNA-Seq libraries were prepared using Illumina mRNATruSEQ kits following manufacturer instructions (Illumina). Library quality and quantity were checked using an Agilent Bioanalyzer. In our study, we implemented the scRNA-Seq analysis based on the Drop-Seq method that was first published in 2015 [30] and sequenced using the Illumina HiSEQ4000 (paired-end 100 bp) following the manufacturer’s reagents and instructions.
Data analysis
Pre-processing workflow and quality check
Raw sequencing data need to be converted into a FASTQ format, and the files need to be aligned to the hg38 reference genome (for human data) provided by 10× Genomics. Seurat package (Satija Lab, NYGC) is used to analyze the data.
If using multiple samples, preprocess, normalize, and scale each individual raw data sample following the Seurat pipeline (https://satijalab.org/seurat/articles/pbmc3k_tutorial).
Consider as outlier cells with <500 or >5000 unique features as well as cells with >25% mitochondrial RNA.
Perform a doublet detection analysis and remove the doublet using the scDblFinder pipeline [31].
Doublet detection and removal can be performed using the standard parameters of 1% doublets per 1000 droplets sequenced. Copy the scDblFinder metadata from the new object into the original dataset.
Droplets identified as "singlets" can be used for subsequent analysis and doublets are discarded.
Sample integration, scaling, and clustering
Integrate the samples using the canonical correlation analysis (CCA) with the integration pipeline in Seurat (https://satijalab.org/seurat/articles/integration_introduction.html). Only variable features present in all samples are selected for dataset integration.
Following CCA integration, scale the dataset and calculate the principal components for plotting and clustering of the integrated dataset. Use ElbowPlot() to estimate the selection of which principal components can be included.
Following dimensionality estimation, “r full.dataset.dims” dimensions are used for UMAP projection and cluster identification of the integrated dataset. Cluster the CCA-aligned data using a Louvain algorithm implemented in Seurat at a resolution of 0.1 and r full.dataset.dims dimensions.
Implement the functions FindNeighbors and FindClusters to cluster the cells that you can then visualize with the function RunUMAP.
Calculate the percentage of cells in each cluster for each sample and, for statistical analysis, perform the hypergeometric distribution analysis using the phyper function [phyper (q=, m=, n=, k=, lower.tail = TRUE, log.p = TRUE)]. In the phyper function, # q = cells in cluster from sample, # m = total cells from that sample, # n = total cells from all other samples, # k = total cells in cluster. Consider P < 0.001 for statistical significance.
Differential gene expression analysis and re-clustering
Perform a differential expression analysis using “FindAllMarkers” function (min.pct = 0.25, log2FC > 1) to identify the cell clusters.
For the visualization of specific markers, use the data visualization methods from Seurat pipeline (https://satijalab.org/seurat/articles/visualization_vignette). DotPlot, VlnPlot, and FeaturePlot are examples of visualization of the data (Figure 2, first output, DotPlot).
Figure 2. Workflow of the scRNA-Seq data analysis and expected output. Images from Corano Scheri et al. [29].
Save the object with the SaveRDS function.
After identification of all cell clusters, re-cluster each cell type individually with the “subset” function indicating the number of the cluster to analyze separately [e.g., stromal <- (subset(original Seurat object, idents = 4))].
Run PCA again, then implement the functions FindNeighbors and FindClusters to find the subclusters of each cell type and visualize them with the function RunUMAP.
Perform a differential gene expression analysis to identify the subclusters in each cell type.
Save the object of each cell cluster individually with the SaveRDS function.
Pathway enrichment, inference, and ligand-receptor analyses
Select the most significant differentially upregulated and downregulated genes (log2FC > 1, logFC<-1 and p-adj < 0.01) to perform a pathway enrichment analysis in each subcluster.
Use the g: GOSt functional profiling tool available on the g: Profiler web server (version e99_eg46_p14_f929183) to identify significant pathways upregulated or downregulated in the clusters.
Generate the dotplots in R using PathFindR package in RStudio (Figure 2, second output).
Monocle3 workflow (https://cole-trapnell-lab.github.io/monocle3/docs/trajectories/) (Becht, Cao, Qiu, Trapnell) can now be used to perform cell inference trajectory analysis or pseudotime analysis. We implemented this workflow to perform the pseudotime analysis on the stromal cluster.
Import the Seurat object (previously saved as .rds file) into Monocle with the Monocle3 conversion tool as. cell_data_set () function.
Use Monocle tools to order the cells.
The differentialGeneTest function can be used to identify the differentially expressed genes among clusters along the trajectory and then generate a UMAP graph to illustrate the trajectory across the clusters (Figure 2, third output).
Perform a ligand-receptor analysis and evaluate the interactions between the cell types using CellphoneDB module.
Calculate the mean gene expression values and p-values from the cell barcode and counts extracted from the Seurat object and use it in conjunction with the statistical_analysis module.
Use the dot_plot module of CellphoneDB with the statistical analysis to generate the dot plots.
The description of these analyses can be found in the “Methods” section of the paper Corano Scheri et al. [29] JCI Insight. The raw data are available on GEO under accession number GSE245561. The code for the inference analysis has been uploaded on GitHub ((https://github.com/katiacoranoscheri/PDR-trajectory-analysis).
Validation of protocol
This protocol has been used in the following research article:
Corano Scheri et al. [29] JCI Insight. Single-cell transcriptomics analysis of proliferative diabetic retinopathy fibrovascular membranes reveals AEBP1 as fibrogenesis modulator.
General notes and troubleshooting
General notes
Limitations of our study might include a small sample size, which may reflect a general limitation of certain cells’ ability to withstand the technical process of single-cell isolation. It is recommended to have a minimum of 10,000 cells with > 90% viability to sequence and allow subsequent further analyses.
Troubleshooting
Problem 1: Cell number and viability can be low throughout the experiment.
Solution(s): It is recommended to use as many specimens as possible and to optimize the protocol, time, and reagents based on the tissue to maximize the yield and viability of the cells. It is important to take into consideration that different cell types and tissues might respond differently to single-cell digestion protocols that need to be changed accordingly.
Acknowledgments
The present work was supported by the following grants: NIH-R01-EY30121-A1 and Juvenile Diabetes Research Foundation Grant INO-2022-1112-A-N (AAF).
This protocol was described and validated in the following research article: Corano Scheri et al. [29], JCI Insight (2023), DOI: 10.1172/jci.insight.172062.
Competing interests
A.A.F. is a consultant to Regeneron, Roche/Genentech, Boehringer Ingelheim, RegenXbio, and T.T. is currently employed at Regeneron, but these entities did not have any relevant role in this article.
Ethical considerations
All human samples were collected at Northwestern, and the study was considered by our Institutional Review Board as exempt from informed consent and exception was granted from Northwestern University (detailed information about the patients are reported in the original paper Corano Scheri et al. [29], JCI Insight).
References
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Miranda, A. M. A., Janbandhu, V., Maatz, H., Kanemaru, K., Cranley, J., Teichmann, S. A., Hübner, N., Schneider, M. D., Harvey, R. P., Noseda, M., et al. (2023). Single-cell transcriptomics for the assessment of cardiac disease. Nat Rev Cardiol. 20(5): 289–308.
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Svensson, V., Natarajan, K. N., Ly, L. H., Miragaia, R. J., Labalette, C., Macaulay, I. C., Cvejic, A. and Teichmann, S. A. (2017). Power analysis of single-cell RNA-sequencing experiments. Nat Methods. 14(4): 381–387.
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Article Information
Publication history
Received: Jun 5, 2024
Accepted: Sep 5, 2024
Available online: Sep 29, 2024
Published: Oct 20, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,097 | https://bio-protocol.org/en/bpdetail?id=5097&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Efficient Gene-Editing in Human Pluripotent Stem Cells Through Simplified Assembly of Adeno-Associated Viral (AAV) Donor Templates
BC Berta Marcó de La Cruz
SM Sanhita Mitra
BH Bingqing He
MÇ Melis Çelik
DK Debora Kaminski
ES Erik Smedler
FS Fredrik H. Sterky
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5097 Views: 416
Reviewed by: Rajesh RanjanGuanghui Yang Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Neuroscience Apr 2024
Abstract
Gene-edited human pluripotent stem cells provide attractive model systems to functionally interrogate the role of specific genetic variants in relevant cell types. However, the need to isolate and screen edited clones often remains a bottleneck, in particular when recombination rates are sub-optimal. Here, we present a protocol for flexible gene editing combining Cas9 ribonucleoprotein with donor templates delivered by adeno-associated virus (AAV) vectors to yield high rates of homologous recombination. To streamline the workflow, we designed a modular system for one-step assembly of targeting vectors based on Golden Gate cloning and developed a rapid protocol for small-scale isolation of AAV virions of serotype DJ. High homology-directed repair (HDR) rates in human pluripotent stem cells (hPSCs), ~70% in ACTB and ~30% in LMNB1, were achieved using this approach, also with short (300 bp) homology arms. The modular design of donor templates is flexible and allows for the generation of conditional and/or complex alleles. This protocol thus provides a flexible and efficient strategy workflow to rapidly generate gene-edited hPSC lines.
Key features
• Versatile approach combining AAV-DJ donors and CRISPR ribonucleoproteins, providing an efficient method for long and short edits, insertions, and deletions in human pluripotent stem cells
• One-step cloning method for rapid generation of customized AAV donor plasmids
• Simplified AAV purification pipeline for ready-to-infect virion preparations
Keywords: AAV purification CRISPR/Cas9 Gene-editing Homologous recombination Human pluripotent stem cells Golden Gate assembly
Graphical overview
Background
The increasing availability of human genetic data offers unprecedented opportunities to understand, diagnose, and treat human diseases, but challenges remain in our ability to interpret the functional significance of specific genetic variants. At the same time, CRISPR/Cas-based gene-editing technologies have vastly simplified the means to introduce specific variants for functional interrogations [1–3]. Human pluripotent stem cells (hPSCs), including human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs), are attractive model systems for such functional studies as they allow experimental manipulations in relevant human cell types [4,5]. Of particular importance, hPSCs allow disease modeling in cell types of the central nervous system, including neurons, astrocytes, and microglia [6], which are generally inaccessible from patients yet are highly relevant to study in the context of psychiatric and neuropsychiatric diseases such as autism and schizophrenia, which are both largely attributable to genetic variants that demand functional studies.
In a typical gene-editing experiment, Cas9 nuclease is introduced to a bulk of cells together with a gene-specific guide RNA (gRNA) and a repair template for homology-directed repair (HDR). This strategy provides a means for flexible and precise edits ranging from single-base-pair substitutions to large insertions. However, sub-optimal editing rates may require tedious screening of clones to obtain cells with the desired edit(s), in particular when bi-allelic edits are desired. Moreover, clonal isolation involves the inevitable risk of acquiring off-target differences between selected cells that may confound the interpretation of subsequent experiments [7], which necessitates validation in multiple clones or programmable strategies in which the same clone can give rise to both experimental and control conditions [8,9].
Recombinant adeno-associated viruses (AAVs) provide superior donor templates for HDR, especially when double-strand breaks (DSBs) are simultaneously introduced at targeted loci [10–12]. Previous studies have demonstrated high HDR rates in hPSCs when Cas9-induced DSBs were combined with AAV donors, reaching ~40%–90% without prior selection [13]. Multiple factors likely contribute to the superiority of AAV virions as vectors of HDR, including its single-stranded DNA genome and the nature of its flanking inverted terminal repeats (ITRs). Several AAV serotypes effectively transduce hPSCs, of which serotype 6 is commonly used [13,14]. AAV-DJ is a synthetic hybrid serotype with expanded tropism that provides comparable rates in hPSCs [13,15]. The use of AAV donor templates may thus overcome challenges related to limited targeting efficiencies. However, contrasting the off-the-shelf availability of Cas9 protein and synthetic sgRNAs, the multi-step cloning and production of AAV virions is time-consuming, which provides a barrier to this approach.
Here, we present a protocol for gene-editing of hPSCs that includes a system for one-step cloning of AAV targeting vectors by Golden Gate [16] assembly as well as a simplified protocol for rapid small-scale isolation of AAV virions. Combined with Cas9 ribonucleoprotein (RNP) complexes, this approach provides a highly efficient yet adaptable approach for gene editing in hPSCs, exemplified by large tag insertions, the generation of conditional alleles, as well as single base edits.
Materials and reagents
Biological materials
AAV-293 cells (Agilent, catalog number: 240073)
hESC H9, WA09 (WiCell, catalog number: WAe009-A)
One shot Stbl3 chemically competent E. coli (Thermo Fisher Scientific, catalog number: C737303)
AAV-GG plasmid vector (Addgene, # 213039)
AAV-packaging plasmids pHelper, pRC-DJ [15]
Reagents
mTeSR Plus (STEMCELL, catalog number: 100-0276)
CloneR2 (STEMCELL, catalog number: 100-0691)
ReLeSR (STEMCELL, catalog number: 100-0483)
P3 Primary Cell 4D-Nucleofector Kit S (20 μL) (Lonza, catalog number: V4XP-3032)
DMEM, high glucose/GlutaMAX/pyruvate (Thermo Fisher Scientific, catalog number: 31966047)
MEM, no glutamine, no phenol red (Thermo Fisher Scientific, catalog number: 51200046)
Fetal bovine serum (FBS) (Merck, catalog number: F7524)
TrypLE Select enzyme, no phenol red (Thermo Fisher Scientific, catalog number: 12563011)
DPBS without calcium and magnesium (Thermo Fisher Scientific, catalog number: 14190250)
Bambanker freezing media (Techtum, catalog number: 23-BB01)
Matrigel hESC-qualified matrix (Corning, catalog number: 354277)
Accutase (STEMCELL, catalog number: 07920)
Thiazovivin (Cayman chemical, catalog number: 14245-5)
PrimeSTAR GXL DNA polymerase (Takara Clontech, catalog number: R050A)
Nuclease-free DEPC water, PCR grade (Thermo Fisher Scientific, catalog number: AM9937)
Alt-R S.p. HiFi Cas9 Nuclease V3, 500 μg (IDT, catalog number: 1081061)
Alt-R CRISPR-Cas9 crRNA, 2 nmol (IDT, custom design)
Alt-R CRISPR-Cas9 tracrRNA, 20 nmol (IDT, catalog number: 1072533)
BpiI FastDigest enzyme (Thermo Fisher Scientific, catalog number: FD1014)
T4 DNA ligase 2000 U/μL (NEB, catalog number: M0202T)
10× T4 DNA ligase buffer (NEB, catalog number: B0202S)
QIAEX II gel extraction kit (Qiagen, catalog number: 20021)
Plasmid Plus Midi Kit (Qiagen, catalog number: 12945)
DNEasy Blood and Tissue kit (50 st) (Qiagen, catalog number: 69504)
CaCl2 (Sigma-Aldrich, catalog number: S9888)
NaCl (Sigma-Aldrich, catalog number: 223506)
KCl (Merck, catalog number: 4936.1000)
Na2HPO4 (Merck, catalog number: 1.06586.0500)
D-(+)-Glucose (Sigma-Aldrich, catalog number: G7021)
HEPES (Sigma-Aldrich, catalog number: H3375)
MgCl2, (Sigma-Aldrich, catalog number: M9272)
Laboratory supplies
6-well culture plates (Thermo Fisher Scientific, catalog number: 140675)
96-well culture plates (Corning, Falcon, catalog number: 353072)
1.5 mL Eppendorf tubes (VWR, catalog number: 89000-028)
15 mL centrifuge tubes (VWR, catalog number: 21008-216)
10 μL racked tips, low-retention sterile (VWR, catalog number: 10017-062)
200 μL racked tips, low-retention sterile (VWR, catalog number: 76322-150)
1000 μL low-retention tips (VWR, catalog number: 10017-090)
Nunc cell culture cryogenic tubes, 1 mL (500) (Thermo Fisher Scientific, catalog number: 377224PK)
PCR tubes and domed caps, strips of 8. (Thermo Fisher Scientific, catalog number: AB0266)
T25 EasYFlask, TC surface, filter cap (Thermo Fisher Scientific, catalog number: 156367T25)
5 mL round-bottom polystyrene tubes (VWR, catalog number: 734-0445)
Amicon Ultra-4 centrifugal filter unit (Merck, catalog number: UFC810024)
35 μm cell strainer cap polystyrene tubes (Falcon, catalog number: 08-771-23)
Equipment
T100 Thermal Cycler (Bio-Rad, model: 1861096)
Microcentrifuge 5418R (Eppendorf, catalog number: 5401000010)
Benchtop refrigerated centrifuge 5430R (Eppendorf, catalog number: 022620601)
4D-Nucleofector X Unit (Lonza, catalog number: AAF-1003X)
NanoDrop 1000 UV/VIS Spectrophotometer (Thermo Fisher Scientific, catalog number: ND-1000)
FACS Aria™ Fusion Flow Cytometer (BD Biosciences, serial number: R656700J40005)
Software and datasets
CHOPCHOP [17] (version 3, http://chopchop.cbu.uib.no)
TIDE [18] (version 3.3.0; http://tide.nki.nl)
Sequence analysis software, e.g., Geneious Prime (version 2024.0.2, BioMatters, 2024-02-14)
Procedure
Design strategy and selection of target gRNA
We design gRNAs for the desired target site aided by the CHOPCHOP design tool (http://chopchop.cbu.uib.no) [17]. Select a gRNA target site that will be disrupted by recombination with the targeting vector, preferably by disrupting its protospacer adjacent motif (PAM).
With this constraint in mind, choose 1–2 gRNAs per site based on their mismatch number and efficiency score from [19], preferably selecting guides with efficiency scores >45 and no off-target sites with <3 mismatches. The gRNAs should ideally cut no more than 30 bp away from the desired recombination site for optimal efficiency [20], and the target sequence should be edited by recombination to prevent recutting.
Design of targeting vector and generation of fragments by PCR or custom synthesis
Design the targeting vector by generating a desired target sequence flanked by ~300–500 bp homology arms (Figure 1A). Each fragment should contain Golden Gate–compatible overhangs listed in Table 1 (ensure that there are no BbsI sites in target sequences). The homology arms can be amplified by PCR with BbsI-compatible primer extensions shown in Table 2, using a high-fidelity polymerase (e.g., PrimeSTAR, Takara). It is preferable to use genomic DNA from the cell line to be targeted as the template for the PCR reaction. Alternatively, the homology arms and/or the edited sequence can be ordered as synthetic fragments (e.g., IDT gBlocks).
Figure 1. Efficient homologous recombination in human pluripotent stem cells (hPSCs) by combining Cas9 ribonucleoproteins (RNPs) and purified adeno-associated virus (AAV)-DJ donor templates. A. One-step assembly of AAV targeting vectors by Golden Gate assembly. The acceptor vector contains unique overhangs flanked by BbsI-sites and interspaced by a “stuffer sequence” to separate the inverted terminal repeats (ITRs). Transgenes, exons, and/or homology arms generated by PCR or as synthetic fragments can be assembled in a single step. The fragments are assembled in the AAV donor vector and used to package the donor template in AAV particles. B. Schematic of the human ACTB locus and strategy to insert 2A-GFP in the 3’-UTR. Target site for the gRNA is marked by a pair of scissors. C. Outline of the experiment (top) to measure homologous recombination efficiencies by FACS analysis. Targeting efficiencies were measured as the percentage of GFP-positive cells following transfection with AAV plasmid or transduction with different amounts of donor templates with ~500 bp or ~300 bp homology arm lengths, respectively (right). Kruskal-Wallis statistical comparison of 500 bp and 300 bp arms targeting for each dilution was not significant (ns, p > 0.999). D. Schematic of the human LMNB1 locus and strategy to insert GFP-2A in the 5’-UTR. Homology arms were of ~350 bp lengths. The target site for the gRNA is marked by a pair of scissors. E. Representative plots (left) and summary statistics on the proportion of GFP-positive cells. Multiplicity of infection (MOI) refers to the ratio of viral genomes (vg), as determined by quantitative PCR, per transduced cell. Data represented as mean ± standard error of the mean (SEM).
Table 1. Example of Golden Gate fragment overhangs used for the modular assembly of AAV vectors (Figure 1A)
Fragment Left junction Right junction
AAV-GG vector AGCG GCAA
Left homology arm (LHA) GCAA AGTG
Edited sequence AGTG TCCA
Right homology arm (RHA) TCCA AGCG
Table 2. PCR primer extensions for amplifying BbsI-compatible fragments.
The underlined sequence corresponds to ligated overhangs (Table 1).
Fragment Forward primer Reverse primer
LHA 5’-CACCACAGAAGACGAGCAA…[primer] 5’-CACCACAGAAGACTCCACT…[primer]
Edited 5’-CACCACAGAAGACGAAGTG…[primer] 5’-CACCACAGAAGACTCTGGA…[primer]
RHA 5’-CACCACAGAAGACGATCCA…[primer] 5’-CACCACAGAAGACTCCGCT…[primer]
Fragments amplified by PCR should be gel-purified, e.g., using the QIAEX II Gel Extraction Kit, resuspended in Milli-Q water. Assess quality and concentration using UV spectroscopy, e.g., using a NanoDrop instrument.
One-step assembly of targeting vector by Golden Gate assembly into universal AAV vector
Set up the Golden Gate assembly reaction in a PCR tube following the recipe in Table 3. Adjust volumes according to the concentrations and lengths of the fragments (total amount of each should be 40 fmol).
Table 3. Typical reaction mix for Golden Gate assembly of three fragments into the AAV-GG destination vector
Fragment Concentration (ng/μL) Volume (μL)
Left homology arm (ca. 500 bp) 50 0.25
Right homology arm (ca. 500 bp) 50 0.25
Edited sequence (ca. 500 bp) 50 0.25
AAV-GG vector (ca. 4,000 bp) 100 1
10X T4 DNA ligase buffer - 1.25
BpiI (isochisomer to BbsI) - 0.75
T4 DNA ligase 2000U/μL - 0.25
Water - 8
Total 12.5
Perform the Golden Gate reaction in a thermal cycler using the following program (Table 4):
Table 4. Golden Gate cycling program
37 °C 1 min 30×
16 °C 1 min
60 °C 5 min
Transform competent bacteria, according to standard procedures. We use chemically competent Stbl3 cells, with 40–50 μL of cells transformed with 1/10 vol of the Golden Gate reaction.
Plate transformed bacterial culture into LB-agar plates with ampicillin and incubate overnight at 37 °C.
Pick 4–6 colonies and grow in LB with ampicillin overnight; expect at least one positive colony. The amount of positive clones obtained differs depending on the purity and size of the fragments but, typically, efficiencies range from 30% to 100%.
Critical: Include a negative control Golden Gate reaction without one of the homology arm fragments to assess for background formed by re-ligation of the backbone plasmid.
Miniprep cultures and test for correct assembly, e.g., using diagnostic restriction enzyme digests.
Amplify a correct clone by growing a 35 mL LB culture and purify the plasmid using the Plasmid Plus Midi Kit, according to instructions.
Rapid isolation of AAV virions from AAV-293 cell conditioned media
Culture AAV-293 cells in DMEM supplemented with 10% (vol/vol) FBS.
One day prior to transfection, split near-confluent cells using TrypLE to a T25 flask (surface area 25 cm2) at 1:3 dilution.
One hour prior to transfection, replace to fresh DMEM-FBS media.
Co-transfect the AAV-293 cells with the AAV targeting vector along with pHelper and RC-DJ plasmids using the Cal-Phos method: Per T25 flask, prepare in a 1.5 mL Eppendorf tube 37 μL of 2 M CaCl2 solution, 3 μg of each plasmid, and water to a total volume of 300 μL. Add this dropwise to a 5 mL round-bottom polystyrene tube containing an equal volume (300 μL) of 2× HBS (0.4 M NaCl, 10 mM KCl, 1.5 mM Na2HPO4, 0.2% glucose, 38.4 mM HEPES pH 7.05) and immediately disperse on top of the cells.
Four hours after transfection, replace media with 5 mL of mTeSR Plus medium.
Collect the media (approximately 5 mL) 72 h post-transfection and clear dead cells and debris by centrifugation at 1,500× g for 10 min at 4 °C.
Equilibrate an Amicon Ultra-4 centrifugal filter (100 kDa) 15 mL spin column with DPBS + 2 mM MgCl2, followed by a brief centrifugation (1 min, until most of the PBS has passed the filter) at 1,500× g at 4 °C.
Load the cleared viral supernatant on the spin filter and centrifuge at 1,500× g for 5–15 min at 4 °C to concentrate the virus solution to ~500 μL, followed by a brief wash with mTeSR Plus medium and further concentration until a final volume of 250 μL of cleaned mTeSR Plus media containing virion particles. Caution: Do not allow the membrane to dry out, as viral titer may decrease due to particles becoming trapped in the filter membrane. Check the volume every 2–3 min during centrifugation to ensure volume reduction without complete drying.Pause point. AAV-DJ particles can be snap-frozen in small aliquots at liquid nitrogen. In contrast to AAV purified over iodixanol gradients, this rapid preparation of AAV particles will contain a significantly higher amount of non-infectious AAV genomes, making the determination of viral titers by quantitative PCR inaccurate. Hence, we empirically determine suitable viral amounts to use (see below).
Nucleofection of hPSCs with CRISPR-RNPs and subsequent transduction of cells with AAV carrying the targeting vector
Culture hPSCs for one week prior to performing nucleofection. Maintain cells in mTeSR Plus medium on Matrigel-coated plastic (0.5 mg/mL dilution in MEM) and keep in a humidified incubator with 5% CO2 at 37 °C. Replace media every other day and passage cells when required using ReLeSR according to the manufacturer’s instructions.
Anneal Alt-R crRNA with Alt-R tracr-RNA (both from IDT) in a 1:1 ratio at 50 μM final concentration of each gRNA by heating to 95 °C for 5 min in a PCR cycler followed by slow cool down to room temperature on the bench top for 3 min; then, place on ice. Annealed guides can be stored at -20 °C for 2–3 weeks and at -80 °C for 6 months.
Prepare Cas9 RNPs by mixing recombinant Alt-R S.p. HiFi Cas9 Nuclease V3 (IDT) with the annealed crRNA and tracr-RNA for a total volume of 4 μL per reaction, as in Table 5. Incubate the mixed gRNA and Cas9 protein at room temperature for 10–15 min before transfection to ensure proper RNP complex formation. Pause point. RNPs can be stored at -20 °C for up to 2 weeks.
Table 5. CRISPR-Cas9 RNP reagents for a single nucleofection reaction
Reagent Final concentration (μM) Volume (μL)
Annealed crRNA and tracr-RNA 50 2,3
Alt-R™ S.p. HiFi Cas9 Nuclease V3 (IDT) 61 1,7
Total 4
Nucleofection of hPSCs:
Treat cells with ROCK inhibitor (we use 2 μM thiazovivin) for ~2 h prior to transfection.
Detach cells with Accutase and count cells using an automated cell counter or Burker chamber. In a 15 mL Falcon tube, spin down (200× g for 5 min) 400,000 cells per transfection. Carefully remove all supernatant and resuspend cells in 20 μL of P3 solution (Lonza). Transfer the cell suspension to an Eppendorf tube and mix with 4 μL of RNP complex. Immediately transfer to a 16-Nucelocuvete strip using a 4D-Nucleofector system (Lonza). Tap strips to avoid the presence of bubbles and bring all material to the bottom of the cuvette.Caution: Work fast and avoid creating bubbles during the resuspension of cells with the P3 solution and transfer to the nucleocuvette.Critical: It is recommended to include transfection with a GFP plasmid (1.5 μg) as a control for nucleofection efficiency. A condition nucleofecting gRNA without Cas9 protein may be included as an additional negative control for downstream applications.
Electroporate in 4D nucleofector (Lonza) using the program CA-137.
Immediately after pulse completion, resuspend cells in 100 μL of mTeSR Plus with ROCK inhibitor (e.g., 2 μM thiazovivin) and plate onto Matrigel-coated 12-well plates previously filled with 1 mL of warm (37 °C) mTeSR Plus medium with ROCK inhibitor. Seed at a density of 200,000 cells per well.
Add AAV particles to the media within 10 min after nucleofection. We normally add 5 μL per well and a no-virus control.Recommended: As viral toxicity can be noticed in some experiments, we recommend empirically testing the optimal amount of viral particles by adding (e.g., 1:5) serial dilutions of the AAV preparation to 3–4 replicate wells of nucleofected cells and subsequently select the condition with the highest titer not showing apparent toxicity.
Place cells back in the incubator and culture for 24–72 h prior to analysis. Change media to mTeSR Plus without ROCK inhibitor 24 h after transfection. Monitor cell viability as high amounts of AAV particles may affect cell viability. Continue with the highest titer that does not show excessive cytotoxicity.
Optional: Isolation of single clones by FACS for expansion and PCR screening.
For isolation and analysis of single clones, we recommend sorting single cells by FACS 24–48 h after Cas9 RNP nucleofection and AAV transduction.
Prepare a 96-well plate coated with Matrigel for 1 h at 37 °C. Fill each well with 100 μL of mTeSR Plus media supplemented with CloneR2 (to promote cell survival).
Detach cells with Accutase, spin down (200× g for 5 min), and resuspend in mTeSR Plus supplemented with CloneR2. Pass cells through a 35 μm cell strainer cap of a polystyrene tube.
Configure gates and flow of an automated cell sorter such as a BD FACSAria III to sort single viable cells into 96-well plates.
Following sorting, place the plate in the incubator. Replace media with mTeSR Plus for clonal expansion 48 h post-sorting and change media every other day. Colonies will appear within one week.
For analysis and banking, expand clones into two duplicate 24-well plates such that one well is used to collect DNA (e.g., using QIAGEN DNAeasy kit) and the other to freeze cells. For freezing, detach cells with Accutase, spin down (200× g for 5 min), and resuspend with 300 μL of Bambanker freezing media.
Validation of protocol
Efficient gene editing in hPSCs by AAV-DJ donors obtained by one-step assembly
A previous study demonstrated high rates of HDR in hPSCs when Cas9-induced DSBs were combined with repair templates delivered by AAVs [13]. However, a drawback of this approach is the requirement to design and generate allele-specific AAV vectors with sometimes complex cloning pipelines. We sought to simplify this workflow by first designing a simplified strategy to assemble donor templates in an AAV backbone by one-step Golden Gate assembly (Figure 1A) [16,21].
To test the utility of our approach, we assembled a targeting vector to insert a sequence encoding a self-cleaving 2A peptide and a green fluorescent protein (P2A-GFP) in the 3’-untranslated region (UTR) of the ACTB locus. GFP expression is thus driven by endogenous ACTB expression in correctly targeted cells only. This approach is analogous to one previously used in mice [22] but employs shorter homology arms of only 300 or 500 bp (Figure 1B) to further simplify vector build. A gRNA sequence (5’-CGTCCACCGCAAATGCTTCT) to cut near the stop codon in exon 6 of ACTB was selected. The AAV targeting vectors were generated by PCR using primer overhangs shown in Table 2. The P2A sequence was added by a long primer, amplifying the GFP fragment of the plasmid pEGFP_N1 (Clontech). Flanking homology arms (300 or 500 bp) were similarly prepared by PCR from synthetic DNA fragments (gBlock; IDT) corresponding to the following genomic coordinates (hg38): LHA300 (chr7:5,527,751-5,528,055); LHA500 (chr7:5,527,751-5,528,304); RHA300 (chr7:5,527,441-5,527,749); RHA500 (chr7:5,527,184-5,527,748); see Supplementary information for full targeting sequence and primers used for cloning. PCR products were gel-purified using the QIAEX II Gel Extraction Kit and subjected to the single-tube reaction assembly into the AAV destination vector (Table 3), using the following program: 30× (37 °C, 5 min, 16 °C, 5 min) 60 °C, 5 min. After completion of the reaction, another 0.5 μL of Bpil was added, followed by a final incubation at 37 °C for 60 min to remove background. Colonies were screened by restriction digests and verified by Sanger sequencing.
Human H9 ES cells were electroporated with the Cas9 RNP and AAV particles, here purified over an iodixanol gradient [23]. Two days later, the proportion of GFP-expressing cells was quantitatively measured by flow cytometry (Figure 1C). We observed a dose-dependent response, with editing rates of up to ~80% editing in conditions using a high titer of AAV-DJ virions (8 × 105 viral genomes per cell). As expected, recombination rates were lower albeit not significantly different when homology arms were shortened to 300 bp, but in the same order of magnitude. The combination of Cas9 RNP and AAV-DJ donor templates thus yields high HDR rates also with relatively short homology arms.
To test the universality of this approach, we took a similar promoterless approach by instead targeting GFP upstream of the coding sequence of LMNB1 (Figure 1D). We similarly designed 300 bp homology arms from synthetic DNA fragments (gBlocks; IDT) corresponding to the genomic coordinates (hg38) LHA (chr5:126,776,986-126,777,467) and RHA (chr5:126,777,597-126,778,005), and assembled with a GFP-2A sequence for one-step assembly in our universal AAV targeting vector, here using a synthetic fragment (IDT gBlock) because a high GC content made genomic PCR challenging (see Supplementary information for full targeting sequences and primers used for cloning). AAV-DJ particles purified by density gradient centrifugation were delivered to human H9 ES cells following electroporation with Cas9 RNPs with a gRNA targeting exon 1 of LMN1B (5’-GGGGTCGCAGTCGCCATGG). The proportion of GFP-positive cells two days later was about 30% (Figure 1E). The lower efficiency compared with the targeting of ACTB may reflect allele-specific differences in targeting efficiencies. The apparent lack of dose-dependent increase with higher AAV concentrations may also reflect toxicity from viral particles, as described in the hematopoietic stem and progenitor cells (HSPCs) [24], and/or allele-specific effects related to manipulation of the LMNB1 locus at the 5’UTR region [22].
Rapid small-scale AAV purification simplifies editing pipeline
In our first experiments, we used dilutions of concentrated AAV particles highly purified through gradient density centrifugation. As this procedure is laborious, here we compared this to simply collecting, briefly washing and concentrating viral particles from the media of AAV-producing cells on spin filter columns (Figure 2A). Using the same assays to test for insertion by HDR as shown in Figure 1, we found high degrees of GFP-positive cells in both ACTB and LMNB1 loci. The rates were almost as high as with the purified vectors, in a dose-dependent manner (Figure 2B-2C). This simplified protocol can reduce the time and cost needed to generate AAV virions for in vitro applications. We have also used this protocol to efficiently introduce a tag in the NRXN1 locus [25].
Figure 2. Rapid small-scale production of adeno-associated virus (AAV)-DJ targeting vectors permits efficient homologous recombination.
A. Schematic comparing conventional purification of AAV particles by density gradient (top) with our simplified protocol for faster, small-scale production for in vitro use (bottom). B. Summary statistics of homology recombination rates in the ACTB locus obtained by small-scale purification of AAV-DJ. Dashed lines indicate the rates obtained with the highest titer of purified viral particles, used in Figure 1C. The 1:1 dilution is estimated to correspond to 8 × 103 viral genomes/cell. C. Summary statistics of homology recombination rates in the LMN1B locus obtained by small-scale purification of AAV-DJ. Dashed lines indicate the rates obtained with the highest titer of purified viral particles, used in Figure 1E. Data represented as mean ± standard error of the mean (SEM).
General notes and troubleshooting
Step Problem Possible reason Suggested solution
Cloning AAV-GG backbone Low yield of bacterial colonies after Golden Gate cloning. Low purity of HDR-template fragments; overhangs incompatibilities due to design inaccuracies; presence of additional BbsI/Bpil recognition sites in fragments; low transformation efficiency. Check design of primers and/or synthetic fragments; ensure purity of fragments by gel electrophoresis and UV spectroscopy; use fresh lots recommended enzymes (Bpil Invitrogen, Cat# FD1014) and high-concentration T4 DNA ligase (NEB, Cat# M0202T) with fresh aliquots of T4 ligase buffer (NEB); include positive and negative controls to test for competency of bacteria and selection on plates.
No bands, multiple bands, or low yield following genomic PCR to generate cloning fragments. Primer or template sequences incompatible with PCR; suboptimal quality of template DNA. Check primers, template GC-content, annealing temperature, and optimize PCR accordingly; use DNA polymerase optimized for challenging templates (e.g., PrimeSTAR GXL, Takara, Cat# R050A); order target sequences as synthetic fragments.
High number of background colonies in the Golden Gate reaction. Incomplete digestion of the AAV-GG backbone plasmid. Perform an extra step of BbsI/Bpil digestion after the Golden Gate reaction to linearize unligated backbone vector and reduce the number of background colonies.
hPSC transfection Insufficient cell survival after transfection and AAV infection. Cell confluency before transfection < 60%; Nucleofection-related toxicity; AAV-related toxicity; gene-editing affects cell survival. Ensure a cell density of at least 80% before transfection and perform gentle pipetting when detaching cells with Accutase. Work swiftly after resuspension of cells with the P3 transfection solution to avoid toxicity-derived cell death and add ROCK inhibitor to media before and after nucleofection; dilute AAV particles even further; consider the potential cell-lethality of the intended gene edit.
Low-efficiency transfection observed as low yield of GFP positive upon nucleofection with control plasmid. Suboptimal cell health prior to nucleofection; suboptimal handling of cells during nucleofection; expired P3 transfection solution. See above; also ensure the quality of reagents including the GFP plasmid and that the P3 solution is not expired.
AAV generation Low efficiency of gene-editing events. Low yield of AAV donor templates; low-efficiency sgRNA targeting or sgRNA degradation; low nucleofection rates; inefficient gRNA. Use AAV-293T cells that produce higher AAV titers than HEK293 cells. Ensure high density (around 70% confluency) of HEK cell culture before transfection; compare targeting efficiencies with or without Cas9 RNP or AAV donors and assess efficiency by a functional readout (e.g., GFP fluorescence) or analysis of bulk DNA (e.g., T7 assay); try alternative gRNAs.
Acknowledgments
We thank Susann Li and Stefanie Fruhwürth for assistance with FACS analysis and Swaroop Achuta for the initial cloning of the AUTS2 targeting vector. Funding: S.M. is supported by a grant from the Assar Gabrielsson Foundation. F.H.S. is supported by a grant from the Swedish Research Council (dnr 2017-03331). F.H.S. and ES are supported by the Knut and Alice Wallenberg Foundation via the Wallenberg Centre for Molecular and Translational Medicine at the University of Gothenburg, Sweden.
Competing interests
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
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Article Information
Publication history
Received: Jun 29, 2024
Accepted: Sep 6, 2024
Available online: Sep 25, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Stem Cell > Pluripotent stem cell > Cell-based analysis
Molecular Biology > DNA > Gene expression
Biological Sciences > Biological techniques > CRISPR/Cas9
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A Real-Time Approach for Assessing Rodent Engagement in a Nose-Poking Go/No-Go Behavioral Task Using ArUco Markers
TS Thomas J. Smith
TS Trevor R. Smith
FF Fareeha Faruk
MB Mihai Bendea
SK Shreya Tirumala Kumara
JC Jeffrey R. Capadona
AH Ana G. Hernandez-Reynoso
JP Joseph J. Pancrazio
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5098 Views: 289
Reviewed by: Edgar Soria-GomezNoah Cowan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eNeuro Mar 2024
Abstract
Behavioral neuroscience requires precise and unbiased methods for animal behavior assessment to elucidate complex brain–behavior interactions. Traditional manual scoring methods are often labor-intensive and can be prone to error, necessitating advances in automated techniques. Recent innovations in computer vision have led to both marker- and markerless-based tracking systems. In this protocol, we outline the procedures required for utilizing Augmented Reality University of Cordoba (ArUco) markers, a marker-based tracking approach, to automate the assessment and scoring of rodent engagement during an established intracortical microstimulation-based nose-poking go/no-go task. In short, this protocol involves detailed instructions for building a suitable behavioral chamber, installing and configuring all required software packages, constructing and attaching an ArUco marker pattern to a rat, running the behavioral software to track marker positions, and analyzing the engagement data for determining optimal task durations. These methods provide a robust framework for real-time behavioral analysis without the need for extensive training data or high-end computational resources. The main advantages of this protocol include its computational efficiency, ease of implementation, and adaptability to various experimental setups, making it an accessible tool for laboratories with diverse resources. Overall, this approach streamlines the process of behavioral scoring, enhancing both the scalability and reproducibility of behavioral neuroscience research. All resources, including software, 3D models, and example data, are freely available at https://github.com/tomcatsmith19/ArucoDetection.
Key features
• The ArUco marker mounting hardware is lightweight, compact, and detachable for minimizing interference with natural animal behavior.
• Requires minimal computational resources and commercially available equipment, ensuring ease of use for diverse laboratory settings.
• Instructions for extracting necessary code are included to enhance accessibility within custom environments.
• Developed for real-time assessment and scoring of rodent engagement across a diverse array of pre-loaded behavioral tasks; instructions for adding custom tasks are included.
• Engagement analysis allows for the quantification of optimal task durations for consistent behavioral data collection without confirmation biases.
Keywords: ArUco markers Automated scoring Computer vision Engagement analysis Go/no-go Real-time tracking
Background
Behavioral neuroscience relies on an accurate assessment of animal behavior to understand complex brain–behavior relationships. Traditionally, such assessments have been performed using labor-intensive manual scoring methods, which are time-consuming, subject to human error, and can become impractical for large-scale studies [1,2]. Recent advancements in computer vision and tracking technologies have led to the development of automated behavioral scoring systems, offering more efficient and objective ways to analyze animal behavior [3–7]. Of these technologies, markerless-based approaches, which leverage machine-learning algorithms for position estimation, have been favored for their ability to capture complex animal movements by tracking an object’s inherent features instead of an attached marker [5,8]. However, they often require high-end computational equipment and extensive training data for robust performance [3,6]. For instance, DeepLabCut, a leading markerless system, recommends a powerful GPU for training models, as relying solely on the CPU can significantly slow execution [3]. Alternatively, marker-based approaches utilize physical markers that can be attached to animals directly, tracking only the markers themselves as a robust estimation of an object’s movement and orientation [4,7,9]. Among these marker-based tracking systems, Augmented Reality University of Cordoba (ArUco) markers have emerged as a promising tool for automated behavioral scoring procedures [10]. ArUco markers consist of black-and-white square grid patterns of various sizes, serving as fiducial target markers that are easily detected and tracked by cameras. Unlike markerless systems, ArUco markers do not rely on machine learning or extensive training data to be tracked, making them computationally lightweight and feasible for real-time analyses [10]. This simplicity enables ArUco markers to be tracked with minimal computational resources, such as low-end CPUs, ensuring accessibility and scalability for a wide range of laboratories. However, one limitation of this approach is the selectivity of the tracked animal behavior. In contrast to a markerless system’s ability to track quick complex movements with predictions between unmarked frames, the ArUco approach is best when utilized for simple movement analyses like locomotion due to camera capture motion blur [10–12]. Nevertheless, ArUco markers offer flexibility in experimental setups where researchers can adapt their tracking to suit their specific experimental needs and study a wide range of behaviors in diverse animal models. Although simple by design, detailed protocols enabling researchers with different backgrounds to readily employ these methods for animal tracking are lacking. Here, we present detailed protocols for establishing a simple and consistent method for automated behavioral scoring of task engagement in an intracortical microstimulation-based nose-poke, go/no-go task using ArUco markers. Our step-by-step approach provides clear instructions, allowing researchers to implement ArUco marker-based tracking systems effectively, thereby advancing the accessibility and reproducibility of behavioral neuroscience research.
Materials and reagents
Biological materials
Male Sprague-Dawley rats (Charles River Laboratories Inc., catalog number: 001CD)
Laboratory supplies
¼ in. diameter vinyl tubing (the length required depends on the setup)
¼ in. diameter, 20 threads/inch, 1 in. long bolt with corresponding nut and (8×) washers
3 mm diameter heat shrink tubing (NTE Electronics, catalog number: HS-ASST-5)
¾ in. binder clip (DUIJINYU, catalog number: W003)
4 in. cable ties (Grand Rapids Industrial Products, catalog number: 54157)
in. binder clip (OWLKELA, catalog number: Clips-Black-0.6inch-120pcs)
6 in. cable ties (Utilitech, catalog number: SGY-CT33)
(2×) 4 ft. Bayonet Neill–Concelman (BNC) cables
6.4 mm diameter heat shrink tubing (Gardner Bender, catalog number: HST-101B)
Acoustic foam wedges (Ultimate Support, catalog number: UA-KIT-SBI)
Break away male header pins, straight (Leo Sales Ltd., catalog number: LS-00004)
Clear packing tape (Duck Brand, catalog number: 287206)
ClearWeld quick-set epoxy (J-B Weld, catalog number: 50112)
Double-sided foam tape
Dustless sugar precision reward pellets (Bio-Serv, catalog number: F0021)
Electrical tape
Hot glue gun with thermal-plastic adhesive (hot glue) sticks
M3 10 mm machine screws with corresponding nuts (6×) (K Kwokker, catalog number: TLEEP2324+FBADE-28)
M3 20 mm machine screws with corresponding nuts (12×) (K Kwokker, catalog number: TLEEP2324+FBADE-28)
M3 30 mm machine screw with corresponding nut (Everbilt, model: 837841)
Medical-grade air tank with regulator (Cuevas Distribution Inc., catalog number: AI USPxxx)
Polylactic acid (PLA) fused deposition modeling (FDM) 3D printer filament
Rubber bands (Advantage, catalog number: 2632A)
Solder wire (Shenzhen Joycefook Technology Co. Ltd, catalog number: JF850)
Stainless steel spring cable shielding (Protech International Inc., catalog number: 6Y000123101F)
Super glue (Loctite, catalog number: 234796-6)
Supplemental food pellets (LabDiet, catalog number: 5LL2, Prolab RMH 1800)
TIP120 negative-positive-negative (NPN) transistors (3×) (BOJACK, catalog number: BJ-TIP-120)
White stock printer paper
Equipment
12V 1A DC power supplies (2×) (Mo-gu, catalog number: B09TXJ9RK6)
150 mm T-Slot 2020 aluminum extrusion rails (4×) (MECCANIXITY, catalog number: mea231123ee000975)
200 mm T-Slot 2020 aluminum extrusion rails (x8) (Tsnamay, catalog number: TSCP5035)
3D-printed ArUco marker mounting assembly hardware (see Procedure)
3D-printed camera mounts (see Procedure)
3D-printed RGB LED strip mounting kit (see Procedure)
3D-printed T-Slot rail connector pieces (see Procedure)
4-pin 12V RGB LED strip
5 mm wide ~4.5 in. long hex wrench
57 in. × 42 in. × 56 in. sound-attenuating chamber (Maze Engineers, catalog number: 5831)
5V activation relay module (Inland, catalog number: 509687)
ArUco tracking cameras (2×) (Logitech, catalog number: 960-001105)
ATmega328 UNO R3 microcontroller (Inland, model: UNO R3 BOARD)
Behavior camera (j5create, catalog number: JVCU100)
Cable tensioner bar (see Procedure)
Commutator (Moog Inc., catalog number: AC6023-18)
Disposable lighter
Electrical Stimulator (Plexon Inc., model: PlexStim)
Fused deposition modeling (FDM) 3D printer
Heat gun (Wagner, catalog number: FBA_503008-cr)
Luxmeter (Leaton, catalog number: HRD-PN-79081807)
Multimeter (Innova, model: 3320)
Omnetics tether adapter (Omnetics Connector Corporation, catalog number: A79021-001)
Operant conditioning chamber (Vulintus, model: OmniTrak)
Oscilloscope (Tektronix Inc., model: TBS 1052B)
Pneumatic solenoid (AOMAG, catalog number: SKUSKD1384729)
Soldering iron (Guangzhou Yihua Electronic Equipment Co. Ltd., model: YIHUA 926 III)
Stackable headers, female 10 pin (2×) (Leo Sales Ltd., catalog number: LS-00009)
Workstation computer with Windows 10–11 (Dell, model: precision 5860 workstation)
Software and datasets
All data and code have been deposited to GitHub: https://github.com/tomcatsmith19/ArucoDetection
Cura v4.13.0 (Ultimaker)
Microsoft Excel
MATLAB vR2023b (MathWorks; requires a student license or higher)
Add-On: MATLAB Support Package for Arduino Hardware v23.1.0 (MathWorks)
Add-On: MATLAB Support Package for USB Webcams v23.1.0 (MathWorks)
Add-On: Instrument Control Toolbox v4.8 (MathWorks)
PlexStim v2.3 (Plexon Inc.)
Prism v10.0.2 (GraphPad; requires a student license or higher)
Python v3.10.11 (Python Software Foundation)
TekVisa v4.0.4 (Tektronix)
Visual Studio Code v1.85 (Microsoft)
Extension: Pylance v2024.6.1 (Microsoft)
Extension: Python v2024.8.0 (Microsoft)
Extension: Python Debugger v2024.6.0 (Microsoft)
Package: numpy v1.25.0
Package: opencv-contrib-python v4.7.0.72 (Open Source Vision Foundation)
Package: scipy v1.13.0 (SciPy)
Microsoft Word
Procedure
Constructing the behavioral chamber
Preparing the sound-attenuating chamber
Adhere acoustic foam wedges around the inside walls of the sound-attenuating chamber (excluding the floor) using super glue or double-sided foam tape. See Figure 1A for an example of the intended outcome.
Figure 1. Behavioral chamber setup. A) Photographic image of the behavioral chamber configured inside the sound-attenuating chamber. B) Cartoon illustration of the behavioral chamber with numbered components: 1) nose-poke module, 2) reward pellet module, 3) OmniTrak controller board, 4) sugar pellet dispenser, 5) T-Slot 2020 rail frame, 6) mounted behavior camera, 7) mounted ArUco marker tracking cameras, 8) cable tensioner bar, 9) commutator, 10) tether, 11) oscilloscope, 12) electrical stimulator, 13) BNC cables, 14) stimulation cable, 15) microcontroller, 16) RGB LED strip, 17) medical-grade air tank and regulator, and 18) pneumatic solenoid.
Setting up the OmniTrak operant conditioning chamber
Place the assembled OmniTrak operant conditioning chamber with nose-poke (Figure 1B.1) and pellet dispenser (Figure 1B.2) modules inside of the sound-attenuating chamber.
Caution: Read the installation instructions provided by Vulintus for the OmniTrak operant conditioning chamber first, then proceed with the protocol instructions listed below.
Mount the matching OmniTrak controller board (Figure 1B.3) to the outside of the sound-attenuating chamber’s right-side panel with double-sided foam tape.
Connect one of the provided RJ45 (ethernet) cables from the OmniTrak nose-poke module to the controller board by feeding it through the upper-right corner hole of the sound-attenuating chamber.
Place the OmniTrak reward pellet dispenser (Figure 1B.4) on the top right edge of the sound-attenuating chamber and connect it to the OmniTrak reward pellet module using the provided vinyl tubing, feeding it through the same hole.
Insert the remaining RJ45 and provided 4-pin power connector cable into both the reward pellet dispenser and the controller board as instructed by Vulintus.
Fill the dispenser reservoir halfway with dustless sugar precision reward pellets.
Connect the OmniTrak controller board to the intended computer using the provided USB cable.
Power on the OmniTrak system by inserting the controller board power cable into a nearby wall socket.
3D printing behavioral chamber components
Visit the highlighted GitHub page for this protocol (https://github.com/tomcatsmith19/ArucoDetection) and download each of the .stl files listed within the 3D Models folder.
Before printing, slice each of the .stl files in Ultimaker’s Cura software to transform them into usable g-code files that are specific to your 3D printer.
Note: In Cura, a layer height of 0.2 mm and cubic pattern infill density of 20% were set for each print, although many setting combinations would work.
Using an FDM 3D printer with PLA filament loaded, print the “RGB LED strip mounting kits” (1×), the “T-Slot rail connector pieces” (8×), and the “camera mount” (3×) files, which include modified models from username: danid87 on Thingiverse (https://www.thingiverse.com/thing:2600507).
Note: Any third-party slicer and FDM 3D printer will work without needing to modify the .stl files as long as the printer has an allocated print volume of at least 220 mm × 220 mm × 250 mm.
Assembling the T-Slot 2020 rail frame
Connect the T-Slot 2020 aluminum extrusion rails with the newly printed T-Slot rail connectors to form an open cube frame. Use 200 mm rails for the top and bottom planes, and 150 mm rails for the vertical sides.
Note: A soft rubber mallet may be used to insert the T-Slot rail connectors completely if they get stuck.
Fix the T-Slot rail frame on top of the operant conditioning chamber (Figure 1B.5) using glue, velcro, double-sided foam tape, cable ties, or a combination of options depending on how permanent the user would prefer the setup to be.
Installing cameras to the T-Slot rails
Assemble three of the 3D-printed camera mounts by referencing the steps outlined in Figure 2.
Mount the behavior camera to the top T-Slot rail closest to the chamber doors (Figure 1B.6) and the two ArUco tracking cameras to the top left and right rails (Figure 1B.7).
Plug in each of the camera’s USB cables to the computer through the hole in the sound-attenuating chamber.
Note: If the cables are too short to reach the computer, USB 3.0 extenders may be utilized.
Figure 2. Instructions for assembling the camera mount. A) Gather the 3D-printed camera mount pieces, M3 10 mm machine screws (2×), M3 20 mm machine screws (4×), M3 30 mm machine screw (1×), M3 nuts (7×), ¼ in. diameter 20 threads/inch 1 in. long bolt (1×) with corresponding nut (1×) and washers (8×), and camera (1×). B) Connect the T-Slot mounting piece to the first segment of the articulating arm with the M3 30 mm screw and nut. C) Connect the remaining articulating arm pieces with the M3 20 mm screws and nuts. D) Thread the 1 in. bolt through the last articulating arm segment with a nut at the bolt’s base and eight washers filling the gap. E) Screw in the camera’s base to the threaded end of the exposed 1 in. bolt. F) Place two of the M3 10 mm screws through the holes of the plastic T-Slot rail mounting piece. G) Fit an M3 nut into each of the two small T-Slot rail adapter pieces. H) Slide the assembled T-Slot rail adapter pieces into the groves of the T-Slot rail (left) and rotate them 90° to lock them into place (right). I) Align the M3 10 mm screws within the T-Slot rail mounting piece with the T-Slot rail adapter pieces and screw them in, mounting the camera arm to the rail.
Creating and attaching the tensioner bar, commutator, and tether
First, modify the commutator by cutting the exposed wires at the top and bottom of the device; the top group of wires should remain 8 in. long and the bottom group should remain 3 in. long. See Figure 3A for an example of the final product.
Assign pin numbers (1–16) to each of the colored wires at the top with matching colors and numbers at the bottom end, leaving two extra wire colors not accounted for; these will become our ground and reference wires.
Solder the top group of wires to two rows of eight male break away pins, arranging the top row pins as 15, 13, 11, 9, 7, 5, 3, and 1 from left to right when looking directly into the pins. The bottom row pins should be arranged as 16, 14, 12, 10, 8, 6, 4, and 2 from left to right when looking directly into the pins (see Figure 3A).
Critical: When soldering any of the wires, cover each of them with a piece of 3 mm heat shrink tube that is just long enough to cover the exposed joints. This will help prevent any of the connections from becoming shorted together.
Now on the bottom end of the commutator, solder all 18 wires to two rows of ten female stackable header pins. The top row pins should be X (blank), 1, 3, 5, 7, 9, 11, 13, 15, and X from left to right when looking directly into the pins. The bottom row pins should be G (ground), 2, 4, 6, 8, 10, 12, 14, 16, and R (reference) from left to right when looking directly into the pins (see Figure 3A).
Wrap the top and bottom ends of the commutator in electrical tape to secure it (see Figure 3A).
Figure 3. Modified commutator and tether cord. A) Image depicting the final product of the commutator modifications with labeled top and bottom end pinouts when looking directly into the pins. B) Image depicting the final product of the tether with labeled top and bottom end pinouts when looking directly into the pins. C) Side-view image of the cable tensioner bar complete with a ¾ in. binder clip, rubber bands, a hex wrench, and the commutator from panel B.
Finally, strip 0.5 cm from the ground wire at the top end of the commutator and leave the reference wire unmodified and unattached.
Moving onto the tether, begin by cutting the Omnetics tether adapter to be 11.5 in. long. Take note that each of the colored wires corresponds to a specific gold pin on the Omnetics adapter end. See Figure 3B for an example of the final product or Video 1 for a demonstration of how to fabricate the tether.
Critical: There are eight gold pins on the top row of the connector with two standoff pins on each side and ten gold pins on the bottom row with one standoff pin on each side. When the gold pins are facing toward you, the top row’s pins are X, X, 1, 2, 3, 4, 5, 6, 7, 8, X, and X from left to right. The bottom row’s pins are X, R, 9, 10, 11, 12, 13, 14, 15, 16, G, and X (see Figure 3B).
Video 1. Fabrication of the tether cord. Instructional video that outlines the steps required to fabricate the exact tether design used within this protocol.
Cut eleven 0.5 in. pieces of the 3 mm heat shrink tube and slide them over all of the Omnetics tether adapter wires, using a heat gun to heat them into place with ¼ in. gaps between each piece.
Cut a 10.5 in. piece of stainless-steel spring cable shielding and slide it over the Omnetics tether cord.
Using the disposable lighter, burn off the insulating layer of each wire at the ends, exposing ~3 mm of bare wire.
Solder the wires to two rows of ten male breakaway pins. The top row wires should match the female end of the commutator when plugged in: X, 15, 13, 11, 9, 7, 5, 3, 1, and X from left to right when looking directly into the pins. The bottom row wires should be as follows: R, 16, 14, 12, 10, 8, 6, 4, 2, and G from left to right when looking directly into the pins (see Figure 3B).
Critical: Before moving to the next step, use a multimeter set in continuity mode to ensure that each connection has continuity with only itself and no other pin.
Coat the exposed wires at both ends of the tether with clear quick-set epoxy to cement the wires in place. Let the epoxy harden for at least 1 h.
Cut six ½ in. pieces of 6.4 mm heat shrinking tube, then slide them onto the tether over the Omnetics end. One at a time, place one piece at each end of the tether and heat it in place with a heat gun. Continue this process until all six pieces are secured (three on each side).
Lastly, create the cable tensioner bar (Figure 1B.8). The cable tensioner used within this experiment is produced from a ¾ in. binder clip, rubber bands, 6-in. cable ties, and a single 5 mm hex wrench that is ~4.5 in. long (see Figure 2C).
Start by clipping a ¾ in. binder clip to the center of the top T-slot rail closest to the back of the chamber around the top corner of the rail that faces the middle of the cage.
Secure the clip in place with two 6 in. cable ties wrapped around each side.
Take the long side of the hex wrench and fasten it to the bottom metal loop of the binder clip using additional cable ties.
Double-wrap 3 rubber bands around the hex wrench and the top metal loop of the binder clip.
Finally, cable-tie the short side of the hex wrench to the commutator so that the commutator is suspended directly above the center of the operant conditioning chamber (Figure 1B.9).
Insert the soldered 20-pin end of the tether into the bottom of the commutator (Figure 1B.10).
Insert the top end of the commutator into the PlexStim stimulation cord with the top pins matching the side of the stimulation cord with the white arrow on it.
Clip the stimulation cord’s metal clamp onto the exposed ground wire at the top of the commutator.
Connecting the oscilloscope
Place the oscilloscope on top of the sound-attenuating chamber (Figure 1B.11).
Insert its provided USB cable into the computer.
Plug in its provided power supply cord into a nearby outlet.
Connecting the electrical stimulator
Place the PlexStim device inside the sound-attenuating chamber to the right of the operant conditioning chamber (Figure 1B.12).
Plug in its provided power supply cord into a nearby outlet through the hole of the sound-attenuating chamber.
Connect the two 4 ft. BNC cables (Figure 1B.13) between the stimulator and oscilloscope’s voltage and current monitor channels through the same hole.
Insert the stimulator’s provided stimulation cable into the top of the commutator (Figure 1B.14).
Connect the stimulator to the computer using the provided USB cable through the hole in the sound-attenuating chamber.
Wiring the ATmega328 microcontroller
Follow the wiring diagram shown in Figure 4 to connect the ATmega328 UNO R3 microcontroller to the RGB LED strip and pneumatic solenoid peripherals.
Critical: To utilize the software without reconfiguration, the red, green, and blue LED wires should be connected to PWM digital pins 9, 6, and 5, respectively, on the microcontroller. The signal, ground, and Vcc pins of the 5V activation relay module should be connected to pins digital 11, ground, and 5V on the microcontroller, respectively. If these pins are not available, they must be reassigned in the ArucoDetection → MATLAB Behavior Program → Required Behavior Functions → LEDColorChange.m and TriggerSolenoidAirPuff.m functions.
Note: ATmega328 UNO R3 microcontroller was selected for its 5V logic, compatibility with MATLAB, (3×) PWM pins, (1×) digital pin, (1×) 5V source pin, and (1×) ground pin. TIP120 NPN transistors were selected for their NPN type, 5V logic, and current support for over 1A. The 5V activation relay module was selected to match the 5V logic of the ATmega328 UNO R3 microcontroller and its ability to support greater than 1A at 12V of power required by the pneumatic solenoid.
Figure 4. Wiring diagram between the microcontroller and peripherals. a) ATmega382 UNO R3 microcontroller. b) 12V 4-pin RGB LED strip with four contact pads labeled from top to bottom: +12V power (+), red LEDs (R), green LEDs (G), and blue LEDs (B). c) TIP120 transistors with three pins labeled from left to right: base (B), collector (C), and emitter (E). d) 12V 1A DC power supply for the RGB LED strip with two wires labeled from top to bottom as ground (GND) and positive (+12V). e) Pneumatic solenoid with two wires labeled from left to right as ground (GND) and positive (+). f) 12V 1A DC power supply for the pneumatic solenoid with two wires labeled from top to bottom as ground (GND) and positive (+12V). g) 5V activation relay module with three pins labeled on the left-hand side of the device from top to bottom as signal (S), ground (G), and Vcc (V); three screw terminals labeled on the right-hand side of the device from top to bottom as normally closed (NC), common (COM), and normally open (NO). h) USB type-B cable for supplying power to the microcontroller alongside data communication with the computer.
Mount the microcontroller to the outside of the sound-attenuating chamber’s right-side panel with double-sided foam tape (Figure 1B.15).
Note: A 3D-printed box can be made to house all the electronics before mounting to the chamber wall.
Insert the microcontroller’s USB type-B cable into the computer.
Mounting the RGB LED strip
Gather the 3D-printed RGB LED strip mounting kit parts and begin by gluing the elongated rectangular strips to the inside of the bracket pieces on the small side (two brackets per rectangle). Reference Figure 1A for further guidance.
With double-sided foam tape, adhere the large end of the bracket pieces and rectangular strips to the outside of the left, back, and right side-panel walls of the conditioning chamber (~90% up from the base), keeping the rectangular strips level with the base of the chamber.
From the outside of the sound-attenuating chamber, push the RGB LED strip through the hole in the chamber so that most of the LEDs are inside while the opposite end remains wired to the microcontroller on the outside.
Thread the RGB LED strip through the bracket mounts, gluing the backside to each of the rectangular strips so that the LEDs face toward the center of the operant conditioning chamber (Figure 1B.16).
Plug the 12V 1A power supply that is wired to the RGB LED strip into a nearby power outlet.
Connecting the pneumatic solenoid for air-puffing punishment
Insert one end of the ¼ in. diameter vinyl tubing to the regulator on a medical-grade air tank (Figure 1B.17). The tank should be securely mounted in place outside of the behavioral chamber.
Cut the vinyl tubing ~4 ft. from the opposite end and set aside.
From the cut end that is connected to the air tank, attach it to the “IN” hole on the pneumatic solenoid with brass fittings (Figure 1B.18).
Then, connect one end of the severed vinyl tube piece to the “OUT” hole on the pneumatic solenoid, again with brass fittings.
Push the opposite end of that tube through the hole in the sound-attenuating chamber and hot glue it to the backside of the nose-poke module hole. The end result should contain the opening of the vinyl tube being flush, secured, and centered with the backside opening of the nose-poke module hole.
Note: A piece of cardboard with a hole can be used to secure the vinyl tube while being glued in place; glue the cardboard piece itself to the backside of the module while sealing the space around the hole with additional hot glue.
Installing and configuring software packages
Behavioral task MATLAB code
Visit the provided GitHub page (https://github.com/tomcatsmith19/ArucoDetection), click on the green < > Code button, and download the entire repository as a zip file to a desired location on the computer.
Locate the downloaded zip file within File Explorer, right-click on the zip file, and select Extract All.
From the extracted folder, open the Excel file located at ArucoDetection → MATLAB Behavior Program → ExcelSessionData → AnimalWeightCheck.xlsx.
Within that file, replace the animal names and their corresponding weights in grams that are specific to your study within the designated cells.
Note: For this protocol’s specific behavioral paradigm, each animal’s weight is monitored weekly due to mild food deprivation practices. The MATLAB behavioral program app possesses a button that evaluates if the animal’s daily weight drops below 90% of its weekly recorded weight from the AnimalWeightCheck.xlsx file; if the animal passes the weight check, it may proceed with the experiment. All animals are fed supplemental food pellets after each behavioral session in quantities calculated by the behavioral app.
Save the AnimalWeightCheck.xlsx file and close it.
MATLAB
With a browser, navigate to the following URL on MathWorks’s website (https://www.mathworks.com/products/new_products/release2023b.html), click the Download Now button, and follow their instructions for downloading and installing MATLAB (vR2023b) to your computer.
Critical: Download the specified version of MATLAB; future versions may not be compatible.
Note: You may need admin privileges for your machine to install this software.
After installing MATLAB, open the application and set its active file directory to the ArucoDetection → MATLAB Behavior Program file path that was extracted from the GitHub repository downloaded in procedure steps B1a–b.
Navigate to the Add-Ons drop-down arrow button within the Environment section of the home menu ribbon. Click the drop-down arrow and select Get Add-Ons.
Within the pop-up window, search for and install the following: 1) MATLAB Support Package for Arduino Hardware, 2) MATLAB Support Package for USB Webcams, and 3) Instrument Control Toolbox. See the Software and Dataset section for version numbers.
Note: Version numbers may differ as each is consistently updated.
Close MATLAB and restart the computer.
Open MATLAB again and, within the active directory folder, double-click to open the BehaviorApp.mlapp file; this will open the MATLAB App Designer window with the corresponding behavioral app already open.
Make sure that the Design View button is selected instead of Code View. Then, navigate to the Session Setup tab within the behavioral app (Figure 5).
Figure 5. Navigating through the behavioral app within MATLAB app designer to update experimental parameters within the session setup tab. a) The drop-down box for selecting the current date. b) A drop-down list for selecting the name of the researcher conducting the current experiment. c) A drop-down list for selecting the name/identification of a specific animal. d) A drop-down list for selecting which microelectrode array channels will be simultaneously stimulated while the Stim Type → Layer option is chosen.
Within the Session Setup → Session Parameter Select panel, update the listed values for Date, Username, and Animal Name (Figure 5a–c) to reflect the current date, the name of the researcher(s) who will be running the behavioral task, and the name of the animal(s) reflected within the AnimalWeightCheck.xlsx file that was updated in step B1d–e. To do so, double-click on each of the boxes. For Date, a pop-up calendar will appear; navigate to the desired date and click on it. For Username and Animal Name, click on any item in the list to reveal a positive and negative sign. Use the positive sign to add names and the negative sign to remove names from the list.
Within the Stimulation Parameter Select panel, click into the Layer 1 Channels list (Figure 5d) and add any combination of stimulated channels (1–16) that are specific to the intended experiment by using the same positive and negative buttons described in the previous step.
Note: The remainder of the Layer 2–6 Channel boxes are currently non-functional for this version of the application.
Next, open the Windows Device Manager application from the computer’s start menu. Then, click the drop-down menu arrow for the section labeled as Ports (COM & LPT); make note of the COM port number that is displayed for which the UNO R3 ATmega328 microcontroller is connected.
Note: You can readily test this by noting the device names, unplugging the USB cable attached to the microcontroller, looking for the difference in device names, and then re-plugging the device.
Go back to the MATLAB behavior app window and switch the Design View option to Code View.
Locate line number 507: app.LED = arduino("COM5", "Uno"); and update the COM number to the one found in Device Manager.
Once finished, switch viewing modes back to Design View and save the newly updated version of the behavioral app.
Close the application.
PlexStim for electrical stimulator
While using a web browser, navigate to Plexon Inc.’s website with the following URL: https://plexon.com/products/plexstim-electrical-stimulator-2-system/.
Click the PlexStim V2.3 – Windows 7 or Windows 10 link to download the PlexStim v2.3 installation file.
Open the downloaded StimulatorV2Setup.exe file and follow the on-screen instructions.
Note: You may need admin privileges for your machine to install this software.
Restart the computer.
TekVisa for oscilloscope
While using a web browser, navigate to Tektronix’s website with the following URL: https://www.tek.com/en/support/software/driver/tekvisa-connectivity-software-v404/.
Click the Download File button to download the correct TekVisa v4.0.4 installation file.
Open the downloaded TekVISA_404_066093809.exe file and follow the on-screen instructions.
Note: You may need admin privileges for your machine to install this software.
Restart the computer.
Python
While using a web browser, navigate to Python’s website with the following URL: https://www.python.org/downloads/release/python-31011/.
Scroll down until the Files section is located, then click on the Windows installer (64-bit) link to download the Python v3.10.11 installation executable file.
Open the file and follow the prompted instructions.
Critical: When prompted, be sure to check the box for Add python.exe to PATH. This is a necessary step.
When prompted, select install now.
Note: You will need admin privileges for your machine to install this software.
When finished, click close.
Restart the computer.
Visual Studio Code and ArUco Marker Tracking Code
While using a web browser, navigate to the following URL: https://code.visualstudio.com/.
Click on the Download for Windows button on the homepage to download the current Visual Studio Code application setup file.
Open that file and follow the prompted instructions.
Note: You may need admin privileges for your machine to install this software.
Open the Visual Studio Code application and click on the Extensions button located on the left-hand side of the screen.
Search and install the newest versions of the following extensions: 1) Python, 2) Pylance, and 3) Python Debugger. See the Software and Dataset section for version numbers.
Note: Version numbers may differ as each is consistently updated.
Close the Visual Studio Code application.
Restart the computer.
Re-open the Visual Studio Code application.
Navigate to the top menu ribbon and select Terminal → New Terminal.
Within the newly opened terminal line, type python --version and press the enter button on the keyboard.
Critical: Ensure that the output text states “Python 3.10.11”.
Troubleshooting: If Python is not found, ensure that it was properly installed and added to PATH during the installation instructions. If a different version of Python appears, be sure to install the following packages to Python v3.10.11 and not the other listed version. Additionally, if another Python version appears, be sure to verify that Python v3.10.11 appears as the selected Python version in the bottom right corner of the Visual Studio Code application window.
Next, type the following three commands into the terminal followed by the enter key after each line:
pip install numpy==1.25.0
pip install opencv-python==4.7.0.72
pip install scipy==1.13.0
Troubleshooting: If an error occurs that states that pip is not installed, then install pip (https://pip.pypa.io/en/stable/installation/).
Go to File → Open Folder at the top left of the screen and navigate to and open the already downloaded ArucoDetection folder from steps B1a-b. The corresponding files within that directory should now appear within the Explorer column of the Visual Studio Code interface.
Note: The drop-down arrow may need to be clicked to show each of the folders/files within the directory.
Find the ArUco Tracking Program folder, click its drop-down arrow to open the folder, and select the AnimalDetect.py file; this is the primary file that runs the ArUco tracking code in parallel with MATLAB.
Open MATLAB and type webcamlist into the command window followed by the enter key. This should output an array in text format, showing you the index value of each camera that is connected to the computer.
With those index values, make note of the two ArUco tracking cameras and subtract 1 from each of their index values found in MATLAB. MATLAB begins indexing at a value of 1, whereas Python begins at 0; the subtraction acts as a conversion to confirm the appropriate camera indexes.
Go back to the AnimalDetect.py file and update the number at the end of line 20: cap = cv2.VideoCapture(1) to the converted index value of the right-side ArUco tracking camera. Additionally, update the number at the end of line 23: cap2 = cv2.VideoCapture(3) to the converted index value of the left-side ArUco tracking camera.
Note: If the wrong cameras were chosen, they can always be updated again as needed.
Save the AnimalDetect.py file and close both Visual Studio Code and MATLAB.
ArUco marker mounting hardware
Assembling the hardware
Follow the instructions listed in step A3 to print the following ArUco marker mounting assembly hardware pieces: (1×) ArUco Marker Mount, (2×) C-Clamp, and (1×) I-Bracket.
Locate one in. generic binder clip and carefully remove each of the two metal loops from the clip using tweezers or pliers.
In Windows File Explorer, open the 15x15mmArUcoMarker3.png file from the ArucoDetection → ArUco Setup Code file path.
Note: Additional markers with various sizes can be produced by modifying the generateMarkers.py file at this location within Visual Studio Code.
Open the image into any image processing application like Microsoft Word or Adobe Illustrator and print the marker on white stock printer paper, then cut it out.
Critical: Make sure that the print size is true to 15 mm × 15 mm. Some applications attempt to resize the image when printing, but the image file itself is set to 15 mm × 15 mm.
Optional: Place a piece of clear packing tape over the ArUco marker. This can help with the longevity of the marker, although a newly printed marker can always be re-glued overtop of the existing one.
Caution: Keep a thin white border around the marker when cutting it out of the computer paper to maintain some contrast if the experimental animals used do not have white fur.
Referencing Figure 6 for guidance, assemble the ArUco marker mounting pieces for a fit test.
Troubleshooting: Reprint any pieces that do not fit properly.
Figure 6. Instructions for connecting the ArUco marker mounting assembly. A) Gather each of the seven ArUco marker mounting pieces: (1×) I-Bracket, (2×) in. binder clip wire loops, (2×) C-Clamps, (1×) ArUco marker mount, and (1×) 15 mm × 15 mm paper printed ArUco marker. B) Pinch the binder clip wire loops together to slot them into the C-Clamps. C) With your index finger and thumb, slide the C-Clamp down the binder clip wire loop until it almost reaches the protruding prongs at the end. Then, insert the prongs into the holes of the I-Bracket. D) Repeat the process described in panel C for the opposite side of the I-Bracket. E) Orient the ArUco marker mount and paper-printed marker as depicted in the image. The tip of the ArUco marker mount should slant down into the workspace surface. F) Place glue on top of the ArUco marker mount 15 mm × 15 mm square surface. G) Adhere the paper-printed marker to the ArUco marker mount. H) Place a 4 in. cable tie into the side hole of one of the C-Clamps. I) Wrap the cable tie around the assembly, threading it through the second C-Clamp hole. J) Slide the cable tie through the marker mount so that the marker is facing away from the assembly. K) Close the cable tie so that it is tightly bound around the assembly. L) Cut the excess length of the cable tie.
Securing the I-Bracket to the animal
While the animal is anesthetized, permanently glue the I-Bracket around the exposed microelectrode array 16-channel Omnetics connector port within the animal’s dental cement head cap (Figure 7). This can be achieved by applying sparing amounts of super glue around each side of the Omnetics connector before slotting on the I-Bracket.
Note: The rectangular center hole and height of the I-Bracket were specifically designed to surround our microelectrode array’s connector port for accessible attachment of the tether to the animal. At the time of surgery, the dental cement head cap was molded to expose enough of the Omnetics connector to fit the I-Bracket at a later post-recovery date. Custom I-Brackets may need to be produced to fit differing devices.
Troubleshooting: If the I-Bracket does not sit flush with the surface of the Omnetics connector, you can gently file down the PLA plastic using a small nail file.
Figure 7. Permanently attaching the I-Bracket to a rat’s head cap. A) An example of a rat that possesses an exposed Omnetics connector port without an I-Bracket attached. B) An example of a rat that has the I-Bracket super glued to the Omnetics connector port and dental cement head cap.
Mounting and dismounting procedures
Follow the instructions outlined in Figure 8 for securely mounting and dismounting the ArUco marker hardware to an awake animal when running the behavioral task.
Note: Steps should almost mirror that of Figure 6.
Once the animal is properly tethered, close the operant conditioning door.
For dismounting, carefully cut the cable tie between the C-Clamps with small scissors, then reverse the steps described in Figure 8.
Figure 8. Instructions for mounting the ArUco marker assembly for behavior. A) With your index finger and thumb, slide the C-Clamp down the in. binder clip wire loop until it almost reaches the protruding prongs at the end. B) While holding the animal still with your non-dominant hand, insert the prongs into the holes of the I-Bracket with your dominant hand and let go. C) Repeat the process described in panels A–B for the opposite side of the I-Bracket. D) Thread the 4 in. cable tie through the side holes of the C-Clamps, keeping the open end of the cable tie facing the animal’s neck. E) Slide the cable tie through the marker mount so that the marker is facing away from the assembly and toward the center of the animal’s neck. Then, barely close the cable tie so that it is loosely bound around the assembly. F) Hold the animal within the operant conditioning chamber and align the tether prongs with the microelectrode array Omnetics port. G) Plug the tether into the Omnetics port. H) Tighten the cable tie so that it is tightly bound around the assembly, securing the position of the marker. The marker should remain aligned with the anterior to the posterior midline of the animal’s neck as close as possible. I) Cut the excess length of the cable tie and release the animal.
Running the behavioral task and ArUco tracking code
Opening the code
Without connecting the animal yet, power on the electrical stimulator and the oscilloscope.
Caution: Make sure to always turn on the stimulator before plugging in the animal to the tether; turning on the device briefly passes current transients that can be seen on the oscilloscope.
Open the medical-grade air tank + regulator valves to allow a small amount of low-pressure air to build at the pneumatic solenoid.
Open the BehaviorApp.mlapp file from MATLAB as outlined in step B2f as well as the AnimalDetect.py file from Visual Studio Code as outlined in step B6m.
In the App Designer window of MATLAB, click the green Run button located at the top of the screen. If you cannot see it, it is located within the Editor tab of the main header menu.
Wait for the pop-up Vulintus code to finish loading and disappear; the behavior app should now be visible.
Navigate back to Visual Studio Code and click the Run Python File play button that is located on the top right of the screen. The Python code will now attempt to connect to MATLAB and open individual windows for the ArUco marker tracking cameras.
Troubleshooting: If an error occurs, please check that your cameras are functioning properly with the computer and that your webcamlist indexes from steps B6n–p are correct. If the error persists, wait a minute or two for MATLAB to finish initializing the behavior app, then click the Run Python File play button again. If Python still refuses to connect to MATLAB and says permission is denied, you may need to contact your administrator for internal host process port access.
Calibrating the cameras
Check to make sure that the cameras are properly oriented so that the blue rectangles shown in each window cover half of the operant conditioning chamber, including the nose-poke and pellet dispenser modules. See Figure 9 for an example of proper camera alignment.
Figure 9. Examples of proper camera alignment for behavioral experiments. a) ArUco tracking camera #1 is mounted on the right side of the operant conditioning chamber and is aligned so that the blue rectangle passes through the midline of the chamber floor. The chamber modules should be located on the right side of the window. b) ArUco tracking camera #2 is mounted on the left side of the operant conditioning chamber and is aligned so that the blue rectangle passes through the midline of the chamber floor. The chamber modules should be located on the left side of the window. c) The behavior camera is mounted on the front side of the operant conditioning chamber. The chamber modules should be located at the top of the window.
Place the ArUco marker mounting piece inside the operant conditioning chamber in different positions to test whether the marker is being properly tracked by the Python code. You should see a colored border around the marker and a red-degree angle listed within one or both tracking camera windows.
Troubleshooting: If the marker is not being tracked well, try the following: 1) clean the camera lenses and the operant conditioning chamber walls, 2) readjust the angle and height of the cameras so that their focus points are calibrated to the approximate rodent head height where the marker will likely be, 3) test the overall brightness of the chamber using a digital luxmeter (see Equipment section). The lux value within the chamber should read at least 3 lux [11], but the average is closer to ~25+ lux. If the value is too low, replace your RGB LED strip with a brighter one.
Testing the behavior code
Once the applications are properly initialized and tracking the ArUco marker, begin selecting custom behavioral task and stimulation parameter settings within the Session Setup panel of the behavior app. Reference [13] regarding information behind the different behavioral task name selections.
Note: For testing purposes, we suggest setting the following parameters: 1) Task Name → DetectAll, 2) Stim Type → Single, 3) Monitor Ch → 1, and 4) Initial Charge (Q) Value → 1 nC/Phase.
Click back to the Session Controls tab within the app and click the Start button.
Using your finger, begin interacting with the nose-poke module within the operant conditioning chamber, testing to see whether or not 1) a pellet is dispensed when poking after a stimulation trial, 2) an air puff is triggered when poking after a silent trial, and 3) that the RGB LED strip changes colors between white, green, and red depending on the trial state.
Next, watch to see if a stimulus pattern is presented on the oscilloscope preceding a stimulation trial.
Note: Since the animal is not plugged in, you may see a non-typical voltage transient pattern. Additional stimulation pattern parameters can be adjusted in the code under the Required Behavior Functions folder → ICMSandCapture.m file.
Test to see if the Feed and Pause buttons work on the Session Controls panel of the behavior app.
Finally, adjust the medical-grade air regulator to output the desired air pressure when an air puff is triggered.
Troubleshooting: If any of the listed operations do not occur, please double-check your equipment setup.
After testing has been completed, click the Stop button within the Session Controls panel of the behavior app. This will properly close both the MATLAB and Python applications.
Running experimental sessions with the behavior code
Once the chamber has been fully tested and working as intended, you may connect the animal subject to the cage using the mounting/dismounting instructions outlined in step C3 and then repeat the steps listed above.
Note: Specific behavioral task parameters can be modified by navigating to their corresponding tab within the behavior app. For the Stim Type option on the Session Setup panel, N/A = no stimulus presented to the animal, Single = only the channel specified by Monitor Ch., Layer = simultaneously stimulating the channels listed within the Layer 1 Channels drop-down selector, and All = simultaneously stimulating all 16 channels of the microelectrode array.
Locating post-session results
Behavioral performance data
After a behavioral session has concluded and ended using the Stop button, an Excel file containing the raw behavioral data for each individual trial is saved into the ArucoDetection → MATLAB Behavior Program → ExcelSessionData → SessionDataSheets folder. Locate this folder and open the specific Excel session file just produced.
The Excel file contains MATLAB variable data that was automatically generated during the behavioral session using the nose-poke sensor module, native MATLAB clock and array indexing functions, and the ArUco tracking program, all without the need for manual data recording. To find a particular data type of interest, navigate through each of the different sheets within the Excel file: Sheet 1 includes a summary of the general session details, Sheet 2 includes parameter details about each of the individual trials that were presented, and Sheet 3 includes arrays for reproducing the Session Timepoint Data plot that was presented and updated in the behavior app after every trial. See Tables 1–3 for examples of each sheet.
Note: During each behavioral session, a 0.15-s delay follows the presentation of any stimulus to prevent the animal from coincidentally nose-poking during stimulus delivery. The reaction times recorded in Table 2 exclude this delay, allowing for flexibility if the user adjusts the delay duration. For accurate post-session analysis based on reaction time, the delay duration should be added to the final values.
Table 1. Exported behavioral session data found in Sheet 1. This table presents an example of raw data generated by the BehaviorApp.mlapp program in MATLAB that was exported to an Excel file. It includes general session summary values used for assessing behavioral performance, with all trials included. This data provides a comprehensive overview of session metrics, allowing for detailed analysis without pre-filtering or exclusion of trials.
Sheet 1
Task Name Go-No-Go
Researcher Name Thomas
Session Number 17
Session Date 11-Jan-23
Animal Name NS008
Animal Weight (g) 494
Stim Type All
Session Pellets 128
Manual Pellets 0
Total Pellets 128
Accuracy Score (%) 93.66515837
Stimulus Trials 111
Silent Trials 110
Hits on Stim Trials 98
Misses on Stim Trials 10
Late Hits on Stim Trials 3
Hits on Silent Trials 1
Misses on Silent Trials 109
Catch Trials 281
Hits on Catch Trials 0
Total Timeout Time (s) 72
Initial Session Time (s) 63299.29
Ending Session Time (s) 66900.93
Total Pause Time (s) 0
Table 2. Shortened example of the exported behavioral trial-specific data found in Sheet 2. This table displays data from 15 sequential trials within an example behavioral session that was generated in MATLAB and exported to Excel using the BehaviorApp.mlapp program. It includes the raw values for each individual trial, providing a detailed record of the animal's responses and the corresponding session metrics. This trial-specific data is essential for analyzing the progression and consistency of behavior throughout the session.
Table 3. Shortened example of the exported behavioral plot data found in Sheet 3. This table presents data from 15 sequential trials (as shown in Table 2) within an example behavioral session that was generated in MATLAB and exported to Excel via the BehaviorApp.mlapp program. It includes raw plot values for each trial type—stimulation, silent, and alternative stimulation—along with their corresponding X and Y-axis values, listed in order of presentation. The session comprised eight stimulation trials, five silent trials, and two alternative stimulation trials.
Sheet 3
Stim X-Axis (min) Stim Y-Axis (s) Sil X-Axis (min) Sil Y-Axis (s) AltStim X-Axis (min) AltStim Y-Axis (s)
0.138583333 0.016 0.397966667 0 21.3667 0.022
0.540683333 0.018 1.008183333 0 23.99553333 0.184
1.336533333 0.018 1.183083333 0 - -
1.57115 0.025 1.829566667 0 - -
2.072966667 0 2.474333333 0 - -
2.2156 0.018 - - - -
2.854116667 0.033 - - - -
3.087666667 0.014 - - - -
Stimulation waveform data
To find records of the individual voltage transients produced from each stimulation trial, navigate to the ArucoDetection → MATLAB Behavior Program → StimDataOutput folder.
Within that folder, a list of additional folders will be produced for each behavioral session that was run. Open the folder specific to a behavioral session that just concluded and here you will find two copies of each stimulus: 1) a MATLAB data file containing the points needed to reproduce an individual waveform and 2) a .png file containing the graphed image of that waveform.
Note: Only the voltage waveforms are saved to these files and not the current. Since the generated pulse is current-controlled, only the voltage waveforms are needed.
ArUco engagement data
The data to determine how well the ArUco marker tracking program scored behaviors of distraction versus engagement can be found within the same session data sheets described in step E1. Open one of the session sheets and navigate to Sheet 2. Here, the Distraction Annotation column within that sheet contains the classification of each individual trial with a value of “0” indicating engagement and a “2” indicating distraction (see Table 2 for an example).
Note: This column does not contain ground-truth data. Compare these results to human-annotated data to quantify performance metrics.
ArUco tracking performance data
After a behavioral session concludes, the AnimalDetect.py ArUco tracking code in Visual Studio Code will produce a text file called detection_results.txt within the same directory. Open this file to view the number of total video frames in which the ArUco marker was detected by camera #1 exclusively, camera #2 exclusively, both cameras, or neither.
Caution: This file is overwritten after each behavioral session.
Verification of ArUco tracking setup
Once the behavioral chamber is fully operational, the user can evaluate the performance of the ArUco tracking setup by calculating accuracy metrics, comparing the automated scoring program against human-generated annotations. To do this, the user must manually score each trial of a behavioral session, marking whether the animal was “engaged” or “distracted.”
Note: Screen recording programs can assist with human annotations.
Afterward, open the corresponding Excel session file and review the Distraction Annotation column in Sheet 2 (see Table 2 for an example). Against your annotations, tally true positives (both human and program agree on engagement), true negatives (both agree on distraction), false positives (program marks engagement, human marks distraction), and false negatives (program marks distraction, human marks engagement) to fill out the components of a confusion matrix.
Once generated, the custom MATLAB file found at ArucoDetection → Analysis Code → Model_Classification_Metrics.m can be used to calculate the following accuracy metrics: accuracy, precision, sensitivity, specificity, F1-score, d-prime, and Matthew’s correlation coefficient (MCC). Upon starting the program, it will prompt the user to input their four confusion matrix values (one at a time), then output the accuracy metric values as text within the terminal.
Note: We leave it up to the user to determine satisfactory accuracy metrics. However, we suggest having an overall accuracy of at least 90% for robust performance. As a benchmark, we previously achieved an overall accuracy of 98% [14].
Troubleshooting: If the expected accuracy value was not met, check the total number of ArUco marker-identified frames (described in step E4). This should inform the user if one or both cameras are underperforming in terms of consistently tracking the marker. If sufficient frames are being detected, see step D2 for further information about recalibrating the cameras.
Data analysis
A full description of the data analysis procedures can be found within the Materials and Methods section for the following research articles: 1) Smith et al., Real-Time Assessment of Rodent Engagement Using ArUco Markers: A Scalable and Accessible Approach for Scoring Behavior in a Nose-Poking Go/No-Go Task, eNeuro [14], and 2) Smith et al., Behavioral Paradigm for the Evaluation of Stimulation-Evoked Somatosensory Perception Thresholds in Rats, Frontiers in Neuroscience [13].
Automated scoring of engagement
The automated assessment and scoring of animal engagement are processed in real-time during each behavioral session using the ArucoDetection → ArUco Tracking Program → AnimalDetect.py program. This program continuously evaluates the ArUco marker's position and orientation to determine whether the animal is "engaged" or "distracted" during any individual trial. Specifically, if the marker is detected within the region of interest—from the chamber’s midline to the wall containing the nose-poke and reward modules—and oriented toward that wall (yaw angle 5°–175°; 90° indicating orthogonality between the marker’s edge and wall), then the animal is scored as "engaged"; otherwise, it is scored as "distracted." Furthermore, to be considered “engaged”, the detected ArUco marker must only meet the listed requirements for a single captured frame. However, for a trial to be scored as distracted, the animal must be distracted throughout the entire trial duration. The results of this scoring process can be found in the Distraction Annotation column data from Sheet 2 of the exported Excel session data sheet (see step E3).
Post-session engagement analysis
Only after the user is satisfied with the ArUco classification performance metrics outlined in section F, should they proceed with the overall post-session behavioral engagement analysis. This process involves combining the Distraction Annotation column data from Sheet 2 of the exported Excel session data sheet (see Table 2) with the x-axis trial values from Sheet 3 (see Table 3) to create a sequential array of trial response data. The data is then analyzed using a rectangular kernel convolution to detect critical transition points between engagement and distraction, identifying the time point where the probability of engagement drops below 50%, thereby determining task disengagement and an optimal task duration for sustained engagement. For this analysis, use the custom MATLAB file found within ArucoDetection → Analysis Code → DynamicEngagementEvaluation.m to evaluate engagement over time in the nose-poking go/no-go behavioral task. To utilize the file, simply run the code in MATLAB and type in the duration of the session being analyzed. Then, when prompted, select the behavioral session file that needs to be analyzed from the ArucoDetection → MATLAB Behavior Program → ExcelSessionData → SessionDataSheets folder. Once the file is fully processed, a MATLAB plot showing the probability of engagement over time will appear. Inside of this plot, the time point at which animal disengagement is calculated will be shown as a blue vertical line. Furthermore, the probability of engagement array values for the session will be displayed in the terminal window as text. See Video 2 for an example of how to run this program effectively. Note that, when using this program to determine a maximum session duration, this analysis should be averaged across multiple sessions and animals to inform of this decision. However, this program can also be used to further define individual session exclusion criteria; in addition to the ArUco tracking program’s standard distraction labeling, the user may choose to exclude all trials following the point where the animal was deemed completely disengaged.
Video 2. Running the post-session engagement analysis program.Instructional video that outlines the steps required to process the exported behavioral session data sheets for determining task disengagement and optimal task durations based on the automated scoring of animal engagement data.
Routine behavioral analysis
Alongside engagement evaluations, behavioral task performance can also be routinely analyzed through post-session programs. Although the standard behavioral program does automatically calculate the overall accuracy for each session, it does not account for the exclusion of distracted trials. To calculate the adjusted accuracy, the user must tally the number of distracted trials for each session type (stim, silent, or alt-stim), then subtract those values from the “missed” category of each session type, providing the updated confusion matrix values: true positives (hits on stim trials), true negatives (misses on silent trials), false positives (hits on silent trials), and false negatives (misses + late hits on stim trials) found within the exported Excel session file following each session. The location of the dataset required for producing this confusion matrix is described in step E1 (see Table 1 for an example). Once generated, the custom MATLAB file found at ArucoDetection → Analysis Code → Model_Classification_Metrics.m can be used to calculate the following revised accuracy metrics: accuracy, precision, sensitivity, specificity, F1-score, d-prime, and Matthew’s correlation coefficient (MCC). Upon starting the program, it will prompt the user to input their four confusion matrix values (one at a time), then output the accuracy metric values as text within the terminal.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Smith et al. [13]. Behavioral Paradigm for the Evaluation of Stimulation-Evoked Somatosensory Perception Thresholds in Rats. Frontiers in Neuroscience. (Figures 1, 3, and 6).
Smith et al. [14]. Real-Time Assessment of Rodent Engagement Using ArUco Markers: A Scalable and Accessible Approach for Scoring Behavior in a Nose-Poking Go/No-Go Task. eNeuro. (Figures 1–3).
Validation of hardware components
Through combined research efforts across multiple studies, including Smith et al. [13,14], the behavioral chamber has been utilized in over 500 sessions, totaling more than 250 h of robust performance, enduring the animals' pulling, biting, and scratching. During this time, the only component that required replacement was the tether, which was replaced three times. Additionally, the ArUco assembly pieces have been used in over 300 sessions, accumulating more than 150 h of use, with the only replacement being the paper-printed ArUco pattern, which was reprinted twice.
Validation of ArUco engagement assessment and scoring
Smith et al. [14] validated the ArUco engagement assessment and scoring procedures described in this protocol, demonstrating the system's accuracy and reliability. In this study, the same ArUco tracking system described in this protocol achieved an overall classification accuracy of 98.3%, precision of 98.5%, sensitivity of 99.7%, specificity of 70.8%, and F1-score of 99.1% when compared with human annotations across ~1,000 individual trials. These results indicated strong classification performance when applied during the go/no-go behavioral task featured in the BehaviorApp.mlapp program.
To further validate the use of this protocol without needing to set up the equipment, researchers can reproduce the findings of the 2024 article [14] by accessing the raw behavioral session data within the relevant GitHub repository (https://github.com/tomcatsmith19/ArucoDetection) under the folder ArucoDetection → All Session Data and Results. The raw data can then be analyzed using the provided analysis code within the ArucoDetection → Analysis Code folder. The corresponding videos for these sessions can be provided upon request.
General notes and troubleshooting
General notes
Automation and confirmation biases
The overall behavioral system was designed to automatically score and record consistent variables such as nose-poking instances, timestamps, electrical stimulation outputs, and estimates of animal engagement. These automated processes help to eliminate confirmation biases during the data collection phase, especially when distinguishing trials of engagement from distraction. This ensures a consistent evaluation approach applicable for large-scale studies involving multiple human evaluators. However, while confirmation biases are minimized during data collection, additional biases may arise during post-session analysis if the exported data is manually edited by the user before being processed in the provided analysis code or outside sources.
System setup and flexibility
The behavioral chamber setup described in this protocol was designed as a flexible starting point rather than a rigid framework. Although step-by-step instructions for reproducing our exact chamber are provided, we understand that each setting may require unique specifications like differing stimulators or stimulation patterns, microelectrode array connectors, number of cameras, etc. Therefore, on the software side, all MATLAB and Python code has been commented or described within this protocol in hopes of easing the customization process with the bulk of the complete program split between the two main files: MATLAB → BehaviorApp.mlapp and Python → AnimalDetect.py. Taking this one step further, the base ArUco tracking components can be extracted from the behavior app, leaving researchers with the ability to insert it into their own configurations. From the BehaviorApp.mlapp file, the only code that is required to receive distraction results from the provided AnimalDetect.py file are the following isolated commands:
% Server socket setup with Python code for ArUco marker detection
import java.net.ServerSocket
import java.io.*
server_socket = ServerSocket(9999);
client_socket = server_socket.accept;
input_stream = client_socket.getInputStream;
d_input_stream = DataInputStream(input_stream);
output_stream = client_socket.getOutputStream;
d_output_stream = DataOutputStream(output_stream);
% Send a 1 to Python script to begin ArUco maker detection
d_output_stream.writeUTF(num2str(1));
% Send Python script a 2 to stop tracking ArUco markers
d_output_stream.writeUTF(num2str(2));
% Receive ArUco marker distraction results from Python script
data_received = str2double(d_input_stream.readUTF());
% Close all ArUco maker detection tracking and Python code.
% Disconnect the server socket and clear variables.
d_output_stream.writeUTF(num2str(4));
pause(5);
input_stream.close;
d_input_stream.close;
client_socket.close;
server_socket.close;
However, if Python is the only desired language, then extract the socket class code from the AnimalDetect.py file. An example of such code in which an ArUco marker is continuously tracked until the key ‘q’ is pressed can be seen below:
import cv2
import numpy as np
font = cv2.FONT_HERSHEY_PLAIN
dictionary = cv2.aruco.getPredefinedDictionary(cv2.aruco.DICT_4X4_50)
# camera setup
cap = cv2.VideoCapture(1)
cap.set(cv2.CAP_PROP_FOCUS, 20)
cap.set(cv2.CAP_PROP_ZOOM, 0)
print("Camera connected")
while True:
# Read frame from the camera
success, frame = cap.read()
if not success:
break
# Detect ArUco markers in the frame
corners, marker_ids, rejected = cv2.aruco.detectMarkers(frame, dictionary)
# if the marker was found
if corners:
for corner, marker_id in zip(corners, marker_ids):
corner = corner.reshape(4, 2)
corner = corner.astype(int)
top_right, top_left, bottom_rght, bottom_left = corner
# Calculate radian yaw angle of the pose
delta_pos = top_left - top_right
yaw = np.degrees(np.arctan2(delta_pos[1], delta_pos[0]))
# Calculate centroid of the marker
centroid = np.mean(corner, axis=0).astype(int)
# Print yaw value in the center of the marker
yaw_text = f"{int(yaw)} deg"
cv2.putText(frame,yaw_text,tuple(centroid),font,1.3,(0,0,255),2)
# Draw marker outline and label corners
cv2.polylines(frame,[corner],True,(255,0,255),3,cv2.LINE_AA)
cv2.putText(frame,"FL",tuple(top_right),font,1.3,(255,0,255),2)
cv2.putText(frame,"FR",tuple(top_left),font,1.3,(255,0,255),2)
cv2.putText(frame,"BR",tuple(bottom_rght),font,1.3,(255,0,255),2)
cv2.putText(frame,"BL",tuple(bottom_left),font,1.3,(255,0,255),2)
# show camera frame with ArUco tracking
cv2.imshow("Camera", frame)
if cv2.waitKey(1) & 0xFF == ord('q'):
break
# close camera stream
cap.release()
cv2.destroyAllWindows()
On the hardware side, most items such as the sound-attenuating chamber, operant conditioning chamber, T-Slot rails, commutator, camera mounts, and the cameras themselves can each be substituted without affecting the application. In addition, the ArUco marker simply needs to be flat, fixed to the animal’s head in the orientation displayed within Figure 9, and hold a direct line-of-sight with the cameras. If the reader is unable to utilize the ArUco mounting hardware in step C1, the reader may investigate other methods such as water-soluble glue or a custom helmet to rigidly and robustly fix the marker to the animal’s head. In terms of electronics, devices meeting the criteria in step A9a will also be viable. Furthermore, if an existing setup utilizes a 3.3V logic microcontroller a level shifter may be utilized to shift the logic voltage to 5V. This overall flexibility enables researchers to tailor the setup precisely to their experimental setup without being constrained by a predefined configuration.
Custom behavioral task integration
To extend the functionality of the pre-loaded MATLAB behavior app with custom behavioral tasks, the code utilizes a structured approach within a switch case framework (lines 550–12388 from the BehaviorApp.mlapp file). Here, each behavioral task is encapsulated within its own case statement, facilitating direct modular expansion. By adding a new case to the switch statement and including the name of that case (Task Name) in the dropdown list located within the Session Setup tab → Task Name dropdown box of the app (see procedure steps B2f–i), researchers can seamlessly integrate additional behavioral tasks into the application. The existing code provides a template and examples from the pre-loaded tasks, offering guidance for developing and integrating new tasks. This approach ensures that the MATLAB behavior app remains adaptable and scalable, supporting the incorporation of novel experimental paradigms.
Acknowledgments
This work was supported in part by the National Institutes of Health, National Institute for Neurological Disorders and Stroke (R01NS110823, GRANT12635723, J.R.C. and J.J.P.), diversity supplement to parent grant (A.G.H-R.), a Research Career Scientist Award (GRANT12635707, J.R.C.) from the United States (US) Department of Veterans Affairs Rehabilitation Research and Development Service, and the Eugene McDermott Graduate Fellowship from The University of Texas at Dallas (202108, T.J.S.). This protocol was derived from the original research paper “Real-Time Assessment of Rodent Engagement Using ArUco Markers: A Scalable and Accessible Approach for Scoring Behavior in a Nose-Poking Go/No-Go Task” [14].
Competing interests
The authors declare no competing financial interests.
Ethical considerations
All animal handling, housing, and procedures were approved by The University of Texas at Dallas IACUC (protocol #21-15) and in accordance with ARRIVE guidelines [15].
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Article Information
Publication history
Received: Jun 28, 2024
Accepted: Sep 8, 2024
Available online: Sep 28, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Neuroscience > Behavioral neuroscience > Sensorimotor response
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5,099 | https://bio-protocol.org/en/bpdetail?id=5099&type=0 | # Bio-Protocol Content
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Optimizing Transmembrane Protein Assemblies in Nanodiscs for Structural Studies: A Comprehensive Manual
FV Fernando Vilela *
CS Cécile Sauvanet *
AB Armel Bezault
NV Niels Volkmann
DH Dorit Hanein
(*contributed equally to this work)
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5099 Views: 563
Reviewed by: Beatrice Li Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Structural Biology Dec 2022
Abstract
Membrane protein structures offer a more accurate basis for understanding their functional correlates when derived from full-length proteins in their native lipid environment. Producing such samples has been a primary challenge in the field. Here, we present robust, step-by-step biochemical and biophysical protocols for generating monodisperse assemblies of full-length transmembrane proteins within lipidic environments. These protocols are particularly tailored for cases where the size and molecular weight of the proteins align closely with those of the lipid islands (nanodiscs). While designed for single-span bitopic membrane proteins, these protocols can be easily extended to proteins with multiple transmembrane domains. The insights presented have broad implications across diverse fields, including biophysics, structural biology, and cryogenic electron microscopy (cryo-EM) studies.
Key features
• Overview of the sample preparation steps from protein expression and purification and reconstitution of membrane proteins in nanodiscs, as well as biobeads and lipids preparation.
• Focus on single-span bitopic transmembrane proteins.
• Includes protocols for validation procedures via characterization using biochemical, biophysical, and computational techniques.
• Guide for cryogenic electron microscopy data acquisition from vitrification to image processing.
Keywords: Nanodisc Single-span Bitopic membrane proteins Cryogenic electron microscopy Structural determination Three-dimensional reconstruction
Graphical overview
Schematic representation of the workflow for membrane protein reconstitution in nanodiscs. The upper blue box illustrates the key steps involved in preparing bare nanodiscs from lipid and membrane scaffold protein preparations (1), removal of detergent (2), and purification of nanodiscs (3). The quality of the samples is verified through various biochemical and imaging approaches (4). The lower red box outlines the step-by-step membrane protein-nanodiscs reconstitution process. Detergent-solubilized and purified membrane protein samples are mixed with the chosen lipid sample (1), followed by the addition of membrane scaffold proteins (2). Dialysis in the presence of bio-beads allows for detergent removal (3), leading to the reconstitution of membrane protein-nanodiscs complexes, followed by size exclusion chromatography purification (4). Finally, we present various biophysical and screening methods to assess the quality and homogeneity of the samples before structural analysis (5).
Background
Transmembrane proteins play a pivotal role in the interactions of cells with their environment. Their significance in pharmacology is underscored by their targeting over 50% of available drugs [1]. A more profound comprehension of the structure and function of these proteins can greatly enhance our understanding of disease development and the creation of effective medications. However, exploring the structure of membrane proteins poses a significant challenge due to the hydrophobic nature of their transmembrane domains, which play crucial functional roles. A comprehensive study necessitates examining these proteins in their entirety, including their transmembrane domains, within stable lipid environments close to their native conditions in the cell. Meeting these prerequisites calls for the development of specialized biochemical tools tailored for the study of transmembrane proteins.
One such tool is the nanodisc (ND), a nanoscale disc-shaped lipid bilayer encircled by a belt formed by two membrane scaffold proteins (MSPs). Nanodiscs provide numerous advantages compared to traditional solubilization methods, including enhanced stability, uniformity, and the ability to explore membrane proteins in their native lipid environment. Since the pioneering work of Sligar and colleagues, MSPs have revolutionized various fields including structural biology and biophysics [2–6]. MSP nanodiscs maintain the solubilization of full-length transmembrane proteins, preventing detergent-induced denaturation. When the size and molecular weight (MW) of the transmembrane protein significantly exceed that of the NDs or when the transmembrane domain size approaches that of the nanodisc and displaces a large fraction of the lipids, the system serves as a potent tool for structural biology investigations. For example, NDs have been instrumental in exploring the structure and function of membrane proteins such as G protein–coupled receptors, ion channels, and transporters [7–9]. However, this scenario changes when the NDs and the transmembrane protein share a similar MW range and the transmembrane displaces only a small fraction of the nanodisc lipids, for example, single-spanning membrane receptors such as integrin. Under the premises where the nanodiscs make a substantial contribution to the overall structural analysis, imperfections in the nanodisc preparations directly translate into lower resolution in cryo-EM structural studies.
Here, we detail comprehensive and reliable biochemical and biophysical protocols that enable the structural examination of transmembrane proteins, particularly when their size and MW are similar to the nanodiscs, and they occupy only a small fraction of the nanodisc. Our focus is a thorough manual for researchers concerning the use of MSPs and MSP-based NDs in the structural and biophysical examination of single-pass alpha-helix transmembrane proteins, often referred to as bitopic transmembrane proteins. While this publication is tailored for bitopic transmembrane proteins, the principles outlined can be applied to the study of proteins with multiple transmembrane domains.
Materials and reagents
Biological materials
E. coli BL21(DE3) chemically competent cells (Sigma-Aldrich, catalog number: CMC0014)
Expression vectors of membrane scaffold proteins (MSP) from Addgene: MSP1D1 (Addgene, catalog number: 20061), MSP1E3D1 (Addgene, catalog number: 20066), and MSP2N2 (Addgene, catalog number: 29520) for 9, 12, and 15 nm nanodiscs, respectively
Purified and monodisperse target membrane protein of your interest. Here, we use ACE2 protein as an example of single-pass transmembrane protein, which was purified in 50 mM HEPES pH 8, 250 mM NaCl, 0.5% DDM, and 2.5 mM desthiobiotin (adapted from Yan et al. [10])
Reagents
POPC lipids in chloroform, 25 mg/mL (Avanti Polar Lipids, catalog number: 850457C)
2× YT medium (Sigma-Aldrich, catalog number: Y2377)
94 mm Petri dish (Grenier Bio-One, catalog number: 633180) containing 20 mL of LB agar media
Kanamycin at 50 µg/mL for selecting bacteria carrying vectors of MSPs (Sigma-Aldrich, catalog number: K0254)
Trizma base (purity ≥ 99.7%) (Sigma-Aldrich, catalog number: 93350)
Hydrochloric acid 37% vol/vol (HCl) (Sigma-Aldrich, catalog number: 258148)
Sodium chloride (NaCl); purity ≥ 99% (Sigma-Aldrich, catalog number: 793566)
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: 03677)
Imidazole (purity ≥ 99.5%) (Sigma-Aldrich, catalog number: 56750)
Ethanol absolute (99.8% analytical reagent grade) (Fisher Chemical, catalog number: E/0650DF/15)
CAUTION: Ethanol is harmful if swallowed, inhaled, or by skin absorption. Wear appropriate protective glasses, gloves, and lab coat. Handle under a chemical hood.
MilliQ H2O
Buffer components required by the target membrane protein of your interest
Methanol (Sigma-Aldrich, catalog number: 322415)
CAUTION: Methanol is toxic if swallowed, inhaled, or in contact with skin. Wear appropriate protective glasses, gloves, and lab coat. Handle under a chemical hood.
Chloroform (99.5%) (Sigma-Aldrich, catalog number: C2432)
CAUTION: Chloroform is toxic if swallowed or inhaled. Always wear googles, gloves, and lab coat. Carry operations under a chemical hood.
Sodium cholate (NaCholate) (Sigma-Aldrich, catalog number: C6445)
Triton X-100 (Sigma-Aldrich, catalog number: T9284)
PMSF (Roche, catalog number: 10837091001)
Anti-proteases cocktail (Roche, catalog number: 11836170001)
Benzonase® (Sigma-Aldrich, catalog number: 1016970001)
Lysozyme (Sigma-Aldrich, catalog number: 1052810500)
Isopropyl b-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, catalog number: I6758)
Bio-Beads SM-2 adsorbents (Bio-Rad, catalog number: 1528920)
Antibody for His-tag detection (Thermo Scientific, catalog number: 15547466)
4–15% Mini-PROTEAN® TGXTM precast protein gels (Bio-Rad, catalog number: 4561086)
Liquid nitrogen (pure from Air Liquide ALPHAGAZ)
CAUTION: Liquid nitrogen can cause severe burns. Always wear required personal safety equipment such as safety glasses, lab coat, and insulated gloves when handling.
Nitrogen (N2) gas (purity ≥ 99,999%) (Air Liquide ALPHAGAZ)
Ethane gas (pure from Air Liquide ALPHAGAZ)
CAUTION: Ethane gas is extremely flammable. Store in tightly closed containers in a cool, well-ventilated area.
Solutions
Lipid solubilization buffer (see Recipes)
Size exclusion chromatography buffer (see Recipes)
Lysis buffer (see Recipes)
MSP wash buffer (see Recipes)
Elution buffer (see Recipes)
Recipes
Lipid solubilization buffer
Reagent Final concentration Quantity or Volume
Trizma pH 7.4 20 mM 1.21 g
NaCl 100 mM 2.92 g
EDTA pH 7.4 0.5 mM 93.06 mg
NaCholate 70 mM 15.07 g
H2O n/a Final volume to 500 mL
Note: Small increments of HCl are used to bring the solution to pH 7.4.
Size exclusion chromatography buffer
Reagent Final concentration Quantity or Volume
Trizma pH 7.4 20 mM 1.21 g
NaCl 100 mM 2.92 g
EDTA pH 7.4 0.5 mM 93.06 mg
H2O n/a Final volume to 500 mL
Note: Small increments of HCl are used to bring the solution to pH 7.4.
Lysis buffer
Reagent Final concentration Quantity or Volume
Trizma pH 8.0 50 mM 454.28 mg
NaCl 300 mM 1.31 g
Triton X-100 1% 750 µL
PMSF 1 mM 13.06 mg
Tablet anti-protease cocktail 1 tablet
Benzonase® 250 unit 1 µL
Lysozyme 0.7 mg per mL of lysis buffer 52.5 mg
H2O n/a Final volume to 75 mL per L of E. coli culture
Note: Small increments of HCl are used to bring the solution to pH 8.0. Initially, a larger volume of solution containing only Trizma, NaCl, and Triton X-100 can be made. On the day of the experiment, use 75 mL of this solution and supplement with PMSF, anti-protease cocktail, benzonase, and lysozyme to perform bacterial lysis.
MSP wash buffer
Reagent Final concentration Quantity or Volume
Trizma pH 8.0 50 mM 3.03 g
NaCl 300 mM 8.77 g
H2O n/a Final volume to 500 mL
Note: Small increments of HCl are used to bring the solution to pH 8.0.
Elution buffer
Reagent Final concentration Quantity or Volume
Trizma pH 8.0 50 mM 3.03 g
NaCl 300 mM 8.77 g
Imidazole 500 mM 17.02 g
H2O n/a Final volume to 500 mL
Note: Small increments of HCl are used to bring the solution to pH 8.0.
Laboratory supplies
Cryo-EM grids such as Quantifoil 1.2/1.3 400 mesh copper (Quantifoil, catalog number: N1-C14nCu40-01)
Filter paper for VitroBot (Grade 595, 50 mm, AA00420S) (Fisher Scientific, catalog number: 09-924-170)
Hamilton syringe 500 Ul (Merck, catalog number: 24523)
Spectra/Por 3 dialysis tubing, 3.5 kD MWCO (Repligen, catalog number: 13272)
Costar® Spin-X® centrifuge tube filters, 0.22 µm pore CA membrane, nonsterile, 100/case (Corning, catalog number: 8161)
Parafilm (Bemis, catalog number: PM996)
10 mL glass tubes (Kimble Chase, catalog number: A407511500611)
His-trap column 5 mL affinity chromatography (Cytiva, catalog number: 17524802 or similar)
Size-exclusion Superdex 200 increase 24 mL chromatography column (Cytiva, catalog number: 28990944 or similar)
Amicon concentrators cutoff 50 kDa (Millipore, catalog number: UFC505096)
InstantBlue® Coomassie protein stain (Abcam, catalog number: ab119211)
Storage dewar for EM grids (Worthington LD Series Liquid Nitrogen Dewars, catalog number: F9143-1EA)
Laboratory glassware and plasticware for preparation, storage, and handling of reagents and samples
NalgeneTM single-use PETG Erlenmeyer flasks with baffled bottom 2 L (Nalgene, catalog number: 4113-2000)
PES (Polyethersulfone) syringe filters (Thermo Scientific, catalog number: CH2213-PES)
Micropipette tips and plastic pipettes cut at about 1/3 from the tip to be used when manipulating Bio-Beads
Equipment
Spectrophotometer NanoDrop (Thermo Scientific, or similar) for OD600 bacterial density and absorbance at 280 (A280) protein concentration measurements
Incubator with temperature control for Erlenmeyer flasks
Rotator TR Series TR-200 (Stuart/Cole-Palmer, model: TR-200)
Mini-PROTEAN Tetra Vertical electrophoresis cell (Bio-Rad, catalog number: 1658004)
Batch sonicator (Fisher Scientific, catalog number: FB15046)
pH meter (Thermo Scientific, catalog number: STARA1115) and calibration reagents (Thermo Scientific, catalog number: 910199)
Dry bath with temperature control such as Eppendorf ThermoMixer F2.0 (Eppendorf, catalog number: 5387000013)
FPLC Equipment, Akta System (Cytiva) or similar
Western blot device iBind and accessories (Thermo Scientific, catalog number: SLF2000)
Imaging system for SDS-Page and western blot (Bio-Rad, model: Gel DocTM EZ System)
Lab bench centrifugation equipment such as Eppendorf centrifuge 5424 R (Eppendorf, model: 5424 R)
Bench microcentrifuge such as Scientific D1008 Palm microcentrifuge (DLAB, catalog number: 9031001012)
Sample storage equipment at -80 °C, -20 °C, and 4 °C
Chemical hood
VitroBot Mark IV (or similar) (Thermo Scientific)
Software and datasets
Chromatography Akta software: Unicorn 6.3 or later version, Cytiva
Imaging software for western blot and SDS page: Image Lab 6.0 or later version, Bio-Rad
DLS software: DYNAMICS software 7.9 or later version, Waters
Data Acquisition software for single particle analysis: EPU 3.3.0.5176REL or later SerialEM 3 or higher version
Data analysis software for single particle analysis: CryoSPARC version 4.3.0 or higher, Relion 4 or equivalent
Procedure
Our protocol describes the reconstitution of single-span, bitopic transmembrane proteins, although it can easily be extended to proteins with multiple transmembrane domains. We begin by outlining the sample preparation steps required before reconstituting nanodisc samples. These steps can be completed several days to weeks in advance. We provide a detailed description of each step, along with essential precautions to ensure success. Finally, we present various methods to evaluate the quality and homogeneity of the samples before structural analysis.
Lipids preparation for reconstituting nanodiscs assemblies
We describe the preparation of POPC lipids for nanodisc reconstitution, though similar protocol can be applied to other lipids or mixed samples (Note 1). This protocol builds on the work of Sligar and colleagues, with contributions from researchers from various fields [2–4].
Prepare a 10 mL glass tube by washing it with H2O MilliQ and ethanol at least three times each step, then dry it with N2 gas.
Add 2 mL of chloroform to the tube and rotate it so the inside walls can be washed. Discard all chloroform. Dry the tube with N2 gas stream.
Use a glass syringe of 500 µL and wash it using the same procedure as for the glass tube.
Gently transfer 500 µL of the POPC lipids chloroform stock (25 mg/mL) to the prepared glass tube (on the bottom of the tube, without touching the walls) using the syringe.
Close the glass tube with parafilm.
Add a gentle stream of N2 gas inside the commercial vial to protect the lipids stock from oxidation. Seal the vial and store it at -20 °C.
Dry the POPC lipids with a gentle stream of N2 gas (10–15 min).
Remove residual chloroform by desiccation under vacuum for at least 2 h. The lipids will form a film at the bottom of the glass tube.
Resolubilize the lipids in 500 µL of lipid solubilization buffer to obtain a solution of POPC at 32.9 mM concentration (Note 2).
Perform a gentle agitation (max. 100 rpm) for 5–10 min until all lipids are well resuspended.
Pursue the lipids resuspension by bath sonicating using a power of 37 kHz and cycles of 1 min of the bottom of the glass tube until the lipid solution becomes transparent (Note 3).
Transfer the new lipid stock to an Eppendorf and keep it at 4 °C for long-term storage.
Preparation of Bio-Beads-SM2 for detergent removal during nanodiscs reconstitution
Weigh 1 g of Bio-Beads-SM2 from the commercial vial into a 50 mL Falcon tube.
Add up to 50 mL of methanol to the solution of Bio-Beads.
Rotate gently (max. 20 rpm) for 2 h at room temperature (RT) using rotator TR-200 under the hood. Make sure the solution of Bio-Beads is well homogenized.
After decanting the Bio-Beads, eliminate the methanol.
Add 50 mL of MilliQ H2O and rotate gently (max. 20 rpm) for 1 h at RT to wash Bio-Beads from methanol.
After decanting the Bio-Beads, eliminate H2O (Note 4).
Repeat this wash two times, to be sure all methanol is eliminated.
Eliminate 30 mL of H2O from your Falcon tube.
Transfer Bio-Beads solution (Bio-Beads + 20 mL of H2O) into the Spectra/Por 3 dialysis tubing membrane (Note 5).
Perform the buffer exchange with 1 L of size exclusion chromatography buffer with agitation (max. 100 rpm) O/N at 4 °C.
Recover all Bio-Beads into a 50 mL Falcon tube (Note 6).
Wait 5–10 min for the beads to sediment in the bottom of the Falcon tube.
Adjust the Bio-Beads concentration for 200 mg of beads per milliliter by eliminating the buffer volume.
Store at 4 °C until use.
Membrane scaffold protein preparation (Note 7)
Perform a bacterial transformation with the desired MSP plasmid (for this protocol, we used the plasmid encoding for MSP1D1 (Note 8) by a heat-shock method at 42 °C for 45 s using the E. coli strain BL21(DE3) Gold (Note 9).
Plate the bacteria after transformation using the antibiotic resistance conditions of the MSP expression vector used, and incubate O/N at 37 °C.
Take one bacterial colony to inoculate 100 mL of 2× YT medium in an Erlenmeyer flask (for a final 2 L culture production).
Incubate O/N at 37 °C with agitation (between 160 and 180 rpm) to saturate the bacterial culture.
Prepare 2 L of 2× YT medium culture using the respective antibiotic resistance conditions (Note 10).
Split the 2 L culture medium into two 2 L Erlenmeyer flasks to ensure good oxygenation of bacteria (1 L in each Erlenmeyer).
Measure the OD600 of the overnight culture.
Dilute the overnight culture into 2 L of 2× YT medium to get an OD600 = 0.2.
Incubate the culture at 37 °C with agitation at 180 rpm.
Induce protein expression with 1 mM final concentration of IPTG when the E. coli cultures are between 0.6 and 0.75 OD600.
Incubate the culture for 4 h at 37 °C as previously.
Collect the bacterial pellets by centrifuging the bacterial culture at 4,000× g for 30 min (Note 11).
Collect all samples to test protein expression via SDS-PAGE analysis.
Resuspend the pellet in 75 mL of lysis buffer.
Incubate the lysis resuspension for 1 h with agitation at 4 °C.
Clarify the lysate by centrifugating at 9,000× g minimum for 1 h at 4 °C.
Recover the soluble fraction.
Filter supernatant through 0.22 µm pore size syringe filters.
Apply the filtered supernatant on a His-tag affinity chromatography using a 5 mL column, previously equilibrated with lysis buffer without detergent.
Wash with 10 column volumes (CV) of lysis buffer and then with 10 CV of MSP wash buffer.
Conduct MSP elution using an imidazole gradient ranging from 0 to 500 mM at a flow rate of 2 mL/min for 7.5 min by adding the elution buffer imidazole to achieve a final imidazole concentration of 500 mM.
Collect all samples of the purification for SDS-PAGE analysis.
Pool the elution fraction with the higher MSP quantity and purity.
Perform dialysis O/N using Spectra/Por 3 dialysis membrane tubing against 1 L of size exclusion chromatography buffer at 4 °C (Note 12).
Recover the sample, centrifuge, and filter it through a 0.22 µm membrane pore.
Measure MSPs concentration by absorbance at 280 nm (Note 13).
Analyze the purity of the purified and dialyzed MSPs by loading an SDS-PAGE 4%–15% acrylamide with 1–2 µg of protein.
Scan the gel using imaging equipment and save the image’s raw data. Then, using image analysis software, evaluate the purity percentage of your MSPs (Note 14).
Aliquot MSP proteins in 100 µL samples of 1–2 mg/mL to avoid multiple freeze-thaw cycles and store them at -80 °C after flash-freezing in liquid nitrogen (Note 15).
Bare-nanodiscs preparation and reconstitution
Thaw MSP purified protein aliquots.
Centrifuge at 12,000× g for 5 min at 4 °C.
Determine the protein concentration of the supernatant by measuring absorbance at 280 nm using Nanodrop equipment or equivalent (Note 16).
Calculate the MSP volume you need for a minimal amount of 20 nmol (Note 17).
Calculate the volume of lipids corresponding to the desired molar ratio (Note 18).
Mix both MSP and lipid using the determined volumes.
Incubate the sample for 1 h at 4 °C with 12 rpm agitation.
During that time, calculate the amount of Bio-Beads needed to completely remove detergents present in the sample used previously to solubilize lipids (Note 19).
Add Bio-Beads and incubate your nanodiscs preparation overnight at 4 °C with 12 rpm agitation.
The following day, transfer the mix to 1 mL filter centrifuge tubes and centrifuge samples using a bench microcentrifuge for a fast spin of 5 s to remove Bio-Beads (Note 20).
Recover nanodiscs sample in the flowthrough of the tube. Be sure that all Bio-Beads are correctly removed from the sample and that you recovered the entire sample.
Using concentrators with a cutoff of 50 kDa, concentrate your nanodiscs preparation to a final volume of 500 µL by centrifugation cycles of 5 min at 5,000× g and 4 °C.
Recover reconstituted nanodisc from the top part of the concentrators.
Resuspend carefully between each centrifugation cycle by up-down pipetting.
Centrifuge samples at 12,000× g for 10 min at 4 °C to eliminate possible protein aggregates.
Keep 10 µL for SDS-PAGE analysis.
Perform a size exclusion chromatography using a 24 mL Superdex 200 increase 10/300 GL.
Pre-equilibrate with size exclusion chromatography buffer at 4 °C.
Set one run per sample, injecting 490 µL with a flow rate of 0.35 mL/min.
Start to collect all samples before the elution volume of 8 mL (Figure 1A). Samples are collected in 500 µL elution fractions (Note 21).
Test all samples on SDS-PAGE to evaluate the purity of your samples (Figure 1C).
Figure 1. Characterization of bare nanodiscs and transmembrane protein nanodiscs assemblies. A. Size exclusion chromatography of bare nanodiscs (MSP1D1-POPC, blue line) and ACE2 nanodiscs (red line). The ACE2 nanodisc chromatogram shows two species corresponding to bare nanodiscs and ACE2-ND. B. SDS-PAGE of the pooled ACE2-ND fraction after gel filtration chromatography. C. SDS-PAGE of the bare nanodiscs. MSP1D1 was loaded first, followed by the MSP1D1-POPC NDs elution, demonstrating a high degree of purity of bare nanodiscs. D. SDS-PAGE of ACE2 nanodiscs purification and the resulting elution. E. Dynamic light scattering (DLS) distribution and its associated autocorrelation curve, showing a hydrodynamic radius of 4.8 nm for MSP1D1-POPC, confirming a monodisperse sample. F. DLS distribution and its associated autocorrelation curve, showing a hydrodynamic radius of 7 nm for ACE2-NDs, also confirming a monodisperse sample.
Pool fractions corresponding to the elution volume peak at 12 mL (Note 22).
Calculate the protein concentration based on the molecular weight (MW) and the measured absorbance at 280 nm of an MSP monomer.
Concentrate your nanodiscs preparations to a final volume of 50 µL using concentrators cutoff of 50 kDa by centrifugation cycles of 5 min at 5,000× g and 4 °C (Note 23).
Resuspend carefully between each centrifugation cycle by up-down pipetting before characterization. If nanodiscs cannot be analyzed directly, they can be stored at 4 °C for a maximum of 2 weeks.
Reconstitution of nanodiscs containing transmembrane proteins
Before proceeding, it is highly advisable to refine the purification protocols to achieve high purity of the transmembrane protein (> 98% on SDS-PAGE), verify monodispersity, and use a detergent compatible with Bio-Beads adsorption (e.g., sodium cholate, Tween 20) [11,12]. Additionally, it is necessary to generate 10–50 nmol of protein to proceed with incorporation experiments.
Define the molar ratios for the overall reconstitution. From our experience, a membrane protein molar ratio of 1:5 to 1:10 is a good starting point (Note 24).
Membrane protein:MSPs.
MSPs:lipids (previously optimized for bare-nanodiscs).
The amount of Bio-Beads per reconstitution.
Incubate the defined amount of membrane protein with lipids for 1 h at 4 °C and 12 rpm agitation.
Add the chosen MSP in the mix of membrane protein and lipids.
Incubate for 1 h at 4 °C with 12 rpm agitation.
Add the needed amount of Bio-Beads to remove the detergents from the sample completely.
Incubate the nanodiscs preparation for 12–15 h at 4 °C with 12 rpm agitation.
The next day, transfer the mix to 1 mL filter centrifuge tubes and centrifuge samples using a bench microcentrifuge for a fast spin of 5 s to remove Bio-Beads.
Using concentrators with a cutoff at 50 kDa, concentrate the nanodiscs preparation to a final volume of 500 µL, through centrifugation cycles of 5 min at 5,000 g at 4 °C.
Resuspend carefully between each centrifugation cycle by up-down pipetting.
Recover reconstituted nanodiscs from the upper quarter of the concentrators.
Centrifuge reconstituted nanodiscs at 12,000× g for 5 min at 4 °C.
Keep 10 µL for SDS-PAGE analysis.
Perform size exclusion chromatography using a 24 mL Superdex 200 increase 10/300 GL.
Pre-equilibrate with size-exclusion chromatography buffer at 4 °C (Note 25).
Set up one run per sample, injecting 490 µL with a flow rate of 0.35 mL/min.
Collect all samples prior to elution volume of 8 mL (Figure 1A). Samples are collected in 500 µL elution fractions.
Test all samples on SDS-PAGE to evaluate their purity and to identify the nanodiscs-incorporated transmembrane protein fractions (Figure 1B, D) (Note 26).
Define the elution volume of your sample (transmembrane proteins nanodiscs’ assemblies) by the elution volume of the elution peak (Note 27).
Pool the fractions corresponding to the elution volume nanodiscs peak.
Measure the protein concentration based on MWs and the extinction coefficient at A280 of an MSP monomer and a monomer of the target membrane protein if your protein is monomeric.
Concentrate your nanodiscs’ assembly preparations to a final volume of 50 µL using concentrators with a cutoff of 50 kDa by performing centrifugation cycles of 5 min at 5,000× g at 4 °C (Note 28).
Resuspend carefully between each centrifugation cycle by up-down pipetting.
Characterization of the quality of the reconstituted nanodiscs’ assemblies
Here, we present a variety of biophysical and biochemical methods to assess sample homogeneity, dispersity, and structural integrity before proceeding with structural studies.
Protein characterization
After reconstitution, a western blot can be employed to confirm the presence of both the His-tagged MSP and the membrane protein in the nanodiscs. Specific antibodies targeting the membrane protein can be used for detection. However, if these are unavailable, adding a tag to the membrane protein should be considered. In such cases, careful selection of the tag is important. Using the same tag, such as His, for both the MSP and the membrane protein could complicate distinguishing the membrane protein, especially if there is not a significant molecular weight difference between the membrane protein and the MSP.
If the concentration of nanodiscs allows, using a native gel can effectively show the ratio of incorporated vs. bare nanodiscs. This method maintains the native state, enabling observation without denaturation. In studies involving integrins, native gels have been particularly useful for accurately distinguishing between different forms [13].
Dynamic light scattering
Dynamic light scattering (DLS) experiments can be used to evaluate the extent of nanodiscs reconstitution. A comparative analysis of the hydrodynamic radius values between bare nanodiscs (NDs) and those incorporating membrane proteins can be performed (Figure 1E, F). The presence of the membrane protein should increase or match the hydrodynamic radius values of the bare NDs [14]. Depending on the instrument, DLS experiments can be conducted with minimal sample consumption, as samples are often recoverable from the plate. Combining DLS results with prior chromatography and western blot data should offer high confidence in determining whether reconstitution was successful or if further optimization is required. Ideally, samples should be monodispersed in terms of the percentage of mass in the sample [15] ensuring that the particles are uniform in size and distribution. This uniformity is crucial for reliable results in downstream experiments.
Analytical ultracentrifugation
Analytical ultracentrifugation (AUC) experiments can also be considered for characterizing reconstituted nanodiscs assemblies. These experiments are particularly valuable in specific cases (for more details, see Inagaki and Ghirlando [16]):
When the membrane protein contains only transmembrane domains and is fully embedded within the lipidic bilayer of the nanodiscs. AUC can differentiate bare-NDs from membrane protein nanodiscs complexes by distinguishing between the densities of the different species, thereby revealing their composition.
When studying membrane protein complexes that exhibit conformational changes, interaction affinities, or stoichiometries.
Cryogenic electron microscopy (cryoEM) of membrane proteins in nanodiscs
For quality control of nanodisc samples, homogeneity and size can be visually inspected using electron microscopy techniques. While negative staining is a convenient method, it requires drying the samples and embedding them in heavy metal salt layer, which can cause sample flattening. We recommend using cryoEM to image, acquire, and analyze the assemblies in their fully hydrated state (for more details, see Cianfrocco and Kellogg [17], Passmore and Russo [18]).
CryoEM sample preparation
i. Turn on Vitrobot [we used Vitrobot Mark IV model (Thermo Fisher)].
ii. Set the parameters to 4 °C and 100% humidity and prepare filter papers.
iii. While the Vitrobot reaches the target temperature, render the TEM substrates (grids) hydrophilic using a glow discharge apparatus (such as PELCO easiGlowTM Glow Discharge Cleaning System or Cordouan Technologies ELMO glow discharge system) or plasma cleaner (such as Gatan/Ametek Solarus II Plasma Cleaner).
iv. Enter the vitrification parameters: 30-s wait before blotting; blot for 3 s at force 2; no drain time (Note 29).
v. Liquefy ethane (Note 30).
vi. Apply 4 µL of the sample solution at the approximate 0.5 mg/mL to the EM grids (Note 31).
vii. Proceed with the plunge freezing in liquid ethane.
viii. Store the vitrified samples in a grid storage container at LN2 temperature.
CryoEM data acquisition
i. Screen the grids before starting data collection (Figure 2).
ii. On a 200 kV or 300 kV cryo-electron microscope, verify the integrity of the grids and the quality and thickness of the vitrified layer [18].
iii. Assess the concentration and homogeneity of nanodiscs assemblies (Figure 2A–G) (Notes 32 and 33).
v. For the selected grids, proceed with data acquisition using the following parameters:
1) Use parallel beam condition on a 200 kV or 300 kV cryo-electron microscope, such as Titan Krios G3i (Thermos Scientific) or a Glacios (Thermos Scientific), preferably equipped with a direct electron detection device (e.g., Falcon4i) in movie mode. If an energy filter is available (e.g., a SelectrisX or Selectris, Thermo Scientific), set the slit width to 10 eV.
2) Use data collection software like EPU (Thermo Scientific) or Serial EM [19].
3) Set magnification to achieve a pixel size of around 1 Å with a defocus range between -0.8 and -3 µm with a step of 0.2 µm; the total dose should be 50 e/Å2.
Figure 2. CryoEM sample and imaging assessment. (A–D) CryoEM micrograph of bare nanodisc samples taken at different defocus values, and (E) the associated 2D classes corresponding to a box size of 256 px with a pixel size of 0.96 Å. A. Defocus 0.5 µm. B. Defocus 1 µm. C. Defocus 1.5 µm. D. Defocus 2 µm. Scale bar 50 nm. Red circles highlight bare nanodiscs visible in the micrographs, with zoomed-in inserts showing the highlighted particles (scale bar: 10 nm). It should be noted that the nanodiscs are easier to identify at higher defocus. (F and G) CryoEM micrographs of nanodiscs at different concentrations: 0.3 mg/mL (F), 0.8 mg/mL (G). Scale bar 50 nm. Ideally, each micrograph should contain approximately 400 particles, aiming for about 1 million particles per dataset. Lower particle concentrations, as seen in (F) will result in a more costly data collection process compared to (G). Therefore, in cases like (F), efforts should be made to optimize the particle concentration.
Feret signature analysis
To assess the homogeneity of nanodisc preparations, we refer to the Feret signature analysis described in our recent publication [14]. This analysis should be done at the onset of data collection, as Feret signature methodology can identify a lack of nanodisc-sample homogeneity with only about 1,000 particles per set. This method enables real-time adjustments of data acquisition parameters for optimizing data collection strategies or aiding in decisions to discontinue ineffective imaging sessions. Feret signatures can also help to detect orientation bias, conformational diversity, and 3D misclassification in samples other than pure nanodiscs [20].
CryoEM data processing
Once data acquisition details are optimized and datasets have been acquired, pursue with initial data processing steps:
i. Import all the movies to CryoEM analysis software such as CryoSPARC [21] or Relion [22].
ii. Follow the Motion Correction and CTF Estimation steps.
iii. Pick particles manually or with software such as Blob Picker in CryoSPARC [23] or in Relion. Then, extract particles with a box size of 400 px corresponding to 384 Å box size for 9 nm nanodiscs.
iv. Generate 2D classes (Figure 2E) (Note 34).
v. Follow by a few rounds of selecting the best 2D class particles.
vi. Repeat steps F4d.iii–v using automatic picker such as Template Picker.
vii. Generate 3 Ab-initio models with the chosen software.
viii. Perform a 3D refinement with all the several generated Ab-initio models created.
ix. Generate 3D reconstruction and perform postprocessing.
Validation of protocol
Membrane scaffold protein (MSP) nanodiscs offer numerous advantages for investigating membrane proteins in biophysics and structural biology [24]. Establishing a robust biochemical workflow is essential to ensure reproducibility. Some of our work has focused on studying challenging systems of membrane proteins embedded in nanodiscs under cryogenic conditions. The protocol described here was indispensable for producing structures for several of these systems. One such system is the B-cell lymphoma 2 (Bcl-2)-associated X protein (Bax) nanodisc assemblies [25–27]. Bax, a 21 kDa protein, is a member of the Bcl-2 family and plays a crucial role in apoptotic regulation. When triggered, Bax assembles into an oligomer and inserts itself into the outer mitochondrial membrane to induce apoptosis [28]. We examined the structural properties of Bax nanodisc assemblies, despite its small molecular weight being closely correlated to those of nanodiscs, mostly fully incorporated within the lipid islands. We have also resolved structures of bitopic membrane proteins using this protocol, such as the integrin protein family, which are bidirectional, allosteric transmembrane receptors playing a pivotal role in hemostasis and arterial thrombosis [29]. The conformational landscape of human integrin αIIbβ3, while bound to its effectors, is strongly affected by the presence of the lipid bilayer [13,30]. Another example of a bitopic membrane protein resolved using our protocol is the B-cell lymphoma xL protein (Bcl-xL; 23 kD), a major suppressor of apoptosis [14]. These systems differ substantially from high-molecular-weight channels, where nanodiscs primarily serve as small solubilizers [31]. When the size of the embedded protein closely matches the size of the nanodisc, heterogeneity in nanodisc preparation can significantly impact alignment and classification accuracy during the structure determination process. Preferred orientation and conformational diversity can further complicate image analysis [32,33]. The proposed biochemical and biophysical characterization protocols, image acquisition, and real-time feedback [14] can advance preparations and lead to superior high-resolution structural determination of these challenging samples.
General notes and troubleshooting
Choosing biologically relevant lipids that closely resemble the natural membrane composition is essential for stabilizing membrane proteins and preserving their structure and function in nanodiscs [24,34]. The physical and chemical properties of the lipids, such as headgroup and tail length, can influence nanodisc size, shape, stability, and rigidity [35]. Additionally, the phase behavior of lipids, including those with high phase-transition temperatures, can enhance nanodisc stability under harsh conditions like elevated temperatures or low pH [4]. Lipid composition also affects interactions between nanodiscs and other biomolecules, which is crucial for certain applications. Specific lipids, such as cholesterol or sphingolipids, can increase the affinity of nanodiscs for other membrane proteins or lipids [24,36]. Therefore, the careful selection of lipids is vital for successful nanodisc formation and functionality.
The concentration of the detergent sodium cholate was prepared at twice the expected concentration of lipids [4]. Thus, we used a concentration slightly above the value demanded to ensure the full solubilization of the lipids. For a final concentration of lipids at 32.9 mM, 70 mM of NaCholate was used.
Once solubilization has been achieved, ensure that all lipids are fully resuspended. The appearance of a film along the wall of the tube typically indicates that some lipids have not yet been properly resuspended.
Exercise caution when handling the supernatant, as it contains a significant amount of methanol dissolved in water.
Use cut micropipette tips and plastic pipettes at about 1/3 from the tip to prevent Bio-Beads from being blocked inside the tips and the pipettes. The loss of Bio-Beads should be limited to the dialysis membrane transfers.
Bio-Beads can stick to the dialysis membrane. To recover more than 99% of the Bio-Beads, add buffer to wash the membrane, allowing retrieving the remaining beads.
It is important to consider the below parameters before selecting the MSP proteins for experiments:
Transmembrane domains: The number of transmembrane domains in the protein is crucial for selecting the appropriate MSP protein for nanodisc reconstitution. Proteins with multiple transmembrane domains may need larger nanodiscs to accommodate the protein and allow proper interaction with the lipid bilayer. Proteins with fewer domains may require smaller nanodiscs to ensure complete protein embedding.
Nanodisc size determination: The optimal size of nanodiscs for a particular protein can be determined experimentally and may vary based on the specific protein and lipid composition [2–4]. The size of nanodiscs is defined by the amount of lipid used and the belt protein incorporated. A range of MSP constructs allows the generation of nanodiscs from 9 to 17 nm in diameter [37]. The desired size of the nanodisc governs the considerations for MSP selection; the diameter of the transmembrane region of the protein can be estimated based on the number of transmembrane domains. Longer MSPs may generate more heterogeneous nanodisc types.
Transformed bacteria are less stable and express less MSP. It may be due to the toxicity of MSP expression. To ensure efficient expression of MSPs, a new bacterial transformation should be conducted prior to each purification.
If the protein is not incorporated into nanodiscs, it may be because the nanodiscs diameter is too small to support the protein (Note 7). We suggest experimenting with the other MSPs listed, which will result in different sizes of nanodiscs.
Use Erlenmeyers flasks that allow a high rate of oxygenation with a ratio between the volume of culture and the maximum volume of at least 1:5.
At this stage, bacteria can be stored at -20 °C or below. If bacteria pellets were previously stored for a long term, the pellet will need to be thawed before resuspension.
Precipitation can eventually occur after changing the buffer.
At the end of the purification, one can aim to obtain between 5 and 10 mg of MSPs per liter of E. coli culture.
The presence of proteolyzed fragments in MSP preparations does not invalidate the successful use of those batches for nanodiscs reconstitution. While proteolytic fragments of MSPs may be present after purification, our study and others have shown that their presence does not significantly affect the reconstitution efficiency. However, it is still essential to consider the ratio percentage of full-length vs. proteolyzed fragments of MSP to ensure accurate quantification of nanodisc components and as importantly to reduce background in cryoEM images.
The volume of aliquots should be optimized depending on the concentration of proteins and the volume needed for nanodisc reconstitution to avoid unnecessary freeze-thaw cycles and degradation of the samples.
Please note that the percentage of proteolyzed MSP present in the sample will not contribute to the effective reconstitution of bare nanodiscs. Therefore, ensure that the 20 nmol amount corresponds to the full-length MSPs. We recommend adjusting the concentration based on the quantity of full-length MSP observed on the SDS-PAGE gel.
The total volume of the nanodisc preparation should not exceed 2 mL [4]. Thus, you may consider setting your initial concentrations of MSPs according to the amount and volumes needed.
To find the best experimental MSP molar ratio for your conditions, we advise testing three different ratios for the reconstitution of bare nanodiscs. For 9 nm nanodiscs using MSP1D1 protein and POPC lipids, you can start with the ratio published by Ritchie et al. (1:65) and also test 1:55 and 1:75 [4]. Perform these experiments in parallel using the same MSP and lipids stocks to compare your preparations.
The amount of Bio-Beads to use is based on their adsorption capacity, which depends on the nature of the detergents. Refer to the values in the papers by Horigome and Sugano (1983) and Rigaud and colleagues (1998) to determine the required volume [11,12]. To ensure efficient detergent removal, consider using double the calculated amount of Bio-Beads. Choosing the right detergents for solubilizing and extracting membrane proteins is crucial for effective detergent removal and stable nanodisc formation. During nanodisc reconstitution, sodium cholate is commonly used. It is important to note that Bio-Beads need to absorb both the detergent used for protein solubilization and extraction as well as the one used for nanodisc reconstitution. Thus, ensure that sodium cholate is compatible with the protein of interest.
Avoid bubbles.
Bare nanodiscs should be eluted at approximately 12 mL.
Nanodisc reconstitution depends on various factors; notably, the amount of full-length MSPs and lipids. Keeping the MSP amount constant, an excess of lipids maximizes the use of MSPs to reconstitute nanodiscs. Low lipid concentration results in fewer nanodiscs and leaves "free" MSPs in the solution. Conversely, excess lipids can form liposomes of different sizes that are not detected on SDS-PAGE.
Bare nanodiscs at a final concentration of 0.5–1 mg/mL are suitable for characterization by biophysical methods. Bare nanodisc concentration is calculated based on the measured absorbance at 280 nm, the extinction coefficient, as well as the molecular weight of one monomer of MSP.
Once the molar ratio between MSPs and lipids for bare nanodiscs is established, the same conditions can be used for initial protein integration attempts. Further optimization of the membrane protein quantity per nanodisc reconstitution may be necessary. The described purification strategy accounts for the increased hydrodynamic radius of nanodiscs incorporating the target membrane protein compared to bare nanodiscs, which will also be present in the mixture.
Buffer conditions should be optimized based on the protein's characteristics, such as pH, stability-improving additives, and salt concentration, as these can impact cryoEM experiments. We recommend starting with buffer conditions used in previous nanodiscs research studies and in our laboratories as a starting baseline [2–4,13,14,25,27].
Carefully analyze the size exclusion chromatography profile to identify contaminants such as free MSPs, liposomes of different sizes, bare nanodiscs, membrane proteins in different-size liposomes, and detergents, compared to your membrane protein nanodiscs of interest.
If your membrane protein has transmembrane domains and an ectodomain, it should elute before 12 mL under the given size exclusion chromatography conditions. If it has only transmembrane domains, its elution volume can be around 12 mL, like bare nanodiscs.
The final concentration of membrane protein nanodiscs samples that can be obtained is around 0.5–1 mg/mL, which is suitable for SDS-PAGE and western blot characterization.
These parameters need to be optimized for each protein-nanodisc preparation and plunge-freezing apparatus. Here, we provide parameters that have worked under our conditions, which can be used as a starting point.
Liquid ethane may cause cryogenic burns, frostbite, or injury and may form explosive mixtures with air. Follow all safety data sheets (SDS) before use.
The concentration of the ND solution should be between 0.17 and 0.8 mg/mL with an optimum concentration around 0.5 mg/mL.
If the Protein-ND is not visible during screening on the cryoEM session, it can be due to a low concentration of samples or the vitrified layer being too thick. It is best to look back to your ND preparation and increase ND concentration before freezing (Figure 2F, G). It is important to try several regions with different thicknesses of the vitrified layer to find the NDs. You can also use a higher defocus or Phase Plate to reenforce the contrast (Figure 2A–D).
During size exclusion chromatography, liposomes and micelles of similar size to nanodiscs may not be fully separated and can still be present in the sample. As a result, these liposomes and/or micelles can be observed during cryoEM screening. Additional purification steps, such as affinity chromatography or ultracentrifugation, may be necessary to reduce these types of contamination and enhance the purity of nanodisc preparations.
To address preferred orientations in vitrified samples, various buffers can be tested, adjusting the vitrification process itself or apparatus, or testing various grids’ types and support films, and possibly adding an extra graphene layer (e.g., a 2 nm graphene film) [38]. If everything else fails, a second tilted data collection can be performed to provide additional orientations [39,40], or sub-tomogram averaging can be considered [33].
Acknowledgments
This work was funded by institutional funds from the Institut Pasteur and the CNRS (D.H. and N.V.). D.H. acknowledges funding from the U.S. Army Research Office under contract W911NF-19-D-0001 for the Institute for Collaborative Biotechnologies award. The authors thank Nicolas Wolff, Sébastien Brûlé, and Bertrand Raynal (Institut Pasteur) for discussions about nanodiscs and membrane proteins, and for support on biochemical and biophysical experiments. The authors acknowledge the biophysical platform (PFBMI) and the NanoImaging Core (NCF) at Institut Pasteur for the provision of the equipment (DLS, Vitrobot, cryo-electron microscope). The NanoImaging Core was created with the help of a grant from the French Government’s Investissements d’Avenir program (EQUIPEX CACSICE - Centre d’analyse de systèmes complexes dans les environements complexes, ANR-11-EQPX-0008).
Competing interests
The authors declare that they have no competing interests.
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Article Information
Publication history
Received: May 29, 2024
Accepted: Sep 1, 2024
Available online: Sep 28, 2024
Published: Nov 5, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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MTT Assay of Cell Numbers after Drug/Toxin Treatment
RC Ran Chen
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Published: Apr 5, 2011
DOI: 10.21769/BioProtoc.51 Views: 25455
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Abstract
MTT assay is a colorimetric method for measuring the activity of enzymes in living cells that reduce MTT to formazan dyes, giving a purple color. It is commonly used to determine cytotoxicity of potential medicinal agents and toxic materials, since these types of materials are expected to stimulate or inhibit cell viability and growth. Here, a general protocol is described to carry out an MTT assay on different types of cells.
Keywords: MTT assay Cell numbers Drug
Materials and Reagents
Raw264.7, MCF-7 or Hela cells
Thiazolyl blue tetrazolium bromide (MTT) (Sigma-Aldrich, catalog number: M5655 )
DMSO
DPBS (Life Technologies, InvitrogenTM, catalog number: 14190-250 )
General chemicals (Sigma-Aldrich)
Equipment
96 well plate
Shaking table
Paper towels
Incubator
Procedure
Plate 500-10,000 cells in 200 μl media per well in a 96 well plate. Leave 8 wells empty for blank controls.
Incubate (37 °C, 5% CO2) overnight to allow the cells to attach to the wells.
Add 2 μl of drug of interest dissolved in DMSO to each well. Place on a shaking table, 150 rpm for 5 min, to thoroughly mix the samples into the media.
Incubate (37 °C, 5% CO2) for 1-5 days to allow the drug/toxin to take effect.
Make 2 ml or more of MTT solution per 96 well plate at 5 mg/ml in DPBS. Do not make a stock as MTT in solution is not stable long-term.
Add 20 μl MTT solution to each well. Place on a shaking table, 150 rpm for 5 min, to thoroughly mix the MTT into the media.
Incubate (37 °C, 5% CO2) for 1-5 h to allow the MTT to be metabolized.
Dump off the media (dry plate on paper towels to remove residue if necessary).
Resuspend formazan (MTT metabolic product) in 200 μl DMSO. Place on a shaking table, 150 rpm for 5 min, to thoroughly mix the formazan into the solvent.
Read optical density at 560 nm and subtract background at 670 nm. Optical density should be directly correlated with cell quantity.
Acknowledgments
This work was funded by 5050 project by Hangzhou Hi-Tech District, Funding for Oversea Returnee by Hangzhou City, ZJ1000 project by Zhejiang Province. This protocol was developed in the Cohen Lab, Department of Genetics, Stanford University, CA, USA [Chen et al. (unpublished)].
References
Hayon, T., Dvilansky, A., Shpilberg, O. and Nathan, I. (2003). Appraisal of the MTT-based assay as a useful tool for predicting drug chemosensitivity in leukemia. Leuk Lymphoma 44(11): 1957-1962.
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An In Vitro Model of Murine Osteoclast-Mediated Bone Resorption
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YT Yi Tang
SW Stephen J. Weiss
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Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5100 Views: 400
Reviewed by: Valérian DORMOYJaira Ferreira de VasconcellosNona Farbehi
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The authors used this protocol in The Journal of Cell Biology Apr 2023
Abstract
Osteoclasts are terminally differentiated multinucleated giant cells that mediate bone resorption and regulate skeletal homeostasis under physiological and pathological states. Excessive osteoclast activity will give rise to enhanced bone resorption, being responsible for a wide range of metabolic skeletal diseases, ranging from osteoporosis and rheumatoid arthritis to tumor-induced osteolysis. Therefore, the construction of in vitro models of osteoclast-mediated bone resorption is helpful to better understand the functional status of osteoclasts under (patho)physiological conditions. Notably, it is essential to provide an in vivo–relevant bone substrate that induces osteoclasts to generate authentic resorption lacunae and excavate bone. Here, we summarize the experimental design of a reproducible and cost-effective method, which is suitable for evaluating the regulatory mechanisms and influence of molecular agonists and antagonists as well as therapeutics on osteoclast-mediated bone-resorbing activity.
Key features
• Experiments are performed using bovine cortical bone slices to simulate bone substrate resorption by murine osteoclasts in vivo.
• The method allows for quantification of bone resorption in vitro.
• The method is suitable for evaluating the regulatory mechanisms that control osteoclast-mediated bone-resorbing activity.
Keywords: Osteoclast Bone resorption Bone slices Resorption pits Cell culture
Graphical overview
Schematic diagram of the protocol for assessing osteoclast-mediated bone-resorbing activity in vitro. Step 1: Euthanizing 8–12-week-old male mice to separate tibias and femurs. Step 2: Flushing bone marrow cells by injector and obtaining the cells. Step 3: Preparation of BMDMs with complete medium containing 20 ng/mL M-CSF in 10 cm Petri dishes. Step 4: Cultivation of osteoclasts with complete medium containing 20 ng/mL M-CSF and 30 ng/mL RANKL atop bovine cortical bone slices in a 96-well plate. Step 5: Evaluating bone-resorbing activity of osteoclasts.
Background
Osteoclasts are exclusive and specialized multinucleated bone-resorbing cells derived from both embryonic and hematopoietic stem cell precursors of erythromyeloid and myeloid lineages [1]. An imbalance of skeletal remodeling as a consequence of increased osteoclast-mediated bone resorption is responsible for a wide range of metabolic skeletal diseases, ranging from osteoporosis and rheumatoid arthritis to tumor-induced osteolysis. Therefore, the construction of in vitro osteoclast-mediated bone resorption models is helpful to better characterize the functional status of osteoclasts under both physiological and pathological conditions.
The transition of cells of the monocyte/macrophage lineage into osteoclasts in vitro requires two essential cytokines—macrophage colony-stimulating factor (M-CSF) and receptor activator of NF-kB ligand (RANKL). The presence of M-CSF promotes the proliferation of osteoclast precursor cells, and RANKL further induces precursor cells to express an osteoclast phenotype [2]. Mature osteoclasts at the bone surfaces undergo a polarization process marked by extensive morphologic changes, including the formation of an actin ring at their basal surface, termed the sealing zone. The sealing zone surrounds a differentiated and intricately folded region of the plasma membrane called the ruffled border, where protons, chloride ions, and proteases are secreted into the underlying resorption lacunae where bone demineralization and extracellular matrix degradation proceed [3]. As the organic phase of bone is dominated by type I collagen, the proteolytic release of C-terminal telopeptide fragments of type collagen (CTX-1) is a characteristic biochemical marker of osteoclast-mediated bone resorption [4]. When cultured on bone or dentin substrate, osteoclasts assemble resorption lacunae, thereby generating resorption pit structures similar to those formed during bone resorption in vivo [5]. Notably, bovine cortical bone slices are superior to synthetic materials or dentine-derived matrix as they better mimic bone substrates encountered in vivo [6]. When combined with an appropriate detection assay, osteoclast-mediated bone-resorbing activity can be quantified under a variety of experimental conditions [7,8].
Here, we describe a step-by-step protocol for in vitro bone-resorbing activity using an osteoclast-cortical bone slice co-culture system. This protocol consists of three main steps: mouse bone marrow–derived macrophage (BMDM) preparation, osteoclast induction and characterization, and bone resorption pit quantification. This method can be universally utilized to evaluate the bone-resorbing activity of osteoclasts in response to both agonists and antagonists.
Materials and reagents
Biological materials
8–12-week-old male C57BL/6J mice (The Jackson Laboratory, catalog number: 000664)
Bovine cortical bone slices (Immunodiagnostic Systems, catalog number: DT-1BON1000-96)
Reagents
Alpha minimum essential medium (α-MEM) (Hyclone, Cytiva, catalog number: SH30265.01)
Fetal bovine serum (FBS) (Sigma-Aldrich, catalog number: F0193)
Penicillin-streptomycin solution (Hyclone, Cytiva, catalog number: SV30010)
Red blood cell lysing buffer (Biosharp, catalog number: BL504A)
Phosphate buffered saline (PBS) (Servicebio, catalog number: G0002)
Ethylenediaminetetraacetic acid (EDTA) solution, 0.5 M (Sigma-Aldrich, catalog number: 03690)
Recombinant murine M-CSF (R&D Systems, catalog number: 416-ML)
Recombinant murine RANKL (R&D Systems, catalog number: 462-TEC)
Bovine serum albumin (Sigma-Aldrich, catalog number: A1933)
Paraformaldehyde powder (Sigma-Aldrich, catalog number: 158127)
Glutaraldehyde solution, 50% (Sigma-Aldrich, catalog number: 49629)
Tartrate-resistant acid phosphatase (TRAP) stain kit (Sigma-Aldrich, catalog number: 387A)
Triton X-100 solution, 10% (Sigma-Aldrich, catalog number: 49629)
Phalloidin-tetramethylrhodamine B isothiocyanate (Sigma-Aldrich, catalog number: P1951)
α-Tubulin mouse mAb (Cell Signaling Technology, catalog number: 3873)
Donkey anti-rat Alexa 488-conjugated secondary antibodies (Invitrogen Molecular Probes, catalog number: A-21202)
Sodium hydroxide (NaOH) powder (Sigma-Aldrich, catalog number: 655104)
Peroxidase-conjugated wheat germ agglutinin (WGA) (Sigma-Aldrich, catalog number: L3892)
Diaminobenzidine (DAB) tablets (Sigma-Aldrich, catalog number: D4418)
FITC conjugate WGA (Sigma-Aldrich, catalog number: L4895)
CrossLaps ELISA kit for quantifying CTX-1 release (Immunodiagnostic Systems, catalog number: AC-07F1)
Solutions
5 mM EDTA solution (see Recipes)
0.1% Bovine serum albumin solution (see Recipes)
Murine recombinant M-CSF solution (see Recipes)
Murine recombinant RANKL solution (see Recipes)
4% Paraformaldehyde solution (see Recipes)
2.5% Glutaraldehyde solution (see Recipes)
NaOH solution (see Recipes)
0.1% Triton X-100 solution (see Recipes)
Peroxidase-conjugated WGA solution (see Recipes)
FITC-conjugated WGA solution (see Recipes)
Recipes
5 mM EDTA solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
0.5 M EDTA solution 5 mM 0.5 mL Room temperature
Sterile PBS NA 49.5 mL Room temperature
Final volume 50 mL 4 1 week
0.1% Bovine serum albumin solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
Bovine serum albumin 0.1% 0.02 g 4
Double-distilled water NA 20 mL Room temperature
Final volume 20 mL -20 1 month
Murine rM-CSF solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
Murine rM-CSF 50 μg/mL 50 μg -80
Sterile PBS containing 0.1% bovine serum albumin NA 1 mL -20
Final volume 1 mL -80 1 year
Murine rRANKL solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
Murine rRANKL 50 μg/mL 50 μg -80
Sterile PBS containing 0.1% bovine serum albumin NA 1 mL -20
Final volume 1 mL -80 1 year
4% Paraformaldehyde solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
Paraformaldehyde powder 4% (w/v) 0.4 g -4
Sterile PBS NA 10 mL Room temperature
Final volume 10 mL -4 1 week
2.5% Glutaraldehyde solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
50% glutaraldehyde solution 2.5% (v/v) 0.5 mL Room temperature
Sterile PBS NA 9.5 mL Room temperature
Final volume 10 mL -4 1 week
NaOH solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
NaOH powder 0.5 N 1 g Room temperature
Double-distilled water NA 50 mL Room temperature
Final volume 50 mL -4 1 month
0.1% Triton X-100 solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
10% Triton X-100 solution 0.1% 0.01 mL Room temperature
Double-distilled water NA 9.99 mL Room temperature
Final volume 10 mL -4 1 month
Peroxidase-conjugated WGA solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
Peroxidase-conjugated WGA 2 mg/mL 1 mg -20
Sterile PBS NA 0.5 mL Room temperature
Final volume 0.5 mL -20 1 year
FITC-conjugated WGA solution
Reagent Final concentration Quantity or Volume Recommended storage condition Recommended storage time
FITC-conjugated WGA 2 mg/mL 1 mg -20
Sterile PBS NA 0.5 mL Room temperature
Final volume 0.5 mL -20 1 year
Laboratory supplies
10 mL syringe with 25 G needle
Sterile absorbent gauze
70 µm cell strainer (Corning, catalog number: 431751)
Pipette
15 mL conical tube (Nest, catalog number: 601001)
50 mL conical tube (Nest, catalog number: 601002)
10 × 10 cm Petri dish with cover (Nest, catalog number: 752001)
Cell lifter (Corning, catalog number: CLS3008)
96-well cell culture plates (Nest, catalog number: 713011)
Equipment
Dissecting tweezers and scissors (Autoclaved)
Cell counter (Thermo Fisher Scientific Inc, model: Countess 3)
Cell culture incubator (37 °C, 5% CO2)
Centrifuge (Eppendorf)
Inverted microscope (Olympus)
Confocal laser scanning microscope (CLSM) (Nikon A1)
Critical point drying (Balzers Union)
Au/Pg sputtered (Polaron Equipment Ltd, model: E-5100)
Field emission scanning electron microscopy (FE-SEM) (AMRAY 1910)
Software and datasets
ImageJ is a Java-based (runs on Mac OS X, Linux, and Windows) freeware available for download at: http://rsb.info.nih.gov/ij/
GraphPad Prism 7
Procedure
Preparation of bone marrow-derived macrophages (BMDMs)
Euthanize 8–12-week-old male C57BL/6J mice by carbon dioxide inhalation and soak in 50 mL of 75% ethanol for 5 min.
Remove the mouse hind limbs intact and bilaterally along the greater trochanter of the femur using sterile scissors and forceps. Remove excess skin and muscle tissues from the femur and tibia with sterile gauze and then place the bones in 10 mL of pre-prepared PBS solution at 4 containing 2% penicillin-streptomycin.
Separate tibiae and femurs using sterile scissors and then wash twice with 2 mL of PBS solution containing 2% penicillin-streptomycin for 5 min. Figure 1B shows the bones and both ends of each bone type cut to expose the bone marrow cavity.
Figure 1. Overview of osteoclastogenesis and bone-resorbing assay. (A) Schematic diagram of separating tibiae and femurs and removing the ends of the bones. (B) Separating the tibia and femur. (C) Irrigating bone marrow cavity. (D) Filtering through a cell strainer. (E) Preparing bovine cortical bone slices. (F) Culturing osteoclasts atop bone slice in a 96-well plate. (G) Preparing for observation of bone resorption. (H) ELISA analysis of the cultured supernatants.
Flush bone marrow cells from both ends of the bone with α-MEM using a 10 mL syringe with a 25 G needle and then pass them through a 70 μm cell strainer into a 50 mL conical tube (Figure 1C, D). Rinse repeatedly until there is no visible redness remaining in the bone marrow cavity. Use approximately 8 mL of α-MEM per bone to flush cells and temporarily store all bone marrow cells at room temperature in 50 mL conical tubes.
Pellet bone marrow cells by centrifuging at 400× g for 5 min at room temperature and discard the supernatant. Resuspend cells thoroughly with 3 mL of red blood cell lysing buffer and then let them stand for 2–3 min at room temperature to lyse red blood cells.
Add 5 mL of room-temperature α-MEM medium supplemented with 10% FBS to terminate lysis, pellet cells by centrifuging at 300× g for 3 min at room temperature, and discard the supernatant.
Resuspend cells in the 20 mL prepared complete medium containing 20 ng/mL rM-CSF and then transfer cells into two 10 cm Petri dishes for overnight incubation in 5% CO2/95% air atmosphere at 37 °C.
After 12 h, collect non-adherent cells and transfer them to fresh α-MEM medium containing 10% FBS and 20 ng/mL rM-CSF in six to eight 10 cm Petri dishes for further expansion of the cell population (Figure 2A). The expected yield of non-adherent cells is approximately 5 × 106 cells/mL.
Change medium every other day for 4–5 days until the adherent cells reach 80%–90% confluence (Figure 2B).
After rinsing twice with PBS, add 5 mL of a 5 mM EDTA solution to each Petri dish. After incubation in 5% CO2/95% air atmosphere at 37 °C for 5 min, detach adherent cells using a cell lifter.
When most cells are observed to be suspended under the microscope, cell detachment is terminated by adding 5 mL per dish of α-MEM medium containing 10% FBS. After centrifugation of cells in 15 mL conical tubes at 400× g for 5 min, resuspend the cells with α-MEM medium with 10% FBS containing 20 ng/mL rM-CSF.
Note: Cells should be used immediately for culture and not prepared for cryopreservation.
Induction and characterization of osteoclast
Disinfect bovine cortical bone slices with 75% ethanol for 24 h and then wash twice with PBS solution for 5 min (Figure 1E). Place bone slices in a 96-well plate (one slice per well) with 200 μL of α-MEM medium containing 10% FBS.
Harvest bone marrow cells and culture atop individual bone slices in the presence of 20 ng/mL rM-CSF at a final cell concentration of 6 × 104–8 × 104 cells/mL in a final volume of 200 μL/well (Figure 1F).
After 12 h, change the medium to α-MEM medium with 10% FBS containing 20 ng/ml rM-CSF and 30 ng/ml rRANKL. Then, change full medium every other day for 6 days to induce osteoclast formation (days 4–5) and bone resorption pits (days 5–6).
Note: If the specific factors or compounds are of interest, add them to the cell culture medium as well. Due to the opacity of the bone slices, it is difficult to observe the state of the cells during this incubation period (Figure 2C); the reference times we have provided are suitable for strict adherence to our protocols.
Figure 2. State of cells at different stages under the light microscope. (A) Non-adherent cells (step A8). (B) Adherent cells reaching 80%–90% confluence (step A9). (C) Culturing osteoclasts atop bone slice (step B3).
Fix bone slices with 4% paraformaldehyde or 2.5% glutaraldehyde in PBS solution for 30 min at room temperature. Transfer the cultured medium to a centrifuge tube, centrifuge at 400× g for 10 min at 4 °C, and then collect the supernatant.
Use the TRAP stain kit to label cells on a 96-well plate according to the manufacturer’s instructions. Then, examine microscopically to observe TRAP-positive osteoclasts. TRAP-positive cells, which are dark red to purple, are identified as osteoclasts if they have three or more nuclei (Figure 3A).
For F-actin staining, fix osteoclasts on bone slices with 4% paraformaldehyde, permeabilize with 200 μL of 0.1% Triton X-100 for 10 min, and block with 2.5% bovine serum albumin for 1 h prior to an overnight incubation at 4 °C with primary antibodies direct against α-Tubulin (1:1,000). Following primary antibody incubations, incubate osteoclasts with donkey anti-mouse Alexa 488-conjugated secondary antibodies for 1 h and subsequently with phalloidin-tetramethylrhodamine B isothiocyanate for 30 min at 37 °C (Figure 3B).
For scanning electron microscopy (SEM) analysis, process osteoclasts atop bone slices for critical point drying and Au/Pg sputter; then, image them on a FE-SEM (Figure 3B).
Quantification of bone resorption pits
After incubating bone slices in 0.5 N NaOH for 30 s, gently scrape off cells using a medical cotton swab.
Label bone slices on the opposite side without cells to serve as a marker for the cell-containing side to observe. Dehydrate samples in ascending ethanol series (70%, 80%, 90%, 95%, 100%) at room temperature.
For SEM analysis, process bone slices for critical point drying and Au/Pg sputter. The resorption pits are imaged on a FE-SEM to clearly observe their morphology (Figure 3C).
For WGA-DAB analysis, soak bone samples in PBS for 5 min, stain with 100 μL of 20 μg/mL Peroxidase-WGA for 45 min, and then incubate with DAB tablets for 15 min. Picture bone resorption pits, which are stained in brown using a light microscope (Figure 3C), and determine the resorbed area in three random sites in one bone slice using ImageJ software. Access the irregular graphics box to outline resorption pits (stained brown) and then use the Analyze and Measure function to determine the resorption area.
For FITC-WGA analysis, sonicate bone samples in PBS and stain with 100 μL of 20 μg/mL FITC-WGA for 45 min in darkness (Figure 1G). Use a confocal laser scanning microscope to characterize three-dimensional profiles of resorption pits (Figure 3C) and process to obtain images of the sagittal angle. Quantitative analysis of resorption pit depth is performed in three random sites per bone slice using ImageJ software. Use ImageJ software to open the image of the sagittal angle, mark the depth of the bone resorption pits with the straight-line tool, and then click on Analyse and Measure to get the straight-line length.
Detect the supernatant from the cultured medium by CrossLaps ELISA kit for CTX-1 according to the manufacturer’s instructions to measure the concentration of collagen fragments (Figure 1H).
Data analysis
Figure 3 shows the characterization of osteoclast morphology and bone resorption pits.
Figure 3. Osteoclasts were induced on bone slices and formed bone resorption lacunae. (A) Bone marrow–derived macrophages (BMDMs) were then harvested and cultured for 4–5 days with rM-CSF (20 ng/mL) and rRANKL (30 ng/mL) on plastic substrata. Following TRAP staining, large multinucleated cells (MNCs) were formed (scale bar = 50 μm). Black arrows indicate large multinucleated cells. (B) BMDMs were cultured atop bovine bone slices in the presence of rM-CSF and rRANKL for 6 days. SEM analysis shows bone adherent osteoclasts. Sealing zone formation is assessed by phalloidin staining to visualize F-actin rings, while microtubules and nuclei are identified by staining with alpha-tubulin and DAPI, respectively. Scale bar = 10 μm, 10 μm, and 2 μm, from left to right panels). (C) Resorption pits were visualized by wheat germ agglutinin–diaminobenzidine (WGA-DAB) staining, scanning electron microscopy (SEM), and 3D imaging of WGA-FITC staining by confocal microscopy (scale bar = 100 μm, 10 μm, and 100 μm, from left to right panels). Black and white arrows indicate bone resorption pits.
Validation of protocol
This protocol has been used and validated in the following research article:
Zhu et al. [9]. Osteoclast-mediated bone resorption is controlled by a compensatory network of secreted and membrane-tethered metalloproteinases. Sci Transl Med. (Figures 2 and 3)
Zhu et al. [10]. A Zeb1/MtCK1 metabolic axis controls osteoclast activation and skeletal remodeling. EMBO J. (Figure 3)
Zhu et al. [11]. Proteolytic regulation of a galectin-3/Lrp1 axis controls osteoclast-mediated bone resorption. J Cell Biol. (Figures 2 and 5)
Ng et al. [12]. Sugar transporter Slc37a2 regulates bone metabolism in mice via a tubular lysosomal network in osteoclasts. Nat Commun. (Figure 8)
Li et al. [13]. Dynamic changes in O-GlcNAcylation regulate osteoclast differentiation and bone loss via nucleoporin 153. Bone Res. (Figures 2 and 3)
General notes and troubleshooting
Problem 1: BMDMs were unable to attach and culture atop bone slices.
Possible cause: Bone slices were not activated by the cultured medium.
Solution: Before BMDMs are inoculated, bone slices need to be soaked in the medium for 2 h.
Problem 2: BMDMs were unable to fuse to form osteoclasts.
Possible cause: Our recommended cell density is 6 × 104–8 × 104 cells/mL, 200 μL each well. The cell density is much higher than conventional RANKL-induced osteoclast in 96-well plates.
Solution: Use our recommended cell densities of BMDMs when inoculating cells on bone slices, since an appropriate density of BMDMs contributes to their osteoclastogenesis and fusion.
Problem 3: There was no F-actin ring formation of osteoclasts.
Possible cause: Osteoclast bone resorption function was not activated.
Solution: Before final fixation of osteoclasts, fresh rRANKL should be stimulated for 12–14 h.
Problem 4: There were few resorption pits on the bone slices despite sufficient osteoclast numbers observed.
Possible cause: Osteoclast and cell debris were not successfully removed.
Solution: Incubate bone slices in 0.5 N NaOH for 30 s and gently scrape off the cells using a medical cotton swab.
Problem 5: No cells or resorption pits were observed on the bone slices.
Possible cause: The cells were cultured on the other side of the bone surface.
Solution: We recommend that the bone slices be marked with a pencil on the surface devoid of cells after fixation.
Acknowledgments
This work was supported by the NSFC 82370914 and 81970919, the Fundamental Research Funds for the Central Universities 2042024YXA010 (L.Z.), the NSFC 82201042 and the Natural Science Foundation of Hubei Province 2022CFB658 (X.S.), the R01-AR075168 from the NIH (S.J.W.)
Competing interests
The authors declare no conflict of interest.
Ethical considerations
All animal experiments were approved by the Animal Research Ethics Committee of Wuhan University, China.
References
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Udagawa, N., Takahashi, N., Akatsu, T., Tanaka, H., Sasaki, T., Nishihara, T., Koga, T., Martin, T. J. and Suda, T. (1990). Origin of osteoclasts: mature monocytes and macrophages are capable of differentiating into osteoclasts under a suitable microenvironment prepared by bone marrow-derived stromal cells. Proc Natl Acad Sci USA. 87(18): 7260–7264.
Uehara, S., Udagawa, N., Mukai, H., Ishihara, A., Maeda, K., Yamashita, T., Murakami, K., Nishita, M., Nakamura, T., Kato, S., et al. (2017). Protein kinase N3 promotes bone resorption by osteoclasts in response to Wnt5a-Ror2 signaling. Sci Signaling. 10(494): eaan0023.
Zhang, Y., Rohatgi, N., Veis, D. J., Schilling, J., Teitelbaum, S. L. and Zou, W. (2018). PGC1β Organizes the Osteoclast Cytoskeleton by Mitochondrial Biogenesis and Activation. J Bone Miner Res. 33(6): 1114–1125.
Inoue, K., Deng, Z., Chen, Y., Giannopoulou, E., Xu, R., Gong, S., Greenblatt, M. B., Mangala, L. S., Lopez-Berestein, G., Kirsch, D. G., et al. (2018). Bone protection by inhibition of microRNA-182. Nat Commun. 9(1): 1114–1125.
Nishi, Y., Atley, L., Eyre, D. E., Edelson, J. G., Superti-Furga, A., Yasuda, T., Desnick, R. J. and Gelb, B. D. (1999). Determination of Bone Markers in Pycnodysostosis: Effects of Cathepsin K Deficiency on Bone Matrix Degradation. J Bone Miner Res. 14(11): 1902–1908.
Rumpler, M., Würger, T., Roschger, P., Zwettler, E., Sturmlechner, I., Altmann, P., Fratzl, P., Rogers, M. J. and Klaushofer, K. (2013). Osteoclasts on Bone and Dentin In Vitro: Mechanism of Trail Formation and Comparison of Resorption Behavior. Calcif Tissue Int. 93(6): 526–539.
Shemesh, M., Addadi, S., Milstein, Y., Geiger, B. and Addadi, L. (2015). Study of Osteoclast Adhesion to Cortical Bone Surfaces: A Correlative Microscopy Approach for Concomitant Imaging of Cellular Dynamics and Surface Modifications. ACS Appl Mater Interfaces. 8(24): 14932–14943.
Zhu, L., Tang, Y., Li, X. Y., Keller, E. T., Yang, J., Cho, J. S., Feinberg, T. Y. and Weiss, S. J. (2020). Osteoclast-mediated bone resorption is controlled by a compensatory network of secreted and membrane-tethered metalloproteinases. Sci Transl Med. 12(529): eaaw6143.
Zhu, L., Tang, Y., Li, X., Kerk, S. A., Lyssiotis, C. A., Feng, W., Sun, X., Hespe, G. E., Wang, Z., Stemmler, M. P., et al. (2023). A Zeb1/MtCK1 metabolic axis controls osteoclast activation and skeletal remodeling. EMBO J. 42(7): e2022111148.
Zhu, L., Tang, Y., Li, X. Y., Kerk, S. A., Lyssiotis, C. A., Sun, X., Wang, Z., Cho, J. S., Ma, J., Weiss, S. J., et al. (2023). Proteolytic regulation of a galectin-3/Lrp1 axis controls osteoclast-mediated bone resorption. J Cell Biol. 222(4): e202206121.
Ng, P. Y., Ribet, A. B. P., Guo, Q., Mullin, B. H., Tan, J. W. Y., Landao-Bassonga, E., Stephens, S., Chen, K., Yuan, J., Abudulai, L., et al. (2023). Sugar transporter Slc37a2 regulates bone metabolism in mice via a tubular lysosomal network in osteoclasts. Nat Commun. 14(1): 906.
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Article Information
Publication history
Received: Jun 20, 2024
Accepted: Sep 3, 2024
Available online: Oct 13, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Developmental Biology > Cell growth and fate > Differentiation
Cell Biology > Cell isolation and culture > Cell differentiation
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Peer-reviewed
Novel Cross-Species Salivary Gland-Parasympathetic Neuron Coculture System
HW Hsueh-Fu Wu *
MI Mohamed Ishan *
MR Md Mamunur Rashid
HL Hong-Xiang Liu *
NZ Nadja Zeltner *
(*contributed equally to this work)
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5101 Views: 292
Reviewed by: Xiaokang WuKrishna Murthy Nakuluri Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Cell Stem Cell May 2024
Abstract
The parasympathetic nervous system is essential for salivary gland development and functionality. Parasympathetic neuron (parasymN) innervation is the main neural network that controls salivary secretion. Therefore, an exclusive model to study parasympathetic neurons and salivary gland tissue circuitry will significantly improve the understanding of the role of parasymN activation on salivary regulation. Harvesting primary rodent parasymNs is challenging due to their body-wide disbursed location. Similarly, the salivary glands are distributed in various locations around and within the oral cavity. Here, we present a coculture model system using human pluripotent stem cell (hPSC)-derived parasymNs and primary mouse von Ebner’s gland cells. We previously reported the first protocol to robustly generate human parasymNs from hPSCs through the Schwann cell precursor (SCP) lineage. The hPSC-parasymNs are functional and have been applied to model several autonomic disorders. We also used a Sox10-Cre::tdTomato (hereafter referred to as RFP) reporter mouse line, which labeled von Ebner’s glands, a type of minor salivary gland connected to the trough of circumvallate and foliate taste papillae. This labeling allowed for visualization and efficient isolation of primary tissues in young adult mice (8–10 weeks). By coculturing the two tissues, human parasymNs control mouse salivary gland cell growth and activation. Both parasymNs and primary salivary gland cells can be frozen and stocked at early stages of differentiation and isolation, making applications easier. This novel coculture model system could also be used to model and study related human diseases in the future, such as dry mouth syndrome.
Key features
• Differentiation of human parasymNs from hPSCs.
• Dissecting RFP+ mouse Ebner’s gland tissues for primary culture.
• Protocol to coculture human parasymNs with mouse primary salivary gland cells.
• Allows developmental and functional assessments of salivary regulation by the parasympathetic nervous system.
Keywords: Parasympathetic nervous system Human pluripotent stem cells Salivary gland von Ebner’s gland Functional parasympathetic salivary gland coculture
Graphical overview
Background
The autonomic nervous system (ANS) regulates involuntary biological functions, such as blood pressure, heartbeat, and gland secretion. The ANS is subdivided into the sympathetic and parasympathetic nervous systems (SNS and PSNS). While sympathetic neurons (symNs) trigger the fight-or-flight response that increases the heartbeat, parasymNs trigger the rest-and-digest response to calm down the body, decreasing the heartbeat [1]. Clinically, the SNS usually draws more attention in autonomic dysregulation, because its dysfunction typically drives more dramatic and detrimental responses and symptoms [2,3]. However, innervation by parasymNs drives important functions, including salivation of the salivary glands [4]. Accordingly, PSNS activation induces elevated calcium (Ca2+) flux in salivary gland cells, which leads to increased saliva production [5].
Harvesting large numbers of primary parasymNs from rodents and humans for multiple assays or high-throughput screening is challenging. The hPSC technology can bridge this problem. hPSC-derived cells provide an unlimited source of any human cell type as desired. This is, however, highly dependent on good in vitro differentiation protocols. To date, there are several well-characterized symN differentiation protocols, including ours [6–11]. In contrast, only two parasymN protocols have been reported so far, and only our protocol generates functional hPSC-parasymNs that are derived via the proper developmental process, i.e., differentiation from neural crest cell (NCC)-derived SCPs [12]. In this protocol, the NCC fate was first induced from hPSCs. This was followed by SCP differentiation in 3D spheroid culture. Finally, SCP spheroids were replated to generate parasymNs [12]. To make research applications easier and secure reproducibility and efficiency, we showed that the protocol can be reproduced using frozen NCC stocks [12]. hPSC-parasymNs express specific genetic markers, including early autonomic markers ASCL1/PHOX2B, later cholinergic markers CHAT/VACHT/CHT/CHRM2/CHRM4, and specific parasymN markers HMX2/3[12,13]. We confirmed the electrical activity of hPSC-parasymNs, and electric activity could be manipulated via nicotine, which mimics preganglionic signaling. In cocultures with cardiomyocytes, nicotine activation downregulated their beating rate [12]. Using hPSC-parasymNs, we studied ANS responsiveness to COVID-19 infection and modeled the genetic autonomic disorder familial dysautonomia [12].
Salivary secretion, important for wetting oral surfaces and maintaining taste acuity, is dependent on the activity of innervating nerves to the salivary glands. Salivary glands include three pairs of major (parotid, submandibular, and sublingual) and numerous minor salivary glands around the mouth cavity. The von Ebner’s glands are a type of minor salivary glands with the ducts open to the trench of circumvallate and foliate taste papillae. Their connection to the taste papillae makes it easy to recognize and dissect. They are innervated by both sympathetic and parasympathetic efferent nerves, yet the salivary fluid secretion (water mobilizing) is largely controlled by parasympathetic activities [14].
Here, we describe a protocol to coculture hPSC-parasymNs with RFP+ mouse primary von Ebner’s gland cells [15,16]. We show that the proliferation of von Ebner’s gland cells is promoted when parasymNs are present. Furthermore, the number of auto-fluorescent secretory granule-positive von Ebner’s gland cells is significantly increased in the cocultures, suggesting the increased salivary cell maturation due to parasymN innervation. We also measured Ca2+ flux to represent salivary production, which is increased after parasymN stimulation in the cocultures. Impaired salivation can be seen in multiple diseases, such as Parkinson’s disease, Wilson disease, Angelman syndrome, various infections, and drug abuse [4]. Our newly established model may therefore be used to study the defects and cytotoxicity in those conditions, as well as the regenerative potential of hPSC-parasymNs on damaged salivary glands in the future.
Materials and reagents
Biological materials
Human embryonic stem cell line (WiCell, WA09, female, NIH 0062)
Sox10-Cre mice (Jackson Laboratory, stock 025807)
RFP Cre reporter mice B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J (Jackson Laboratory, stock 007914)
Reagents
Geltrex (Invitrogen, catalog number: A1413202)
Poly-L-ornithine hydrobromide (PO) (Sigma, catalog number: P3655)
Mouse laminin I (LM) (R&D Systems, catalog number: 3400-010-01)
Human fibronectin (FN) (VWR/Corning, catalog number: 47743-654)
Essential 8 medium (E8) (Gibco, catalog number: A15169-01)
Essential 8 supplement (E8) (Gibco, catalog number: A15171-01)
Essential 6 medium (E6) (Gibco, catalog number: A15165-01)
Neurobasal media (Gibco, catalog number: 21103-049)
Stem cell banker (Amsbio, catalog number: 11924)
DMEM/F12 (Thermo Fisher/Life Technologies, catalog number: 11330-057)
DMSO (Thermo Fisher/Life Technologies, catalog number: BP231-100)
Fetal bovine serum (FBS) (Atlanta Biologicals, catalog number: S11150)
B27 supplement (Thermo Fisher/Life Technologies, catalog number: 12587-010)
N2 supplement (Thermo Fisher/Life Technologies, catalog number: 17502-048)
L-Glutamine (Thermo Fisher/Gibco, catalog number: 25030-081)
GlutaMAX (Gibco, catalog number: 35050061)
Antibiotic-antimycotic (Fisher, catalog number: 15 240 096)
Accutase (Innovation Cell Technologies, catalog number: AT104500)
EDTA (Sigma, catalog number: ED2SS)
Trypsin-EDTA (Gibco, catalog number: 25200056)
Phosphate-buffered saline (PBS) (Gibco, catalog number: 14190-136)
SB431542 (Tocris/R&D Systems, catalog number:1614), make 10 mM stock
BMP4 (R&D Systems, catalog number: 314-BP), make 10 µg/mL stock
CHIR99021 (R&D Systems, catalog number: 4423), make 6 mM stock
Y27632 (R&D Systems, catalog number: 1254), make 10 mM stock
FGF2 (R&D Systems, catalog number: 233-FB/CF), make 10 µg/mL stock
NRG1 (PeproTech, catalog number: 100-03), make 100 µg/mL stock
GDNF (PeproTech, catalog number: 450), make 10 µg/mL stock
BDNF (R&D Systems, catalog number: 248-BD), make 10 µg/mL stock
CNTF (R&D Systems, catalog number: 257-NT), make 100 µg/mL stock
Ascorbic acid (Sigma, catalog number: A8960), make 100 mM stock
dbcAMP (Sigma, catalog number: D0627), make 100 mM stock
Retinoic acid (Sigma, catalog number: R2625), make 1 mM stock
Collagenase A (Tribioscience, catalog number: TBS2116-01), make 2 mg/mL stock
Dispase II (Tribioscience, catalog number: TBS2117-01), make 5 mg/mL stock
DPBS without Ca+2 and Mg+2 (Thermo Fisher/Life Technologies, catalog number: 14190144)
70% ethanol
Solutions
NCC induction medium, D0, 1 (see Recipes)
NCC induction medium, D2-10 (see Recipes)
SCP differentiation medium, D10-16 (see Recipes)
ParasymN differentiation medium, D16 (see Recipes)
Salivary gland cell medium (see Recipes)
Tongue tissue separation solution (see Recipes)
Salivary gland cell dissociation solution (see Recipes)
Dissociation termination solution (see Recipes)
Recipes
NCC induction medium, D0, 1 (50 mL)
Reagent Final concentration Quantity or Volume
E6 n/a 50 mL
SB431542 (10 mM stock) 10 µM 50 µL
BMP4 (10 µg/mL stock) 0.2–1 ng/mL 1–5 µL
CHIR99021 (6 mM stock) 300 nM 2.5 µL
Y27632 (10 mM stock) 10 µM 50 µL
Note: Please see Troubleshooting 1 for more information.
NCC induction medium, D2-10 (100 mL)
Reagent Final concentration Quantity or Volume
E6 n/a 100 mL
SB431542 10 µM 100 µL
CHIR99021 0.75 µM 12.53 µL
SCP differentiation medium, D10-16 (100 mL)
Reagent Final concentration Quantity or Volume
Neurobasal media n/a 100 mL
N2 supplement 1% 1 mL
B27 supplement 2% 2 mL
L-glutamine 1% 1 mL
CHIR99021 3 µM 50 µL
FGF2 (10 µg/mL stock) 10 ng/mL 100 µL
NRG1 (100 µg/mL stock) 10 ng/mL 10 µL
ParasymN differentiation medium, D16 (100 mL)
Reagent Final concentration Quantity or Volume
Neurobasal media n/a 100 mL
N2 supplement 1% 1 mL
B27 supplement 2% 2 mL
GlutaMAX 1% 1 mL
FBS 1% 1 mL
GDNF (10 µg/mL stock) 25 ng/mL 250 µL
BDNF (10 µg/mL stock) 25 ng/mL 250 µL
Ascorbic acid (100 mM stock) 200 µM 200 µL
CNTF (100 µg/mL stock) 25 ng/mL 10 µL
dbcAMP (100 mM stock) 200 µM 200 µL
Retinoic acid (1 mM stock) 0.125 µM 12.5 µL
Note: Retinoic acid should be added to the medium freshly every feeding.
Salivary gland cell medium (10 mL)
Reagent Final concentration Quantity or Volume
DMEM/F12 1× 9.6 mL
FBS 1% 100 µL
B27 1× 200 µL
Antibiotic-antimycotic 1× 100 µL
Tongue tissue separation solution (2 mL)
Reagent Final concentration Quantity or Volume
Collagenase A (2 mg/mL stock) 1 mg/mL 1.0 mL
Dispase II (5 mg/mL stock) 2.5 mg/mL 1.0 mL
PBS 1×
Salivary gland cell dissociation solution (3 mL)
Reagent Final concentration Quantity or Volume
Trypsin-EDTA 0.25% 3 mL
Dissociation termination solution (2 mL)
Reagent Final concentration Quantity or Volume
DMEM/F12 1× 1800 µL
FBS 10% 200 µL
Laboratory supplies
TC-treated 6-well tissue culture plate (Corning, catalog number: 3516)
TC-treated 24-well tissue culture plate (Corning, catalog number: 3526)
TC-treated 10 cm tissue culture plate (Corning, catalog number: 430167)
Ultra-low attachment plate (Corning, catalog numbers: 07 200 601 and 07 200 602)
15 mL conical tissue culture tubes (VWR/Corning, catalog number: 89039-664)
50 mL conical tissue culture tubes (VWR/Corning, catalog number: 89039-656)
Cryovial (Thermo Fisher/Life Technologies, catalog number: 375353)
Trypan blue (Corning, catalog number: MT-25-900-CI)
P2-10, 20, 200, 1000 pipettes and tips (Eppendorf)
Parafilm (ULINE)
5 mL low retention vials (Eppendorf, catalog number: 30122348)
35 mm dish (Genesee Scientific, catalog number: 32-103G)
100 mm culture dishes (Genesee Scientific, catalog number: 32-107G)
3 mL syringes (BD, catalog number: 8194938)
30-G needle (BD, catalog number: 9193532)
70 µm cell strainer (Fisher Scientific, catalog number: 352350)
35 µm cell strainer (Electron Microscopy Science, catalog number: 64750-25)
100% CO2 gas (Airgas, CD USP50)
Plastic CO2 euthanasia chamber (Length: 10 cm, wide: 7 cm, and height: 5 cm)
Equipment
Water bath (VWR)
CellDrop automated cell counter (DeNovix)
Racks
Liquid nitrogen tank (Custom Biogenic Systems)
Freezer (-20 °C)
Refrigerator (2–8 °C)
Water bath (37 °C)
Centrifuge (Eppendorf)
37 °C incubator with 5% CO2
Class II biological safety hood
Lionheart FX microscope (BioTek)
Dissecting microscope (Olympus, model: SZX16 upright fluorescent and brightfield)
Biosafety cabinet (NUAIRE, model: GellGard, Class II, Type A2)
CO2 incubator (NUAIRE)
Centrifuge (Thermo Scientific, model: LEGEND XTR)
Light microscope (EVOS XL Core)
Tank with CO2 (Airgas)
Instruments
Surgical and fine scissors (Moria, catalog number: 9601; Fine Science Tool, catalog number: 91604-09)
Surgical and fine forceps (Inox electronic, catalog number: 91150-20; Fine Science Tool, catalog number: 112900)
Spatula (Fine Science Tool, catalog number: 10360-13)
Software and datasets
Gen5 (BioTek)
Prism v10.0.3 (GraphPad, 9/25/2023)
Fiji (Image J)
Adobe Photoshop
CellSens Dimension
Procedure
Caution: The procedures for sections A and B will take 1.5–2.0 h without an opportunity to pause. The experiments need to be done as efficiently as possible for harvesting healthy cells. Please make sure you have all the instruments, solutions, and equipment ready before euthanizing the animals.
Salivary gland dissection
Euthanize the 8-week-old Sox10-Cre::RFP mice by placing them in a CO2 chamber. Ensure mouse death by cervical dislocation.
Place the mouse in the surgical area. Wet the mouse head using 70% ethanol to disinfect the surface and prevent fur from getting into the oral cavity.
Decapitate the mouse. Hold the head in one hand and open the mouth cavity by pulling the skin over the skull and below the mandible (Figure 1A).
Cut the corners of the mouth along the cheek using dissecting scissors to further open the oral cavity (Figure 1B).
Figure 1. Opening the mouse oral cavity. (A) Image of ventral view of a mouse head. (B) Image of a mouse head with the mouth cavity widely open.
Dissect the tongue with the mandible and remove the skin covering the mandible (Figure 2). Transfer the tongue on the mandible to 30 mL of fresh sterile DPBS (Ca+2 and Mg+2 free) in a 50 mL tube. Wash five times using DPBS.
Figure 2. Dissected tongue with the mandible. A representative image of the dorsal view of an adult mouse tongue sitting on the mandible. Black dotted line encircles the single circumvallate taste papilla in the midline of posterior tongue. Scale bar: 500 μm.
Using surgical forceps to hold the tongue under a dissecting microscope, inject ~0.5 mL of the tongue tissue separation solution (a mixture of Collagenase A and Dispase II enzymes) into the subepithelial space of posterior tongue as described [17] (Figure 3A and B) and incubate for 30 min at 37 °C (Figure 3C).
Note: See Troubleshooting #3 and #5 for precautions.
Figure 3. Intralingual injection of tongue tissue separation solution. (A) An image to illustrate the entry and location of the needle for injecting the solution into the posterior part of the tongue. (B) An image of dorsal view of the posterior tongue after 0.5 mL solution injection. (C) An image of dorsal view of the posterior tongue after 15 min of incubation at 37 °C. Black dotted lines encircle the single circumvallate taste papilla. Scale bars: 500 μm.
After the incubation, dissect the posterior tongue region with RFP+ von Ebner’s glands. Use small scissors to cut the epithelium of the circumvallate papilla region (Figure 4A), then use fine forceps to peel off the epithelium of the circumvallate papilla region (Figure 4B).
Figure 4. Peeling the circumvallate epithelium. (A) Brightfield image of the dorsal view of the posterior tongue region after removing part of the epithelium. Black dotted line outlines the cutting edge of epithelium in the circumvallate papilla area with the epithelium removed. (B) Brightfield image of the ventral view of the peeled epithelium. White dotted line encircles the circumvallate papilla epithelium. Scale bars: 500 μm in A; 100 μm in B.
Under a fluorescent stereomicroscope, we were able to see the RFP+ von Ebner’s glands in the circumvallate region (Figure 5A). Dissect and collect ~0.1 mL of von Ebner’s salivary gland tissue in the surrounding of the circumvallate papilla under the fluorescent microscope (Figure 5B).
Caution: If you collect more gland tissues for more cells, you will need to use more solution of 0.25% trypsin-EDTA accordingly.
Figure 5. Dissection of von Ebner’s gland tissue from Sox10-Cre::RFP mice. (A) RFP fluorescent image of the dorsal view of the posterior tongue region under the fluorescent stereomicroscope. Bright RFP+ glands were readily visible in the regions surrounding and under the circumvallate papilla. White dotted line in A encircles the circumvallate papilla. White dashed squares encircle the circumvallate papilla-adjacent regions from which glands were collected. (B) RFP fluorescent image of two pieces of dissected tissue enriched RFP+ Ebner’s glands (white arrowheads). Scale bars: 500 μm.
Wash the salivary glands in 15 mL of fresh sterile DPBS in a 100 mm sterile culture dish.
Salivary gland cell dissociation
Transfer salivary glands into a 35 mm dish.
Use fine-tip scissors to cut the salivary gland tissues into small pieces (Figure 6).
Figure 6. Tissue pieces of Ebner’s glands after mincing. RFP+ Ebner’s gland tissue (Figure 5B) was cut into smaller pieces using fine scissors under a fluorescent stereomicroscope. Arrows point to a few pieces of RFP+ Ebner’s gland tissue after mincing. Scale bar: 200 μm.
Incubate the tissue pieces in 3 mL of salivary gland cell dissociation solution (0.25% trypsin-EDTA) for 30 min at 37 °C. Gently agitate the tissues every 5 min using 1 mL pipette tips. Do not cut the tip, as the sharp edge of the cutting end may injure the cells.
Note: See Troubleshooting 4 for precautions.
Add 1 mL of dissociation termination solution to stop the reaction and transfer all of the solution with cells into a 5 mL low-binding vial.
Centrifuge the cell suspension at 450× g for 10 min at 4 °C and remove the supernatant as much as possible. Be careful not to disturb the cell pellet.
To completely remove the remaining trypsin-EDTA, repeat step B4, adding 1 mL of dissociation termination solution to suspend the cells. Then, repeat step B5 for the centrifugation.
Remove the supernatant and leave one-fourth of the medium (250–300 mL) without disturbing the pellet and filter the cells using a 70 µm cell strainer, followed by a 35 µm cell strainer.
Collect the filtered cell suspension and proceed straight to Section C (Figure 7).
Figure 7. Cell suspension. Photomicrographs of the dissociated cells and some small cell clusters from RFP+ Ebner’s glands. Scale bar: 100 μm.
Salivary cell passaging
Centrifuge the cell suspension at 200× g for 5 min.
Aspirate the medium.
Wash cells twice with PBS (1 mL/24-well, 3 mL/6-well, and 4 mL/10 cm plates).
Add prewarmed (5 min in a water bath) trypsin-EDTA (1 mL/24-well, and 3 mL/10 cm plates).
Incubate for 5 min in a 37 °C incubator.
Pipette gently with a P1000 pipette 3–5 times to fully dissociate the cells.
Transfer the trypsin-EDTA and cell solution to a conical tissue culture tube that contains at least 5 times higher volume of salivary gland cell medium.
Centrifuge at 200× g for 5 min.
Resuspend the cells with 1 mL of salivary gland cell medium.
Count the cells.
Replate the cells on desired tissue culture plates at 15 × 103/cm2 density with salivary gland cell medium (0.5 mL/24-well, 2 mL/6-well, and 7 mL/10 cm plates).
Change the medium the next day, and every 2–3 days after (1 mL/24-well, 3 mL/6-well, and 10 mL/10 cm plates).
Cells should reach >80% confluency in one week (Figure 8) and will be ready for replating/freezing.
The ideal passage number of the cells for the experiment is P3–8. Please see also troubleshooting 2.
Figure 8. Cell morphology of salivary gland cells after replating. Brightfield images showing the isolated salivary gland cells after (A and A’) 24 h, and (B and B’) 4–6 days. Cell density in B is ideal for making frozen stocks or replating the SCPs to start the coculture. Scale bars = 200 µm.
(Optional pause point) Salivary cell cryo-banking
After passage 3, resuspend 80% confluent and dissociated cells from a 10 cm dish into 1 mL of salivary gland cell medium that contains 10% DMSO.
Transfer the medium with the cells into a cryogenic tube.
Freeze the vial at -80 °C fridge in a wrapped Styrofoam box.
Next day, move the vial to a LN tank for long-term storage. The cells can be safely preserved at this stage until the hPSC-derived cells are ready.
NCC induction
Prepare Geltrex-coated plates.
Thaw Geltrex at 4 °C overnight.
Dilute Geltrex with ice-cold DMEM/F12 at 1:100 dilution.
Coat the plates with Geltrex (1 mL/24-well, 2 mL/6-well plates) and seal with parafilm.
Incubate the plates at 4 °C at least overnight before use.
Plating hPSCs on day 0
Resuspend dissociated hPSCs (15 min by EDTA in order to gain single cells) in 1 mL d0, 1 media. hPSC is maintained in E8 medium. For details of hPSC replating, please also check our previous protocols [10,18].
Aspirate the Geltrex solution from the plates coated overnight.
Plate hPSCs at 125k cells/cm2 density (0.5 mL/24-well, 2 mL/6-well plates).
Incubate cells at 37 °C.
Next day (day 1), fully change the medium with NCC induction, d0, 1 media (1 mL/24-well, 3 mL/6-well plates).
On day 2, fully change the medium with NCC induction, D2–10 medium (1 mL/24-well, 3 mL/6-well plates).
From now on, feed the cells by full medium change every two days (days 2, 4, 6, 8) until day 10 (1 mL/24-well, 3 mL/6-well plates).
On day 10, differentiated NCCs should actively aggregate and form the condensed “dark ridge” structure (Figure 9) [10].
Caution: If you fail to see such structures, it is not recommended to proceed to the next step.
Figure 9. Cell morphology during Schwann cell precursor (SCP) differentiation. Brightfield images showing the differentiated D10 neural crest cell (NCC) “dark ridges” (indicated by the yellow arrow). Scale bar = 200 µm.
(Optional pause point) Dissociate NCCs for cryobanking
On day 10, aspirate the culture medium.
Wash cells twice with PBS (1 mL/24-well, 3 mL/6-well, and 4 mL/10 cm plates).
Add Accutase (1 mL/24-well, and 2 mL/10 cm plates).
Incubate for 20–30 min in a 37 °C incubator.
Pipette gently with a P1000 pipette 3–5 times and fully dissociate the cells.
Test if cells are ready for dissociation by gently pipetting 1–2 times. If more times are needed, incubate 5 more minutes each time and test for up to 30 min.
Transfer the Accutase and cell solution to a conical tissue culture tube that contains at least 5 times higher volume of PBS.
Centrifuge at 200× g for 5 min.
Resuspend the cells with ice-cold stem cell banker.
Count the cells.
Transfer the cells (8 × 106 cells/1 mL/vial) in ice-cold stem cell banker to cryogenic tubes.
Freeze the vial at -80 °C freezer in a wrapped Styrofoam box.
Next day, move the vial to a LN tank for long-term storage. The differentiation can be safely paused at this stage.
Dissociate NCCs for SCP differentiation
On day 10, aspirate the culture medium.
Wash cells twice with PBS (1 mL/24-well, 3 mL/6-well, and 4 mL/10 cm plates).
Add Accutase (1 mL/24-well, and 2 mL/10 cm plates).
Incubate for 20–30 min in a 37 °C incubator.
Pipette gently with a P1000 pipette 3–5 times and fully dissociate the cells.
Test if cells are ready for dissociation by gently pipetting 1–2 times. If more times are needed, incubate 5 more minutes each time and test for up to 30 min.
Transfer the Accutase and cell solution to a conical tissue culture tube that contains at least 5 times higher volume of PBS.
Centrifuge at 200× g for 5 min.
Resuspend the cells with day 10–16 medium.
Count the cells.
Replate the cells on 24-well ultra-low attachment tissue culture plates at 1 × 106/well density with SCP differentiation, D10–16 medium (0.5 mL/well).
Add day 10–16 medium on the next day (0.5 mL/well), which makes the total volume 1 mL.
Feed cells on days 13 and 15 by full medium change.
In order to keep all the cells, tilt the plate and aspirate the medium slowly. Always keep the tip on the surface of the culture medium away from the cells (Figure 10A).
On day 16, SCPs should be differentiated in suspending spheroids (Figure 10B).
Caution: If you fail to see such structures, it is not recommended to proceed to the next step.
Figure 10. Cell morphology during Schwann cell precursor (SCP) differentiation. (A) Cartoon illustration of the method to change medium in suspension cultures. (B) Brightfield images showing the differentiated D16 SCP spheroids. Scale bar = 200 µm.
SCP replating for parasymN differentiation/coculture
Please make sure you have 80% confluent salivary cells at passages 3–8 before proceeding.
On day 16, tilt the plate and collect all the SCP spheroids in the culture medium into a conical tissue culture tube and fill up the tube with PBS (at least the same volume of cell suspension).
Centrifuge at 200× g for 3 min.
Aspirate the supernatant and add Accutase to the spheroids (1 mL for less than 12 wells, 2 mL for less than 24 wells).
Transfer the Accutase that contains the spheroids back to a low attachment well.
Incubate for 20–30 min in 37 °C incubator.
Gently pipette the spheroids with a P1000 pipette, transfer the cells to a conical tissue culture tube, and fill up the tube with PBS.
Centrifuge at 200× g for 5 min.
Resuspend the cells with 1 mL day 16 parasymN medium.
Count the cells.
Replate SCP cells at a density of 1 × 105/cm2 on top of 80% confluent salivary cells (passage 3–8) in a 24-well plate containing 0.5 mL parasymN differentiation medium.
Fully change the medium on the next day and every 2–3 days after (1 mL/24-well plates).
After 7–10 days, aggregated cell bodies of parasymNs should be observed on the salivary cell layer with well-developed neurites (Figure 11).
Figure 11. Cell morphology of the coculture. Brightfield images showing the cell bodies (indicated by the yellow arrows) and the axons (indicated by the red arrows) of hPSC-parasymNs on the salivary cells (indicated by the white arrow) 10 days after the coculture. Scale bar = 200 µm.
Data analysis
After 7–10 days, well-developed parasymNs should be observed on the salivary cell layer. Cell bodies of parasymNs should be clearly distinguishable from the salivary tissue, given that parasymNs will aggregate with bright and round cell bodies (Figure 11, yellow arrows). The development of neurons and their connectivity with salivary cells can be clearly observed by immunostaining with specific markers as desired (for instance, PRPH in Figure 12A) [19]. Secretory granules in salivary cells are autofluorescent and should be observable at high resolution (Figure 12B). Both total salivary cell number and granule+ salivary cells are increased with parasymNs, suggesting improved development and maturation (Figure 12C). We can also monitor Ca2+ dynamics in the salivary cells (Figure 12D). The timeline of the entire process is summarized in Figure 13.
Figure 12. Sample result of functional activity of the coculture. A. Immunofluorescent staining for peripheral neural marker PRPH shows parasymNs growing on the salivary tissue with extensive neurites and aggregating cell bodies. This feature can be mostly seen in peripheral neuron cultures since it might resemble the ganglion formation in the body. B. RFP signals representing SOX10+ salivary cells as well as the autofluorescent granules. C. Representative quantification result showing the improved proliferation and maturation of the salivary tissues when cocultured with parasymNs. D. Representative Ca2+ image (using fluo-4) for the salivary cells shows the functional connectivity between parasymNs and salivary cells upon parasymN activation by nicotine in the coculture. Scale bars = 200 µm.
The replating density of D16 SCP is critical for parasymN differentiation especially in neural culture alone. However, given that current understanding suggests that there might be mutual effects on the development and maturation of cocultured cells [8], the cell density in cocultures might be adjusted; however, we have not tested this. Nevertheless, if the density is too high, the chance that the cells detach after coculture will be higher. It is also worth mentioning that, while hPSC-derived parasymNs allow an unlimited neuron source for research, the nature of small amounts and limited passage numbers of primary salivary tissues still restrain the scale-up option of this coculture model system. Establishing a coculture model with both hPSC-derived parasymNs and hPSC-derived salivary tissues would be an important future task.
Figure 13. Timeline of the coculture
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Wu et al. [12]. Parasympathetic neurons derived from human pluripotent stem cells model human diseases and development. Cell Stem Cell (Figure 6, panel H, and supplementary figure 9).
Yu et al. [16]. The duct of von Ebner’s glands is a source of Sox10+ taste bud progenitors and susceptible to pathogen infections. Frontiers in Cell and Developmental Biology (Figure 1, panel A and B).
General notes and troubleshooting
General notes
Efficient and successful early NCC induction for the first 10 days of differentiation is critical. For more detailed methods and tips, please also see our previous protocols [10,18].
It is always recommended to replate a neuron-only group and a salivary gland-only group in parallel to the cocultures to validate the neuron differentiation efficiency and functionality as well as the salivary gland cell integrity of each batch. When differentiating parasymNs alone, coat the plate with PO (15 μg/mL in DPBS)/LM (2 μg/mL in DPBS)/FN (2 μg/mL in DPBS).
Troubleshooting
Problem 1: D16 SCP spheroid is poorly formed.
Possible cause: D10 NCC is not properly differentiated (very few or no NC ridges can be observed).
Solution: It is critical to make sure that each stage of the differentiation, including D0 hPSC and D10 NCC, has passed the quality control steps as previously described [10]. If NCC differentiation is not ideal, a BMP4 titration test is highly recommended, as detailed in [18]. In general, concentrations between 0.2 and 1 ng/mL are ideal. In our case, for example, we had to drop the concentration from 1 to 0.4 ng/mL due to the batch-to-batch variability of BMP4.
Problem 2: ParasymNs in the cocultures do not regulate the activity of the salivary tissues.
Possible cause: The parasymN or salivary cell batch is bad.
Solution: Only use salivary cells between passages 3 and 8. Always replate a neuron-only group in parallel to the coculture to validate the neuron differentiation efficiency and functionality of each batch.
Problem 3: Difficulty in injecting tongue tissue separation solution into subepithelial space.
Possible cause: Incorrect injection technique or improper handling of the tongue.
Solution: Practice the injection technique to ensure accurate delivery of solution. Use a dissecting microscope for better visualization and precision. Be careful not to poke and injure the tongue epithelium; otherwise, the solution will leak, leading to incomplete enzyme digestion.
Problem 4: Incomplete or over-enzyme digestion for tongue tissue separation.
Possible cause: Insufficient enzyme incubation time, incorrect enzyme concentration, or improper injection of enzyme. Solution: Verify the concentration and activity of Collagenase A and Dispase II. Ensure proper mixing and incubation at the correct temperature for the specified duration.
Problem 5: Primary cell clumping or low viability.
Possible cause: Over-digestion of tissues for the separation or excessive mechanical stress during pipetting.
Solution: Control the incubation time ≤ 30 min at 37 °C after the injection of tongue tissue separation solution. Agitate gently during the incubation with salivary gland cell dissociation solution. Consider optimizing the trypsin-EDTA concentration and incubation time.
Acknowledgments
This work was supported by 1R01NS114567-01A1 to NZ and R21DC018910 and R21DC018089 to HXL. This protocol was adapted from Wu et al. [12] and Yu et al. [15].
Competing interests
The authors declare no competing interests.
Ethical considerations
This study employed human embryonic stem cell lines (WA09), the use of which by the Zeltner lab was approved by WiCell. All iPSCs employed in this work were reprogrammed from human samples obtained through the public repository Coriell Research Institute. The use of animals was approved by the Institutional Animal Care and Use Committee at The University of Georgia (AUP# A2022 06-030-A6) and complied with the National Institutes of Health Guidelines for the care and use of animals for research.
References
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Scott-Solomon, E., Boehm, E. and Kuruvilla, R. (2021). The sympathetic nervous system in development and disease. Nat Rev Neurosci. 22(11): 685–702.
Muo, E. C., Cardona, J. J., Chaiyamoon, A., Iwanaga, J. and Tubbs, R. S. (2023). The Parasympathetic Root of the Submandibular Ganglion: A Review. Cureus. 15(1): e33775.
Takano, T., Wahl, A. M., Huang, K. T., Narita, T., Rugis, J., Sneyd, J. and Yule, D. I. (2021). Highly localized intracellular Ca2+ signals promote optimal salivary gland fluid secretion. eLife. 10: e66170.
Frith, T. J. R. and Tsakiridis, A. (2019). Efficient Generation of Trunk Neural Crest and Sympathetic Neurons from Human Pluripotent Stem Cells Via a Neuromesodermal Axial Progenitor Intermediate. Curr Protoc Stem Cell Biol. 49(1): e81.
Kirino, K., Nakahata, T., Taguchi, T. and Saito, M. K. (2018). Efficient derivation of sympathetic neurons from human pluripotent stem cells with a defined condition. Sci Rep. 8(1): 12865.
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Wu, H. F., Saito-Diaz, K., Huang, C. W., McAlpine, J. L., Seo, D. E., Magruder, D. S., Ishan, M., Bergeron, H. C., Delaney, W. H., Santori, F. R., et al. (2024). Parasympathetic neurons derived from human pluripotent stem cells model human diseases and development. Cell Stem Cell. 31(5):734–753.e8.
Espinosa-Medina, I., Saha, O., Boismoreau, F., Chettouh, Z., Rossi, F., Richardson, W. D. and Brunet, J. F. (2016). The sacral autonomic outflow is sympathetic. Science. 354(6314): 893–897.
Proctor, G. B. and Carpenter, G. H. (2014). Salivary secretion: mechanism and neural regulation. Monogr Oral Sci. 24: 14–29.
Yu, W., Ishan, M., Yao, Y., Stice, S. L. and Liu, H. X. (2020). SOX10-Cre-Labeled Cells Under the Tongue Epithelium Serve as Progenitors for Taste Bud Cells That Are Mainly Type III and Keratin 8-Low. Stem Cells Dev. 29(10): 638–647.
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Article Information
Publication history
Received: Jul 19, 2024
Accepted: Sep 9, 2024
Available online: Oct 13, 2024
Published: Nov 5, 2024
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© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Semi-Automated Assessment of Long-Term Olfactory Habituation in Drosophila melanogaster Using the Olfactory Arena
CR Camilla Roselli
MR Mani Ramaswami
MA Marcia M. Aranha
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5102 Views: 372
Reviewed by: Geoffrey C. Y. LauRaghavendra Yelahanka NagarajaXiaochen Sun
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Abstract
Long-lasting memories are a core aspect of an animal’s life. Such memories are characterized by unique molecular mechanisms and often unique circuitry, neither of which are completely understood in vivo. The deep knowledge of the identity and connectivity of neurons of the fruit fly Drosophila melanogaster, as well as the sophisticated genetic tools that allow in vivo perturbations and physiology monitoring, make it a remarkably useful organism in which to investigate the molecular mechanisms of long-term memories. In this protocol, we focus on habituation, a non-associative form of learning, and describe a reliable, semi-automated technique to induce and assess long-term olfactory habituation (LTH) in Drosophila using the olfactory arena, thus providing a method aligned with recent technological progress in behavioral measurement. Prior work has shown that LTH is induced by a 4-day exposure to an odorant and is characterized by a long-lasting (> 24 h) reduction in behavioral response to the exposed odorant, measured using a manual and skill-intensive Y-maze assay. Here, we present a semi-automated protocol for obtaining quantifiable measures of LTH, at the level of detail required for other investigators in the field. Unlike previously described methods, the protocol presented here provides quantitative and detailed behavioral measurements obtained by video recording that can be shared with the scientific community and allows sophisticated forms of offline analysis. We suggest that this procedure has the potential to advance our understanding of molecular and circuit mechanisms of olfactory habituation, its control via neuromodulation, and its interactions with other forms of memory.
Key features
• A protocol to induce olfactory aversive long-term habituation in Drosophila melanogaster.
• Video recording and analysis of Drosophila in an olfactory arena.
Keywords: Long-term habituation Olfactory habituation Olfactory arena Drosophila melanogaster
Background
Habituation is a fundamental form of learning that reduces an animal’s behavioral response after continuous or multiple exposures to an innocuous stimulus [1,2]. This reduction in behavioral response is adaptive, allowing animals to dedicate cognitive resources to potentially salient elements of their environment [3]. A defining feature of habituation is that it can be instantly reversed, as revealed by an animal’s ability to reengage either volitionally or in a new context, with a familiar, previously ignored stimulus [4]. Habituation has also been described in Drosophila melanogaster using a wide range of sensory modalities—visual [5,6], gustatory [7–9], olfactory [10,11], or of other types of stimuli, such as shock [12] or courtship [13]. However, long-term forms of habituation (LTH) have only been studied for olfactory behavior [14–17]. LTH requires molecular mechanisms shared with other forms of long-term memory, notably de novo transcription and protein synthesis [12,15,18,19]. Hence, studies on LTH have the potential to reveal not only circuit mechanisms that flexibly gate stimulus responses but also in vivo molecular mechanisms, such as translational control of mRNAs, involved in the consolidation of long-term memories.
The protocol we describe for LTH is distinctive in its use of the olfactory arena [20–22], which allows flies to exhibit more naturalistic, non-binary behavioral responses. Being based on video recording and machine vision, this protocol also allows offline analyses of multiple aspects of flies’ behavior, with the potential to provide a broader and deeper understanding of circuit and molecular mechanisms for long-term habituation.
Materials and reagents
Biological materials
Canton S
Oregon R
Reagents
Malt extract (Muntons, amber malt extract, 25 kg, catalog number: 84260)
Polenta (Heera-Corn Meal Fine, 1.5 kg)
Golden Syrup (GEM, 7.26 kg)
Dry yeast (Oxioid, catalog number: LP0021)
Agar (Apollo Scientific-Micro Agar, catalog number: BIM1002)
Propionic acid (Thermo Fisher Scientific, catalog number: 149300010)
Methyl 4-hydroxybenzoate (Merck, catalog number: H5501-1KG)
Paraffin oil (Sigma-Aldrich, catalog number: 18512-2.5L)
Ethyl butyrate (Sigma-Aldrich, catalog number: E15701-500mL)
Methyl hexanoate (Sigma-Aldrich, catalog number: 259942-100mL)
3-octanol (Fluka Chemica, catalog number: 74870-50mL)
CO2 (BOC Gases Ireland Limited)
Agar for starvation vials (Sigma-Aldrich, catalog number: A1296-500)
Solutions
Fly food (see Recipes)
Starvation vials (see Recipes)
Recipes
Fly food
Reagent Quantity
Malt extract 82 g
Polenta 77 g
Golden syrup 16 g
Dry yeast 18 g
Agar 11 g
Propionic acid 4.5 mL
10% methyl 4-hydroxybenzoate (in ethanol) 10 mL
Water Up to 1 L
Starvation vials
Reagent Quantity
Agar 0.75 g
Water Up to 100 mL
Laboratory supplies
Fly vials (Regina Industries, catalog number: P1014T)
Fly bottles (Genesee Scientific, catalog number: 32-129F)
PCR tubes (Azenta, catalog number: 4TI-0790)
Microlance needles 21 G × 38 mm (BD, catalog number: 304432)
Copper wire (RS, catalog number: 779-0707)
PRO White PTFE Tape, 5 m × 12 mm × 0.2 mm (RS, catalog number: 231-964)
Equipment
Clear plastic box 1.6 L, dimensions 195 × 135 × 110 mm (Really Useful Boxes)
Incubators (Memmert, model: IPP 400)
Vortex-Genie 2 (Scientific Industries, Inc., catalog number: 103-SI-0256)
Waterproof silver and black blackout UV-protect water-resistant fabric (Amazon)
Climate box with red light bar (Con-Elektronik/Konrad Oechsner)
Odor delivery system
One olfactory arena for fruit flies [20]
Two mass flow controllers 0-500 SCCM (Omega Engineering Limited, catalog number: FMA5512A)
TYGON SE200 Tubing 1/16 (I.D.) × 1/8 (O.D.) (Fisher Scientific, catalog number: 11727293)
TYGON SE200 Tubing 1/8 (I.D.) × 1/4 (O.D.) (Fisher Scientific, catalog number: 15590275)
PTFE Tubing [50 ft (L) × 1/8 in. (O.D.) × 0.063 in (I.D.)] (Supelco, catalog number: 58699)
PTFE Tubing [25 ft (L) × 1/4 in. (O.D.) × 0.228 in (I.D.)] (Supelco, catalog number: 20533)
Flowmeter system, 65 mm direct reading flow tube for air; 500 mL/min (Fisher Scientific, catalog number: 15949117)
Vacuum pump (KNF LABOPORT, model: N 86 KT.18)
Connectors:
Diameter straight tube-to-tube connector, push in 3.2 mm to push in 6 mm (Radionics, catalog number: KQ2H23-06A)
Flangeless fittings 1/8" PK/5 (Supelco, catalog number: 58686)
Tee tube-to-tube adapter (3.2 mm OD tubing) (Radionics, catalog number: KQ2T23-00)
Fitting plug 1/4-28 (NResearch, catalog number: FITM128)
Luer to threaded fitting, straight adapter (1/16 female luer to 1/4-28 UNF thread) (Fisher Scientific, catalog number: 15196303)
Male luer with lock ring 1/16" ID (Fisher Scientific, catalog number: 14889980)
Y union barbed fitting 1/16" ID (Cole Parmer, catalog number: WZ-50111-30)
Barbed fitting 10:32 × 1/16" (NResearch, catalog number: FITM328)
Male pipe adapter barbed fittings (Fisher Scientific, catalog number: 15277536)
Two 5-port low-pressure manifold body, 0.062" bore, 1/8" OD tubing, 1/4-28 flat bottom; 1/EA (Cole Parmer, catalog number: P-155)
Six screw cap GL 14 for hose connection for Duran bottles (Duran Group, catalog number: 1129814)
Twelve insert for screw cap, GL 14, 3.2 mm (1/8 Zoll) ID (Duran Group, catalog number: 1129817)
Six 100 mL SCHOTT Duran bottles with GL45 neck (Sigma-Aldrich, catalog number: Z305170-10EA)
Two 3-way isolation valve 12VDC (NResearch, catalog number: 225T031)
Four 2-way normally closed isolation valve 12VDC (NResearch, catalog number: 225T011)
One CoolDriveSolo multi-board with IDC connectors (NResearch, CDSX8UK-IDC)
Six CoolDriveSolo Universal valve drivers (trigger voltages 4.5–24 VDC) (NResearch, catalog number: CDS-V01)
MAPLIN 36W switched mode DC variable voltage compact bench power supply 6 output (Amazon)
Insulated breadboard jumper wire (Radionics, catalog number: 791-6454)
Arduino MEGA 2560 (Radionics, catalog number: 715-4084)
USB cable, type A to B (for Arduino) (Radionics, catalog number: 282-4023)
One Point Grey Flea3 digital camera (ClearView Imaging, catalog number: FL3-U3-32S2M-CS)
Point Grey USB 3.0 cable, Type-A to Micro-B (ClearView Imaging, catalog number: ACC-01-2301)
Computar lens M0814-MP2 F1.4 f8 mm 2/3" (MultiPix imaging, catalog number: 2000034697)
Desktop computer (Windows PC, ≥8 GB RAM, nice GPU)
Software and datasets
Bonsai Visual reactive programming [Bonsai (2.8.0), 30/07/2023] [23]
MATLAB_R2023a [R2023a (9.14), 22/02/2023]
Prism 10 (GraphPad, 11/07/2023)
Setting up the olfactory arena
All details for manufacturing the various parts for the olfactory arena and the LED panel were obtained from the Janelia Farm Tech-Transfer office as described by Aso and Rubin [20]. Additionally, we developed an odor delivery system where saturated odor vapor can be independently delivered to two diagonally opposing quadrants (Figure 1). Two mass flow controllers ensured a constant flow rate of 100 mL/min in each quadrant. The airflow then converged to a central hole of the arena and was extracted at 400 mL/min with a vacuum pump. Odors were delivered to the arena by rapidly diverging the airflow stream to selected odor vials using solenoid valves. Experimenters could choose among three different odors to be delivered on two diagonally opposite quadrants of the arena.
Figure 1. Schematic of the odor delivery system. Saturated odor stream, produced by bubbling the air stream through a selected odor vial (e.g., Odor 1) can be independently delivered to two diagonally opposing quadrants. The air stream can be rapidly diverged to odor vials 2 and 3 by simultaneously opening two solenoid valves (V1 and V2 or V1 and V3, respectively). A constant flow rate of 200 mL/min on each odor line is ensured by two mass flow controllers. This flow bifurcates prior to entering the arena such that each quadrant receives a constant flow rate of 100 mL/min. The airflow then converges to a central hole of the arena and is extracted at 400 mL/min with a vacuum pump. Valves are energized using a 12 V power supply via CoolDriveSolo valve drivers. Open and close commands are sent from the computer to the valves through an Arduino.
Connect each mass flow controller to solenoid valves (V1) using odor-inert PTFE tubing and connectors, as shown in Figure 2A and B. Push the 1/4" OD (outer diameter) tubing through the push connector of the mass flow controller. Then, using a different diameter straight connector, attach one end of the 1/8" OD tubing to this connector and the other end to the common port of the 3-way valves using the flangeless fittings (Figure 2B).
Connect 3-way solenoid valves and odor vials (odor vial 1). Attach one end of the 1/8" OD PTFE tubing to the normally open port of the 3-way valve using the flangeless fittings and pass the other end of the tubing through the odor bottle screw cap with the insert, making sure the tubing is close to the bottom of the odor vial (Figure 2C). Push a separate piece of tubing through the second screw cap and insert, ensuring the end protrudes slightly from the odor vial lid. Connect the other end of this last tube to the manifold using the flangeless fittings. We obtained similar success using either SCHOTT Duran bottles with lids or scintillation vials with rubber caps as odor vials.
To allow diversion of the air stream to other odor vials, connect the normally closed port of the 3-way valve with two 2-way valves (V2 and V3) by using a T connector to bifurcate the odor stream onto the two 2-way valves (Figure 2D). Connect the 2-way solenoid valves and odor vials (odor vial 2 and 3) as explained above, then connect the output tubing from odor vials with the manifold (Figure 2E). Each manifold can be connected to up to four different odor vials. Use a fitting plug to block the unused manifold port.
Attach the odor inert flexible tubing TYGON SE200 1/16" ID (inside diameter) to the output port of the manifold. Use the luer to thread and the male luer fittings to attach the tubing (Figure 2E). Then, connect the other end of the tubing to two opposing quadrants, first by bifurcating the tubing into two using a Y connector (Figure 2F) and then by attaching each of the two tube ends to the barbed fittings at two diagonally opposite quadrants of the arena (Figure 2K).
Connect the arena’s vacuum fitting and the flow meter using a TYGON SE200 flexible tubing (1/8" ID) and a male barbed fitting (Figure 2G–H), then connect the flow meter to the pump. Finally, direct the pump output to an extractor to prevent odor accumulation in the room.
Cut a piece of red and black hookup wire. Connect one end of each wire to the 12 V power supply and the other end to the CoolDriveSolo multi-board (Figure 2I). Make sure there are no exposed wires outside the mount housing. Insert the CoolDrive valve controller driver on the multi-board (Figure 2J). You will need as many drivers as valves, which means 6 in our case; each board allows independent operation of up to eight solenoid valves. Connect the two wires of the valve to the CoolDriveSolo multi-board. Connect the CoolDriveSolo multi-board to the Arduino via insulated breadboard jumper wires, then plug the Arduino with the computer.
Place a high-resolution camera with a lens above the arena to record behavior and plug it into the computer with a USB 3.0 cable (Figure 2K). A custom-made infra-red LED panel and LED controller, made as described by the Janelia Farm Tech-Transfer office [20], is used as a light source. A desktop Windows PC with at least 8 GB RAM is needed for managing the experimental session and controlling the camera, valves, and LED panel.
The arena and associated odor delivery system should be placed in a climate box covered with a blackout cloth. We highly recommend checking all the connections for potential air leakage before operating the arena for the first time. From our experience, the most common source of air leakage is the Duran bottles used to contain the odors. Ensure that the bottles are evenly closed and use PTFE tread seal tape around the neck of the Duran bottles to avoid air leakage. While checking that connections are tight, make sure that you do not overtighten the connections with the solenoid valves—they are made of soft Teflon and can easily be damaged if overtightened.
Figure 2. Odor delivery assembly steps. (A) Add a mass flow controller to the odor delivery system by pushing one end of the PTFE tubing through the push connector. (B) Connect the mass flow controller with the common port of the 3-way valve, and then connect the normally open port with odor vial 1. Valve wires will be connected to the MultiBoard as described further ahead in panel I. (C) Connection to odor vial #1 is established by passing the tubing through a screw cap and an insert. Make sure the input tubing is close to the bottom of the odor vial. (D) Using three separate pieces of tubing and a T connector, bifurcate the output from the normally closed port of the 3-way valve, and connect them with two 2-way valves. Each of the 2-way valves will then be connected to odor vials #2 and #3. (E and F) Connect the output from odor vials #1, #2, and #3 to individual ports of the manifold, and then connect the output port with the arena, making sure to use a Y connector to bifurcate the odor flow into two and then connecting each tubing end to diagonally opposed odor inputs in the arena. (G and H) Connect the arena vacuum fitting with a flow meter using a flexible and odor-inert piece of tubing, then connect the flow meter output with the vacuum pump using a separate piece of flexible tubing. (I) Plug the 12 V power supply wires in the MultiBoard, as well as the two valve wires from each valve. A total of six valves will be connected to the board (two 3-way valves and four 2-way valves). (J) Insert six valve drivers in the board, one for each valve, and then use insulated breadboard jumper wires to connect each driver to the Arduino. You will need an extra wire for ground connection. (K) After assembling the odor delivery, place an LED board below the arena and plug the LED controller for illumination. Insert a high-resolution camera and a lens on top of the arena for video recording.
Procedure
Fly preparation
Ten days before the experiment, cross 25–30 virgin D. melanogaster females of the desired genotype A with 10–12 males of the desired genotype B. Prepare at least four bottles containing 40–50 mL of fresh fly food for each group. Keep all the crosses in an incubator at 25 °C, with 60% relative humidity, and a 12/12 h light/dark cycle. Expand the crosses to new bottles every 7 days.
Note: Crosses can also be set at lower or higher temperatures, depending on experimental requirements.
Empty the bottles by discarding any adult fly the night before the collection to make sure only newly enclosed flies will be collected the morning after. LTH can only be induced in newly eclosed flies, no more than 12 h after eclosure [24].
Anesthetize flies using CO2 and sort between males and females of the desired genotype. Transfer 20–25 newly eclosed flies of either sex to fresh food vials and distribute vials among two groups: Control group, to be mock exposed, and Test group, to be odorant exposed (see section B for odorant preparation and section C for training protocol).
Induction of habituation
This protocol was established by exposing flies to either ethyl butyrate (EB) or methyl hexanoate (MH). It is possible that other odorants may be used to induce LTH, but their odor concentrations and efficacies will need to be standardized.
Prepare as many 200 μL PCR tubes as the total number of fly vials collected.
To induce habituation to EB, add 80 μL of paraffin oil (PO) and 20 μL of EB to the PCR tube, to obtain a final concentration of 20% EB. Alternatively, induce habituation to MH and add 20 μL of MH to 80 μL of PO. For the control group (mock exposed), prepare PCR tubes containing 100 μL of PO.
Mix the odorant by vortexing the PCR tube for 10–15 s at ~2,500 rpm. Cut off the lids of the PCR tubes and cover each of them with cling film squares (1.5 × 1.5 cm) (Figure 3A).
Figure 3. Odor preparation setup. (A) In a 200 µL PCR tube, prepare 100 µL of either paraffin oil (PO) for the mock-exposed or 20% ethyl butyrate (EB) in PO for the odor-exposed (steps B1–3). Remove the PCR cap and cover the PCR tube with cling film (step B4). (B) Wrap a piece of copper wire around the PCR tube neck and fold it in order to create a hook on the other end; pierce the cling film 10 times with a needle (step B5). (C) Hang the PCR tube containing the odor inside a fly vial using the copper hook and seal the vial by wrapping parafilm around the cap (step B6).
Using approximately 5 cm of copper wire, wrap the wire around the top of the PCR tube and make a hook with the loose end (Figure 3B). Pierce the cling film with a Microlance needle 21 G × 38 mm approximately 10 times to allow the odor vapor to flow out of the vial.
Tap the flies to the bottom of the vial and quickly hang the PCR tube inside the fly vial using the wire hook (Figure 3C). Close the vial and seal it by wrapping the top of the vial with parafilm.
Transfer vials of each condition, mock and odor, onto the dedicated clear plastic box and put each box in separate incubators at 25 °C and a 12/12 h light/dark cycle. For experiments with TubGal80ts, incubators have to be set to higher temperatures to allow for gene expression during odor exposure.
Note: Keep in mind that the odor will leak from the odor-exposed box; so, it is important to keep the two conditions in separate incubators.
Incubate the flies with the odors for four consecutive days.
On the late afternoon/evening of the fourth day of odor exposure, transfer the flies onto starvation vials (see Recipes) in the absence of odors and incubate overnight in an odor-free incubator at 25 °C with 60% humidity and a 12/12 h light/dark cycle.
Habituation testing
Turn on the temperature of the climate chamber (or any other heating device) to 25 °C and allow it to reach the temperature for 1 h before starting the experiment.
Move the fly vials from the incubator to the testing room and let them acclimatize for 30 min before starting the experiment.
Prepare the odor bottles for the behavioral test by pipetting 20 mL of PO onto six odor vials and label four of them PO and the remaining two EB. Add 20 µL of EB to each EB-odor vial to obtain a final concentration of 1:1,000. For MH exposure, we also used a concentration of 1:1,000. Close the vials and vortex for a few seconds to mix. Remove the caps and connect the odor bottles to the arena by inserting the caps.
Ensure the arena is closed before you turn on the vacuum pump and confirm that the airflow is stable at 200 mL/min for two opposing quadrants (the flow rate should be 100 mL/min on each quadrant).
Turn on the computer and set the infrared light to 25% intensity. We use the command set originally obtained from Janelia Tech-Transfer office to communicate with the LED panel via Bonsai [23] software. This light intensity was empirically determined based on the overall quality of the acquired image and might have to be readjusted in a different apparatus/camera. In such a case, aim for low light intensity to prevent heat-related artifacts.
Adjust camera settings using the camera software. We used an image size of 1,552 × 1,552 pixels while the speed was set to 22 frames per second.
Open the acquisition protocol file (Figure 4). Our protocol was developed using the Bonsai programming language, but other software can be used for running a protocol and interfacing with the different hardware, namely the camera, LED board, and Arduino.
Figure 4. Acquisition protocol workflow. Bonsai workflow describing the different operations during the odor response test. To activate the valves, we communicate with the Arduino and send transistor-transistor-logic (TTL) signals using the Digital Output node. Each Step contains the status of each of the six valves for a given duration. For example, if the first minute of the protocol is composed of air stimulation, then the status of all valves in Step 1 will be 0 (closed) for a specified duration of 1 min. Upon odor stimulation, the valve status of the designated valve should change to 1 (open). We use the Fly Capture node to collect information from the camera and the VideoWriter operator to record the sequence of images collected. A Timestamp node was introduced to timestamp the stream of information, which is then outputted in a csv file via the CsvWriter node.
Direct the airflow to each of the vials for 10 min by toggling the solenoid valves. This will ensure that the headspace of the odor vial is saturated with odor vapor. After priming each of the odor lines, turn off the pump and open the arena lid to start a behavioral test.
Using the fly aspirator, transfer approximately 20 flies of the same genotype from a vial into the arena and immediately slide the glass lid to prevent them from escaping. Close the four-field chamber by pushing down the clamps, turn on the vacuum pump, and cover the apparatus with the blackout cloth. Let the airflow stabilize to 100 mL/min in each quadrant while flies get acclimatized for 3 min and start the test.
The test is composed of three steps, each with 1 min duration (Figure 5). During step 1, a baseline is established by allowing the airflow to be directed toward two PO-containing bottles for 1 min such that all four quadrants are equally stimulated with the solvent-neutral odor. Flies should be equally distributed among the four quadrants during this time.
Figure 5. Habituation testing protocol. During the first minute of the test, flies are exposed to paraffin oil (PO) in all four quadrants to establish a baseline (step 1). To stimulate flies with odor, valves are opened for 1 min in two opposing quadrants (step 2). Finally, all valves are closed to allow the odor to be flushed out for another 1 min (step 3).
Open the valves for 1 min at step 2 to stimulate flies with odor in two opposite quadrants of the olfactory arena. This is achieved by diverging the airflow into one PO bottle to stimulate two opposite quadrants, and into one EB-containing bottle (1:1,000 EB in PO) to stimulate the other two quadrants. The quadrants being stimulated with odor should be flipped in different experiments.
Close the valves at the third and last step of the protocol to allow airflow from PO-containing bottles in all four quadrants, washing out the odor used in step 2 from the arena.
Turn off the pump and carefully collect the flies with an aspirator under red light.
Confirm the existence of two output files at the end of a test—a recorded movie and a csv file containing the timestamps and valve status. They will be required for subsequent analysis.
Generate around 5–8 repeats for each condition.
Cleaning the setup
Clean the bottom of the chamber, the chamber spacer, and the covering glass with water and 70% ethanol between each run. Dry it out with a dry tissue and load a new set of flies.
At the end of the session:
Prepare six bottles with 70% ethanol and allow the airflow to go through all bottles for 5 min to clean the odor and remaining PO from the delivery system.
Discard the ethanol from the bottles and add water to each of them. Perform another round of cleaning by diverging the airflow to each bottle for 5 min.
Clean the bottom of the chamber, the chamber spacer, and the covering glass as indicated in section D point 1.
Dry all components with a dry tissue. Turn off the light, the power supply, and the computer.
Data analysis
Count the number of flies on each quadrant for each frame (or calculate the pixel area on each quadrant for each frame). We processed each acquired movie using a machine vision protocol in Bonsai to automatically extract the number of flies on each quadrant on each frame (Figure 6). This process entails background subtraction and thresholding, followed by counting the number of flies within previously defined regions of interest (ROI). Total pixel area selected after thresholding can also serve as a good measure for the number of flies.
Figure 6. Screenshot of the Bonsai workflow used to analyze videos and calculate preference index. We start by uploading a movie of an empty arena (A) and a movie that we want to analyze (B) (FileCapture nodes). We will use one frame of the empty arena movie to perform background subtraction (Subtract node) followed by conversion of the input image into greyscale (Grayscale node). We then perform thresholding in order to have a well-defined foreground (Threshold node). The threshold value is empirical and may be adjusted depending on the image quality. We next define the areas of each of the two opposing quadrants, as dictated by the boundaries of the four quadrants, using the CropPolygon node. On each of the defined regions, we count the number of flies (C) (FindContours and Count) and calculate the pixel area (D) (Area). Both measures (number of flies per quadrant and pixel area per quadrant) are suitable for calculating a preference index, even though we choose to use the pixel area in our study. Finally, we compute the preference index by performing the arithmetic operations required to calculate the preference index as dictated by the following formula: (n° flies in odor quadrants − n° flies in air quadrants)/total n° flies. The square area depicts the preference index calculation using the number of flies per quadrant and the pixel area per quadrant. The calculated preference index is then loaded in a csv file (E) (CsvWriter), which is subsequently aligned with the valve status information using a MATLAB script.
Calculate the preference index (PI) by subtracting the number (n°) of flies in the air quadrants from the number of flies in the odor quadrants and dividing by the total number of flies:
PI = (n° flies in odor quadrants − n° flies in air quadrants)/total n° flies.
The PI range is between +1 and -1. A positive PI occurs when the animals are attracted to the odor; a negative PI corresponds to repulsion; when PI is 0 the animals are equally distributed across the arena.
We used a single Bonsai workflow to automatically extract the number of flies, calculate the PI, and output a csv file with the PI on each frame. The Bonsai workflow for the analysis is depicted in Figure 6.
Using MATLAB, align the PI data with the valves’ onset and offset. Calculate the overall PI by averaging the PI of the last 30 s of the odor stimulation period. Flies take on average 20–30 s to react and stabilize at their final position upon odor stimulation. The PI of the last 30 s therefore reflects their final behavioral choice.
Calculate the average response index (RI) by reversing the sign of the overall PI (RI = −PI).
Statistical analyses were performed using Graph Pad Prism 8. Both unpaired t-test and 2-way ANOVA are used to compare two or more groups, followed by the Tukey-Kramer test for multiple comparisons.
Validation of protocol
To validate this protocol, we assessed whether wild-type strains, Canton S (CS) and Oregon R (OrR), could exhibit a reduction in the behavioral response to EB after four days of exposure to this odorant.
We observed that odor-exposed animals showed a 50% reduction in behavioral response (Video 1) in comparison to mock-exposed animals (Video 2) (Figure 7).
Video 1. Behavioral response to ethyl butyrate after odor exposure.The movie shows the moments before (step 1) and during odor stimulation (step 2). Notice that odor-exposed flies can be found in quadrants with the control odor (PO), as well as in quadrants with the test odor (EB) (highlighted in red).
Video 2. Behavioral response to ethyl butyrate after mock exposure.The movie shows the moments before (step 1) and during odor stimulation (step 2). Mock-exposed flies readily leave EB-stimulate quadrants (highlighted in red), crossing into quadrants stimulated with control odor (PO).
Figure 7. Wild-type strains Canton S (CS) and Oregon R (OrR) habituated to an odorant after long-term exposure. Naïve flies were either exposed to paraffin oil (PO) as control (mock-exposed, n = 7) or to 20% ethyl butyrate (EB) (odor-exposed, n = 6). Error bars show ± SEM. Two-way ANOVA: CS *p = 0.0451, OrR *p = 0.0264.
To test whether the protocol is functional with other aversive odors, we used another repulsive odor, methyl hexanoate (MH), to perform the training and testing (Figure 8). We observed that wild-type animals (CS) show a 60% decrease in behavioral response after MH-odor exposure in comparison to mock-exposed animals, confirming that this protocol can function with aversive odors other than EB.
Figure 8. Prolonged exposure to methyl hexanoate (MH) results in olfactory long-term habituation (LTH). Canton S (CS) flies were exposed to either paraffin oil (PO) (mock-exposed, n = 8) as control or 20% MH (odor-exposed, n = 8). Their behavioral response after exposure was assessed using 1% MH. Bars represent the average response index (RI) of CS flies. Error bars show ± SEM. T-test. *** p value < 0.0001.
Subsequently, we wanted to test whether the canonical characteristics of habituation can be observed in this behavioral paradigm. The two characteristics tested were spontaneous recovery and specificity [1,2]. We first assessed spontaneous recovery, which looks at whether odor avoidance can be restored in odor-exposed animals after a certain number of days. We observed that animals tested 7 days after odor exposure do not show a decrease in behavioral response as they do at 1 day after odor exposure (Figure 9), arguing that the olfactory habituation undergoes spontaneous recovery.
Figure 9. Avoidance to ethyl butyrate (EB) is restored 7 days after training. Canton S (CS) flies were either exposed to 20% ethyl butyrate (EB) (odor-exposed, n = 7) or to paraffin oil (PO) (mock-exposed, n = 7). Their behavioral response to 1% EB was tested 1 and 7 days after exposure. Error bars show ± SEM. Two-way ANOVA test. 1 day p-value *= 0.0352; 7 days p-value = 0.9915.
Lastly, we addressed the second characteristic of habituation, specificity. To investigate the specificity of decreased response, we exposed animals to EB but tested their behavior with two other different odors—CO2 and 3-octanol—to then confirm that the decreased behavioral response is only exhibited toward the exposed odorant. We observed that the behavioral response of EB-exposed flies was not different from mock-exposed animals when they were tested with CO2 and 3-OCT, suggesting that the behavioral decrease observed is selective for the training odorant (Figure 10).
Figure 10. Habituation is specific for the training odor. Flies were either trained with 20% ethyl butyrate (EB) (odor-exposed) or with paraffin oil (PO) as control (mock-exposed) and tested with either 1% EB (n = 9), 0.2% CO2 (n = 7), or 1% 3-OCT (n = 5). Error bars show ± SEM. Two-way ANOVA EB test p-value *= 0.0141; CO2 p-value ns = 0.863; 3-OCT p-value ns = 0.978.
Discussion and conclusion
Habituation is a form of non-associative learning in which the animal filters out familiar inconsequential stimuli resulting in a reduction of behavioral response after continuous or multiple exposure to a stimulus. However, the exact molecular and neuronal mechanisms underlying habituation remain elusive.
Here, we establish an updated behavioral paradigm for long-term olfactory habituation in free-walking groups of adult Drosophila. Our training protocol consists of a 4-days passive-diffusion odor-training, which generates an olfactory memory that lasts 24 h. This protocol differs from previously established protocols for studying long-term olfactory habituation and overcomes behavioral measurement limitations by allowing quantitative and detailed behavioral measurements obtained by video recording with little interference or manual skill from the experimenter, which might be crucial for the purpose of circuit and molecular mapping.
General notes and troubleshooting
General notes
In this type of behavior, the genetic background of the animal plays a key role in its ability to form habituation. It appears that animals that have a white background are unable to form habituation. For this reason, the fly strains should be in a red-eye background, such as Canton S.
Troubleshooting
Unbalanced airflow in the four quadrants of the arena.
This can happen mainly because of two reasons: 1) Air leaks unexpectedly from connections or Duran bottles, or 2) airflow in one odor line is more difficult than in the other. Ensure that all odor bottles are securely sealed and the connectors are tightened to avoid air leaks. From our experience, the most common source of air leakage is the Duran bottles used to contain the odors. We recommend using PTFE tread seal tape around the neck of the Duran bottles to avoid air leakage. Make sure that the solenoid valves you choose provide the same flow once they are opened to avoid variations in airflow between the two odor lines. Select the solenoids that match because we observed slight differences in airflow between different solenoid valves.
Animals are not showing a decreased behavioral response in line with long-term habituation.
This could be due to the age of the animal when the training protocol started. LTH can only be induced in newly eclosed flies, less than 12 h from enclosure [24]. Make sure to collect just newly eclosed flies.
Acknowledgments
This protocol was adapted and modified from McCann et al. [15]. We thank Jens Hillebrand for help with the video editing and members of the Ramaswami lab for useful discussions. C.R. was funded by the Irish Research Council Postgraduate scholarship. M.R. acknowledges support from funded by a Wellcome Trust-SFI-HRB Investigator grant, an Irish Research Council Laureate Award and an SFI Frontiers for the Future grant.
Competing interests
The authors declare no competing interests.
References
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McCann, C., Holohan, E. E., Das, S., Dervan, A., Larkin, A., Lee, J. A., Rodrigues, V., Parker, R. and Ramaswami, M. (2011). The Ataxin-2 protein is required for microRNA function and synapse-specific long-term olfactory habituation. Proc Natl Acad Sci USA. 108(36): e1107198108.
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Article Information
Publication history
Received: Jul 10, 2024
Accepted: Sep 8, 2024
Available online: Oct 13, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Fluorescent Staining and Quantification of Starch Granules in Chloroplasts of Live Plant Cells Using Fluorescein
SI Shintaro Ichikawa
YK Yutaka Kodama
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5103 Views: 462
Reviewed by: Wenrong HeWenyang LiSriema L. Walawage
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Original Research Article:
The authors used this protocol in Plant Physiology Oct 2023
Abstract
Plants use CO2, water, and light energy to generate carbohydrates through photosynthesis. During daytime, these carbohydrates are polymerized, leading to the accumulation of starch granules in chloroplasts. The catabolites produced by the degradation of these chloroplast starch granules are used for physiological responses and plant growth. Various staining methods, such as iodine staining, have previously been used to visualize the accumulation of chloroplast starch granules; however, these staining methods cannot be used to image live cells and/or provide confocal images with non-specific signals. In this study, we developed a new imaging method for the fluorescent observation of chloroplast starch granules in living plant cells by staining with fluorescein, a widely available fluorescent dye. This simple staining method, which involves soaking a leaf disk in staining solution, shows high specificity in confocal images. Fluorescent images of the stained tissue allow the cellular starch content of living cells to be quantified with the same level of accuracy as a conventional biochemical method (amyloglucosidase/α-amylase method). Fluorescein staining thus not only enables the easy and clear observation of chloroplast starch granules but also allows for precise quantification in living cells.
Key features
• Visualizes chloroplast starch granules stained with fluorescein in living cells.
• Requires only simple specimen preparation with no reagents needed other than the staining solution.
• Fluorescein is readily available worldwide.
• Highly specific method for identifying chloroplast starch granules in confocal images.
• Enables estimation of cellular starch content using fluorescent images.
Keywords: Starch granule Staining Live-cell imaging Fluorescein Chloroplast Confocal microscope
Graphical overview
Background
Plants perform photosynthesis during the day to convert light energy into usable chemical energy. Many plant species store this energy as polysaccharides by accumulating starch granules within chloroplasts. At night, starch granules are broken down into soluble sugars, which are then distributed to sink organs for use in growth and development [1,2]. The products mobilized from starch degradation also mediate osmotic stress tolerance [3] and facilitate stomatal opening in guard cells [4]; thus, the biosynthesis and degradation of chloroplast starch granules are crucial processes in plants.
Several staining methods have been developed for observing chloroplast starch granules in leaves, enabling the quantification and investigation of their morphology. Conventional staining methods such as iodine, toluidine blue, and periodic acid Schiff staining have been widely used [5–7]; however, these staining methods involve multiple procedures and chemical fixation, preventing rapid and/or live-cell imaging. Chloroplast starch granules have also been visualized using fluorescent staining methods, including modified pseudo-Schiff propidium iodide [4,8] and safranin O staining [9], but these methods generate non-specific signals. It is therefore necessary to develop a simple staining method that enables the observation of living cells and shows good specificity for chloroplast starch granules.
To address this need, we recently established a fluorescein staining method for visualizing chloroplast starch granules fluorescently in various living plant cells [10]. Fluorescein staining only requires the submergence of leaf tissue in staining solution for 10 min. Due to its high specificity for chloroplast starch granules, fluorescent images of stained leaf tissue can be used to quantify the cellular starch content. Fluorescein staining is therefore a valuable analytical tool for understanding chloroplast starch granules in various plant species.
Materials and reagents
Biological materials
Arabidopsis thaliana accession Col-0 (Arabidopsis) (see General Note 1)
Reagents
99% ethanol (Japan Alcohol Trading Co.)
Molecular sieves pack 3A (Wako, catalog number: 131-13531)
Fluorescein (Wako, catalog number: 065-00252)
Dimethyl sulfoxide (Wako, catalog number: 048-21985)
Fluorescein diacetate (FDA) (TCI, catalog number: F0240) (see General Note 2)
Solutions
Fluorescein staining solution (see Recipes)
FDA staining solution (see Recipes)
Recipes
Fluorescein staining solution
Reagent Final concentration Amount
10 mM fluorescein in ethanol treated with molecular sieves (100% ethanol) 10 μM 1 μL
Ultrapure water n/a Up to 1,000 μL
The staining solution should be prepared just before use. Molecular sieves were used as dehumidifiers for 99% ethanol following manufacturer’s instructions.
FDA staining solution
Reagent Final concentration Amount
10 mM FDA in dimethyl sulfoxide 10 μM 1 μL
Ultrapure water n/a Up to 1,000 μL
The staining solution should be prepared just before use.
Laboratory supplies
Hole puncher (2.0 mm in diameter) (Natsume Seisakusho Co., catalog number: KN-291-2)
Disposable 10 mL syringe and plunger (Terumo, catalog number: SS-10SZ)
Slide glass (super frost) (Matsunami Glass, catalog number: S024420)
Cover glass, 20 × 50 mm, thickness 0.16–0.19 mm (No. 1S) [Matsunami Glass, catalog number (similar product): C025501]
Microtube, 1.5 mL (WATSON, catalog number: 131-7155C)
Equipment
Confocal microscope (Leica Microsystems, model: Leica TCS SP8X)
Water-immersion lens (Leica Microsystems, model: HC PL APO 63×/1.20 W CORR CS2)
Software and datasets
LAS X (Leica Microsystems)
Fiji/ImageJ (https://imagej.net/software/fiji/downloads)
Procedure
Sample preparation and fluorescein staining
Use a hole puncher to excise a leaf disk with a diameter of 2.0 mm (Figure 1A–B) (see General Note 3).
Add 2–5 mL of ultrapure water and the leaf disk to a syringe using a tweezer and insert the plunger. Be careful not to spill water from the top of the syringe.
Deaerate the leaf disk in water by pulling the plunger while holding the top of the syringe (Figure 1C). Repeat this process several times to adequately deaerate the leaf disk.
Transfer the deaerated leaf disk into a 1.5 mL microtube with 1 mL of 10 μM fluorescein solution using a tweezer and gently invert the tube several times to immerse the leaf disk (Figure 1D).
Incubate the leaf disk for 10 min with one or two gentle inversions of the tube to sufficiently stain the leaf disk.
Mount the stained leaf disk onto a slide glass without rinsing and cover it with a cover glass. Ensure the slide glass is moistened with ultrapure water before mounting the leaf disk (Figure 1E).
Figure 1. Staining and sample preparation. A. Hole puncher used to excise a leaf disk. B. Excision of a leaf disk from an Arabidopsis leaf using the hole puncher. C. Deaeration of a leaf disk using a syringe and plunger. D. Staining of a leaf disk in the fluorescein staining solution. E. Preparation of a wet-mount slide for a stained leaf disk.
Observation of starch granules using confocal microscopy
Place the slide on the confocal microscope and observe the fluorescein fluorescence (which represents stained starch granules) using the settings shown in Table 1 (Figure 2) (see General Note 4).
Table 1. Confocal microscope (Leica TCS SP8X) settings for detection of fluorescein and chlorophyll fluorescence
General condition for fluorescein Chlorophyll
Light source White light laser (WLL) WLL
Laser intensity 21% (30% intensity from the 70% power WLL) 21%
Objective HC PL APO 63×/1.20 W CORR CS2 (water-immersion lens) HC PL APO 63×/1.20 W CORR CS2
Format 100 Hz/512 × 512 pixels 100 Hz/512 × 512 pixels
Detector Hybrid detector (HyD) HyD or photomultiplier tube (PMT)
Excitation 488 nm 488 nm
Emission range 500–550 nm 680–720 nm
Time gating 0.5–1.2 ns Off
Figure 2. Confocal images of fluorescein-stained starch granules in mesophyll and guard cells. Leaf cells of 3-week-old Arabidopsis plants stained with 10 µM fluorescein for 10 min. The green channel displays the fluorescence of fluorescein-stained starch granules, and the magenta channel displays the fluorescence of chlorophyll. Scale bars, 10 μm.
Data analysis
Measurement of the area of the fluorescein signals representing the starch granules
Export confocal images of fluorescein-labeled starch granules as Tagged Image File Format (TIFF) files with a scale bar using LAS X software.
Open the images in Fiji/ImageJ (Figure 3A).
Convert the image format from RGB color to 8-bit to perform the following binarizing process (Image > Type > 8-bit).
Use the Straight tool to draw a line along the scale bar and then set the scale (Analyze > Set Scale) (Figure 3B).
Binarize the image by adjusting the threshold (Image > Adjust > Threshold) (Figure 3C). Visually adjust the threshold value until the signal area of the binarized image closely matches that of the original confocal image (see General Notes 5 and 6).
Use the Wand tool to select the binarized starch granule signals (Figure 3D).
Measure the area of the selected starch granule signals (Analyze > Measure) (Figure 3E) (see General Note 7).
Figure 3. Quantification of the area of chloroplast starch granules stained with fluorescein. A. Fluorescein-stained starch granules in Arabidopsis mesophyll cells irradiated with white light (approximately 50 μmol photons m–2 s–1) for 0, 4, and 8 h after a 24 h dark treatment. B. Setting the scale using the Straight tool in Fiji and the bar in the TIFF image. C. Setting the threshold using 8-bit color images. D. Selecting the starch granule signals with the Wand tool of Fiji. E. Quantifying the area of the selected chloroplast starch granules. All scale bars, 10 μm. These quantitative data are used in Figure 5 and our previous study [10].
Validation of protocol
This protocol or parts of it were used and validated in the following research article:
Ichikawa et al. [10]. Fluorescein staining of chloroplast starch granules in living plants. Plant Physiology (Staining experiments with chloroplast starch granules: Figure 1A, C, F; Figure 3; Figure 4 A; Supplementary Figure S1B, C; Supplementary Figure S2; Supplementary Figure S3; Supplementary Figure S4A; Supplementary Figure S6B; Supplementary Figure S8; Supplementary Figure S9A; and Supplementary Figure S10. Correlation analysis of the area of fluorescent starch granules in chloroplasts and the starch content in leaves: Supplementary Figure S4. Spectrum measurement: Supplementary Figure S6C).
General notes and troubleshooting
General notes
Applicability of fluorescein staining: Fluorescein staining can be used in many plant species including Arabidopsis, Nicotiana benthamiana, soybean (Glycine max), strawberry (Fragaria × ananassa), lettuce (Lactuca sativa), tomato (Solanum lycopersicum), cucumber (Cucumis sativus), and the model liverwort Marchantia polymorpha [10]. In our previous tests, chloroplast starch granules were successfully observed in a variety of land plants using fluorescein staining.
Enhanced fluorescence with FDA: The use of FDA instead of fluorescein greatly enhances the fluorescence intensity of chloroplast starch granules in living cells [10]. The recipe for FDA staining solution is noted in the Recipes section. The protocol for FDA staining is the same as that for fluorescein staining [10].
Leaf disk size: Any size of leaf disk can be used, but a smaller size is preferable for staining and observing leaf cells. A leaf punch is not essential.
Fluorescein fluorescence detection: To accurately detect fluorescein fluorescence in plant cells, we provide the spectrum of excitation and emission wavelengths of fluorescein in the starch granules (Figure 4). Although we employed the confocal microscopy system made by Leica (SP8X), confocal microscopy systems made by other manufacturers can also be used. Furthermore, starch granules stained by fluorescein could be observed using conventional fluorescence microscopy.
Figure 4. Spectrum of fluorescein in the chloroplast starch granules of living cells. Excitation and emission spectra of fluorescein adhering to the chloroplast starch granules in Arabidopsis mesophyll cells were measured at 5 nm intervals using xyΛ and xyλ modes, respectively, within LAS X. The measurement was repeated for 10 starch granules, and relative means are plotted. These data were used in a previous study [10].
Thresholding for binarization: Typically, a constant threshold value or auto-threshold is desirable for binarization [11]; however, these thresholding processes may not precisely delineate the area of the starch granules because fluorescein accumulates not only at starch granules but also in the stroma after the treatment of plant cells [10]. Fluorescence from the stroma represents a non-specific signal, so users should manually adjust the threshold value for each image to achieve optimal delineation.
Manual tracing options: The Polygon selection tool or Freehand selection tool within Fiji/ImageJ can also be used to determine the area of chloroplast starch granules by manual tracing, but we employed the threshold process because it is laborious to manually trace the small fluorescence signals of starch granules.
Correlation with starch content: There is a strong positive correlation (R2 = 0.9995) between the area of chloroplast starch granules and the enzymatically quantified starch amount [10]. This indicates that fluorescein signals from starch granules represent the starch content of leaves (Figure 5).
Figure 5. Correlation between chloroplast starch granule size and the biochemically determined starch content in Arabidopsis leaves. Plots of the starch granule size within chloroplasts (fluorescein imaging) and the leaf starch content (amyloglucosidase/α-amylase method; Millipore Sigma, STA20) at 0, 4, and 8 h of white light illumination (approximately 50 μmol photons m–2 s–1) after a 24 h dark incubation. At each time point, the size of 10 starch granules was measured, and the starch content was measured four times. Average values for both measurements were plotted. These data were used in a previous study [10].
Troubleshooting
Problem 1: Chloroplast starch granules are not observed using confocal microscopy.
Possible cause: Inadequate staining by fluorescein.
Solutions: Improve staining efficiency by using the fluorescein staining solution instead of water during the deaeration step. Extend the staining time; staining for less than 3 h is typically sufficient to visualize chloroplast starch granules in living plant cells [10]. Consider using FDA instead of fluorescein, as it enhances the fluorescence intensity of the chloroplast starch granules.
Problem 2: Chlorophyll autofluorescence interferes with the detection of fluorescein fluorescence.
Solutions: Use the time-gating method to exclude chlorophyll autofluorescence [12]. If time-gating is not available, ensure that the samples are well-stained with fluorescein or FDA to reduce interference. Set the excitation wavelength of fluorescein to a longer wavelength (e.g., 514 nm) to decrease chlorophyll excitation [12]. However, note that this may reduce fluorescence intensity of fluorescein due to non-optimized excitation. Bleached chlorophyll has an elevated autofluorescence [12]; therefore, lengthy irradiation of the specimen by the excitation laser should be avoided.
Acknowledgments
We thank Ms. Akari Masaki (Utsunomiya University, Tochigi, Japan) for preparing the figures.
Competing interests
The authors declare no conflict of interest.
References
Smith, A. M. and Stitt, M. (2007). Coordination of carbon supply and plant growth. Plant, Cell Environ. 30(9): 1126–1149. https://doi.org/10.1111/j.1365-3040.2007.01708.x
Stitt, M. and Zeeman, S. C. (2012). Starch turnover: pathways, regulation and role in growth. Curr Opin Plant Biol. 15(3): 282–292. https://doi.org/10.1016/j.pbi.2012.03.016
Thalmann, M., Pazmino, D., Seung, D., Horrer, D., Nigro, A., Meier, T., Kölling, K., Pfeifhofer, H. W., Zeeman, S. C., Santelia, D., et al. (2016). Regulation of leaf starch degradation by abscisic acid is important for osmotic stress tolerance in plants. Plant Cell. 28(8): 1860–1878. https://doi.org/10.1105/tpc.16.00143
Horrer, D., Flütsch, S., Pazmino, D., Matthews, J. S., Thalmann, M., Nigro, A., Leonhardt, N., Lawson, T. and Santelia, D. (2016). Blue light induces a distinct starch degradation pathway in guard cells for stomatal opening. Curr Biol. 26(3): 362–370. https://doi.org/10.1016/j.cub.2015.12.036
Abt, M. R., Pfister, B., Sharma, M., Eicke, S., Bürgy, L., Neale, I., Seung, D. and Zeeman, S. C. (2020). STARCH SYNTHASE5, a noncanonical starch synthase-like protein, promotes starch granule initiation in Arabidopsis. Plant Cell. 32(8): 2543–2565. https://doi.org/10.1105/tpc.19.00946
Seung, D., Soyk, S., Coiro, M., Maier, B. A., Eicke, S. and Zeeman, S. C. (2015). PROTEIN TARGETING TO STARCH is required for localising GRANULE-BOUND STARCH SYNTHASE to starch granules and for normal amylose synthesis in Arabidopsis. PLoS Biol. 13(2): 1–29. https://doi.org/10.1371/journal.pbio.1002080
Szydlowski, N., Ragel, P., Raynaud, S., Lucas, M. M., Roldán, I., Montero, M., Muñoz, F. J., Ovecka, M., Bahaji, A., Planchot, V., et al. (2009). Starch granule initiation in Arabidopsis requires the presence of either class IV or class III starch synthases. Plant Cell. 21(8): 2443–2457. https://doi.org/10.1105/tpc.109.066522
Truernit, E., Bauby, H., Dubreucq, B., Grandjean, O., Runions, J., Barthélémy, J. and Palauqui, J. C. (2008). High-resolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. Plant Cell. 20(6): 1494–1503. https://doi.org/10.1105/tpc.107.056069
Liu, Q., Li, X. and Fettke, J. (2021). Starch granules in Arabidopsis thaliana mesophyll and guard cells show similar morphology but differences in size and number. Int J Mol Sci. 22(11): 5666. https://doi.org/10.3390/ijms22115666
Ichikawa, S., Sakata, M., Oba, T. and Kodama, Y. (2023). Fluorescein staining of chloroplast starch granules in living plants. Plant Physiol. 194(2): 662–672. https://doi.org/10.1093/plphys/kiad528
Shihan, M. H., Novo, S. G., Le Marchand, S. J., Wang, Y. and Duncan, M. K. (2021). A simple method for quantitating confocal fluorescent images. Biochem Biophys Rep. 25: 100916. https://doi.org/10.1016/j.bbrep.2021.100916
Kodama, Y. (2016). Time gating of chloroplast autofluorescence allows clearer fluorescence imaging in planta. PLoS One. 11(3): 1–8. https://doi.org/10.1371/journal.pone.0152484
Article Information
Publication history
Received: Jun 21, 2024
Accepted: Sep 13, 2024
Available online: Oct 12, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Plant Science > Plant cell biology > Cell imaging
Cell Biology > Cell staining > Carbohydrate
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The on-Site Monitoring and Specimen-Making of Ectoparasites on Rodents and Other Small Mammals
PY Peng-Wu Yin
XG Xian-Guo Guo
WS Wen-Yu Song
WD Wen-Ge Dong
YL Yan Lv
DJ Dao-Chao Jin
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5104 Views: 162
Reviewed by: Zeeshan BandaySatya Ranjan Sahu Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Parasites & Vectors May 2023
Abstract
The ectoparasites of rodents and other small mammals usually involve five categories of arthropods—fleas, sucking lice, gamasid mites, chigger mites, and occasionally, ticks. These ectoparasites are medically important, serving as vectors for diseases such as plague, murine typhus, scrub typhus, forest encephalitis, Lyme disease, and other zoonoses. Field surveys, collection, and specimen preparation of ectoparasites are crucial for studying taxonomy, faunistics, ecology, and epidemiology. They are also essential for vector surveillance. The present protocol summarizes the on-site monitoring and specimen-making of ectoparasites of rodents and other sympatric small mammals. Besides the collection and specimen preparation of small mammal hosts, the protocol describes in detail the collection, fixation, specimen-making, and taxonomic identification of ectoparasites and provides some monitoring indices. The on-site monitoring indices include the host density index and the infestation indices of ectoparasites (prevalence, mean abundance, mean intensity). The methodologies outlined in this protocol provide technical guidance and references for vector monitoring (surveillance) and control.
Key features
• Collection and specimen preparation of small mammal hosts, including rodents (rats, mice, and voles) and other sympatric small mammals (shrews, tree shrews, and pikas)
• Collection, fixation, specimen-making, and taxonomic identification of ectoparasites, including fleas, sucking lice, gamasid mites, chigger mites, and ticks
• On-site monitoring indices—host density index and infestation indices of ectoparasites: prevalence, mean abundance, and mean intensity
Keywords: Rodent Small mammal Ectoparasite Flea Sucking louse Gamasid mite Chigger mite Tick Collection Specimen-making
Graphical overview
Flowchart for collection and specimen-making of ectoparasites and their hosts. A. Process for the collection and specimen preparation of small mammal hosts. B. Collection and fixation of ectoparasites. C. Process for specimen-making for different categories of ectoparasites.
Background
Rodents (rats, mice, and voles) and other sympatric small mammals (shrews, tree shrews, and pikas) often share habitats. They serve as significant infectious sources and reservoir hosts for numerous zoonotic diseases, including plague, murine typhus, scrub typhus, and hemorrhagic fever with renal syndrome [1,2]. These small mammals frequently harbor a multitude of ectoparasites such as fleas, sucking lice, gamasid mites, chigger mites, and occasionally ticks [3,4]. Rodents are of paramount importance among small mammal hosts [5]. Ectoparasites found on rodents and other small mammals act as vectors for a variety of zoonotic diseases, with fleas transmitting plague and murine typhus, ticks transmitting encephalitis and Lyme disease, chigger mites transmitting scrub typhus, and gamasid mites transmitting rickettsialpox. Gamasid mites may also serve as potential vectors for hemorrhagic fever with renal syndrome (HFRS) [6–12]. Although ectoparasites are of medical importance, to date, few sources of literature have provided a systematic and comprehensive protocol for on-site monitoring and specimen preparation of ectoparasites. Based on relevant literature and our research experience on ectoparasites, this protocol systematically describes methods for on-site monitoring and specimen-making of ectoparasites.
Small mammals in this protocol refer to rodents (Order Rodentia) and other sympatric small mammals found in the same habitats. These include insectivores (Order Eulipotyphla or Insectivora), tree shrews (Order Scandentia), and pikas (Order Lagomorpha), excluding flying bats (Order Chiroptera). Small mammals serve as hosts of ectoparasites.
Fleas and sucking lice, two groups of ectoparasites on small mammals, belong to the class Insecta from the phylum Arthropoda [13,14], and their collection and specimen preparation methodologies are similar. Gamasid mites and chigger mites, two tiny categories of arthropods, belong to the subclass Acari, class Arachnida of Arthropoda [15,16], and their collection and specimen preparation methodologies are also similar. Chigger mites are unique among Acari, with only the larval stage acting as ectoparasites; other stages are free-living [17–20]. Ticks predominantly parasitize large and medium-sized mammals, but occasionally, a few tick larvae and nymphs can be found parasitizing rodents and other small mammals [21-23]. Although ticks belong to the subclass Acari of Arachnida [24], their collection and specimen preparation methods differ from those of gamasid mites and chigger mites due to their relatively larger size.
Materials and reagents
Reagents
Cresol solution (Shanghai Hengyuan Biochemical Reagent, catalog number: 7784-24-9)
Potassium aluminum sulfate (Shanghai Hengyuan Biochemical Reagent, catalog number: 7784-24-9)
Arsenic trioxide (Yunnan Tin Co, catalog number: 1327-53-3)
Camphor powder (Shanghai Anpu Experimental Technology, catalog number: 464-49-3)
Xylene (Shanghai Aladdin Biochemical Technology, catalog number: 1330-20-7)
Distilled water
Canadian balsam (Shanghai Macklin Biochemical Technology, catalog number: 8007-47-4), neutral balsam (Shanghai Macklin Biochemical Technology, catalog number: 96949-21-2), or fir balsam (Aibixin Biotechnology, catalog number: 8016-42-0)
Pomegranate oil (Xi’an Rongbai Biological Technology, catalog number: 84961-57-9) or clove oil (Jiangxi Baicao Pharmaceutical, catalog number: 8000-34-8)
Arabic gum (Shanghai Yiji Industrial, catalog number: 9000-01-5)
Formaldehyde (Beijing Qinling Pharmaceutical Technology, catalog number: 79083-29-7)
Glycerin (Shanghai Aladdin Biochemical Technology, catalog number 56-81-5)
10% sodium hydroxide (Shanghai Macklin Biochemical Technology, catalog number 1310-73-2) or potassium hydroxide (Shanghai Aladdin Biochemical Technology, catalog number 1310-58-3) solution
Solutions
5%–10% cresol soap solution (Lysol) (see Recipes)
Alum arsenic preservative (see Recipes)
Hoyer’s solution or Berlese’s solution (see Recipes)
Different concentrations of ethanol (30%, 50%, 70%, 80%, 90%, 95% and 100% anhydrous ethanol)
The different concentrations of ethanol can be prepared by diluting anhydrous ethanol with distilled water. Anhydrous ethanol is a kind of very commonly used reagent with a wide choice of catalog numbers and manufacturers. We use anhydrous ethanol from Wuhan Jisi Instruments & Equipment (catalog number: 64-17-5).
Recipes
5%–10% cresol soap solution (Lysol)
5 mL of cresol solution + 95 mL of distilled water for making 5% cresol soap solution.
10 mL of cresol solution + 90 mL of distilled water for making 10% cresol soap solution.
In a beaker, mix the cresol and water and shake or stir until the cresol is completely dissolved in the water.
Alum arsenic preservative
Potassium aluminum sulfate (powder, accounting for 40% or 70%) + arsenic trioxide (powder, 20%) + camphor powder (40% or 10%). Mix them together.
Hoyer’s solution or Berlese’s solution
Hoyer’s solution (or Hoyer’s medium)
50 mL of distilled water, 30 g of Arabic gum, 200 g of formaldehyde, 20 mL of glycerin.
Berlese’s solution (or Berlese medium)
20 mL of distilled water, 15 g of Arabic gum, 160 g of formaldehyde, 20 mL of glycerin.
Laboratory supplies
Collection and specimen preparation of small mammal hosts
Fieldwork equipment (field tent, backpack, raincoat, non-slip shoes, flashlight)
Disinfection and protection materials: 70% or 75% ethanol (Wuhan Jisi Instruments & Equipment, catalog number: 64-17-5), protective clothing and hat, medical latex gloves, mask
Trapping materials
Rat cage (18 cm × 12 cm × 9 cm, Jiangxi Guixi Li’s Rat Trap Equipment Co., Ltd.)
Food bait (peanuts, corn, cooked meat, fried dough sticks, steamed buns)
Disposable white cloth bag
Recording materials (pen, marker, pencil, collection information record book)
Specimen preparation materials for small mammals [lancet, surgical scissors, tweezers, steel ruler or tape measure, electronic scale, degreased cotton, bamboo sticks, wooden sticks, suture needles, specimen labels, beakers, alcohol lamps (for boiling small animal skulls), ether, alum arsenic preservative]
Host tissue sample collection and preservation materials: Cryopreservation tubes and 95%–100% alcohol
Collection and fixation of ectoparasites
White square plate (large size: 565 mm × 400 mm, small size: 385 mm × 285 mm)
Lidded centrifuge tubes
Ophthalmic forceps
70% or 75% ethanol
Marking pens
Specimen-making of flea and sucking lice
Glass slides
Coverslips
Baking plate or baking rack
Specimen box
Marker pen
Label paper
Specimen-making of gamasid mites and chigger mites
Petri dish
Brush
Slide
Coverslip
Marker pen
Label paper
Specimen-making of ticks
Wide-mouth bottle
Cover glass
Sealing wax
Equipment
Stereomicroscope (Leica, model: DM3000)
Baking oven (Shanghai Yiheng Scientific Instrument, model: DHG-9420A)
Procedure
Collection and specimen preparation of small mammal hosts
Collection of small mammal hosts
Select the survey site and set rat cages: Prior to placing the rat cage, you should observe signs such as food residues, feces, rat trails, and rat holes. Identify areas frequently visited by rodents and strategically place the rat cage along their activity route to enhance the capture rate (Figure 1). Concurrently, conceal the rat cage with branches or weeds and mark its location for easy retrieval.
Choose food baits: This can be flexible according to the specific situation. For trapping house rats and mice in residential areas, choose baits such as cooked meat, fried dough sticks, and steamed buns. For trapping wild rodents (rats, mice, and voles) in outdoor environments, choose baits such as peanuts and corn.
At the chosen field survey site, place rat cages (trap cages) with baits in the afternoon or evening of the survey day to trap rodents and other sympatric small mammals (hosts) in various geographical and ecological environments, including residential areas (mainly indoor habitats) and different types of outdoor habitats such as farmlands, grasslands, shrubs, woodlands and forests [25,26]. Based on our experience, it is advisable to set rat cages between 4:00 PM and 6:30 PM in the afternoon.
In monitoring the density of small mammal hosts, quantitative placement of rat cages is required. In indoor habitats (human houses, poultry sheds, pig pens, cattle stalls, stables, barns, and warehouses), place one cage every 15 square meters (per 15 m2) along the foot of the wall. In outdoor habitats, place all 25 cages in a group in a straight line, with a spacing of 5 m and a row spacing of 20 m. In paddy fields, place cages along the ridges.
The number of cages placed at each survey point should be as uniform as possible to ensure the homogeneity and comparability of the samples from each survey point.
After being placed for three consecutive days, relocate to a new location and repeat the process [27].
Collect the captured small mammal hosts: The following morning, collect the captured rodents and other sympatric small mammal hosts and place them, along with the rat cage, into a white cloth rat bag, ensuring it is securely tied to prevent ectoparasites from escaping (Figure 1). Adhere to the principle of “one host (one small mammal), one bag” when placing host mammals into the rat bag.
Transport the bag to a temporary on-site laboratory for ectoparasite collection.
Prior to reuse, the rat bag must be thoroughly cleaned and disinfected to prevent cross-contamination of ectoparasites between different host mammals [28].
Collect ectoparasites: Rodent hosts considered pests can be euthanized by cervical dislocation or anesthesia (using ether). Afterward, collect their ectoparasites. Small mammal hosts that do not require euthanasia can be anesthetized into a deep coma for ectoparasite collection.
Once collection is complete, release the small mammals back into their natural habitats after the effects of anesthesia have subsided.
After ectoparasite collection, collect tissue specimens (muscle, bone, or internal organ specimens) from the host mammals, if needed. Store these collected tissue specimens in cryopreservation tubes with 95%–100% alcohol or other specialized fixatives for future use (e.g., DNA extraction).
Record the relevant information of the collected host mammals according to Tables 1 and 2.
Critical: Rodents and other small mammals are frequently carriers of numerous pathogens. During the collection and preparation of specimens from these hosts, it is advised to wear protective clothing, hats, medical gloves, and masks to ensure personal safety. The carcasses and internal organs of euthanized rodents should be immersed in a 5%–10% cresol soap solution (Lysol) for disinfection for a period of 24–48 h. Subsequently, they should be buried deep to prevent the spread and transmission of pathogens present in the rodent body [29].
Figure 1. Collection of small mammal hosts. A. Place rat cages (trap cages) at the chosen site. B. Strategically place the rat cage along the activity route of rodents and other sympatric small mammals (hosts) and mark the location for easy retrieval. C. A captured host inside a rat cage. D. Place the animal and the rat cage into a white cloth rat bag, ensuring it is securely tied to prevent ectoparasites from escaping.
Table 1. Basic information registration form for survey and collection sites of small mammal hosts
Date (year/month/day) Collection site name Geographic landscape, habitat, and altitude (m) of the collection site Climate factor records of the collection site
Landscape Habitat Altitude Weather conditions Temperature ℃ Relative humidity % Other
2024/6/11 Dali Flatland Grasslands 2,005 m Sunny 27 ℃ 83% -
2024/6/12 Heqing Mountainous Forests 2,196 m Rainy 26 ℃ 76% -
Note: Geographic landscapes include mountainous and flatland landscapes. Habitats include residential areas (mainly indoor habitats) and different types of outdoor habitats such as farmlands, grasslands, shrubs, woodlands, and forests. Weather conditions include sunny, cloudy, and rainy (drizzle, moderate rain, heavy rain) conditions.
Table 2. Basic information registration form for small mammal hosts
Collector: Host animal identifier: Information recorder:
Host number Collection time and location Host mammal name Host sex and age Host body index measurement Other information
Date Collection site Sex Age Weight (g) Body length (cm) Tail length (cm) Ear height (cm) Hind foot length (cm)
1 2021/6/21 Bingchuan Mus caroli Male Adult 20 95 85 14 18 -
2 2021/6/21 Bingchuan Apodemus chevrieri Female Adult 34 110 90 15 21 -
Note: Collection time and collection site are the same as in Table 1. Host age refers to adult or juvenile.
Preparation of host fur specimens:
Skinning: Euthanize small mammals via cervical dislocation or anesthesia. Under stringent personal protection measures, use surgical lancets and scissors to dissect and fully excise all muscles, bones, and internal organs from the euthanized rodent (dead rodent), ensuring a complete rodent skin is obtained.
Preservation: Uniformly paint an alum arsenic preservative to the inner surface of the rodent skin. This preservative serves to prevent the decay and pest infestation of the animal fur.
Filling or prosthesis support: If a posture specimen is required, first construct a dummy frame using wires, comparable in size and shape to the rodent body. Fill this frame with cotton or other suitable materials. For a general taxidermy specimen, provide support to the feet and tail using bamboo sticks or wooden sticks, filling the remaining parts directly.
Suturing and shaping: Sew the rodent skin using a needle and thread and adjust its shape and posture through a process termed “shaping.”
Fixation: Upon successful creation of the taxidermy specimen, affix the posture specimen to an appropriate platform for ease of display. Accompany each specimen with a label (either label paper or a label card) with information such as the scientific name of the rodent (ordinary name and complete Latin name), collection location and habitat, altitude, collection time, collector’s name, identifier’s name, and identification time (Figure 2, Figure 3A).
Figure 2. Posture specimen (Rattus tanezumi) (modified by Pan et al. [30])
Figure 3. Representative images of small animal host specimens (Rattus tanezumi). A-1. Dorsal view of taxidermy specimen. A-2. Ventral view of taxidermy specimen. A-3. Lateral view of taxidermy specimen. B-1. Overall dorsal view of skull specimen. B-2. Overall ventral view of skull specimen. B-3. Overall lateral view of skull specimen. B-4. View of mandibular tooth surface. B-5. Mandibular side view.
Preparation of host skull specimens
The skull and dental characteristics are important in the precise taxonomic identification of rodent hosts, thereby requiring the preparation of host skull specimens. Typically, rodent skull specimens are prepared using the boiling method:
Place the entire skull of the rodent (rat, mouse, or vole), detached from the stuffed specimen, in a container filled with water and boil it.
Once nearly cooked, remove the skull and eliminate all soft tissues such as muscles and fascia. If any soft tissue remains, return the skull to water (a small amount of sodium carbonate or sodium hydroxide can be added if necessary) and boil until all remaining soft tissue is removed.
Once all soft tissues are removed and the skull is naturally dried, it becomes the host skull specimen (Figure 3-B).
Collection and fixation of ectoparasites
Collection of ectoparasites
The collection procedures and methods for the aforementioned ectoparasites are similar and are thus described collectively here (Figure 4).
To prevent ectoparasites from attacking and biting the operator, anesthetize both the host mammals and the ectoparasites with ether prior to the collection of ectoparasites: place the host mammals, which have been captured on-site and enclosed in a sealed rat bag, into a sealed container (a large plastic bucket with a lid suffices).
Subsequently, add several cotton balls soaked in ether for anesthesia until the host mammals are either deceased or deeply comatose.
Then, place host mammals in a pre-prepared simple device, known as the “double-plate device,” to facilitate the collection of all ectoparasites from each host mammal individually. This device consists of a small white square plate of suitable size placed within a larger white square plate. Add an adequate volume of water to the large square plate and apply insect repellent around the edges of both plates. This special device is designed to prevent the escape of ectoparasites that have not been fully anesthetized by ether or those that have revived after anesthesia during the collection process. Here, the water in the large plate and the insect repellent form two types of barriers.
Use a toothbrush or comb to brush the host mammals from head to tail 2–3 times, aiming to brush the majority of the ectoparasites into the small plate. Combing 2–3 times allow most fleas, sucking lice, and gamasid mites to be obtained.
Use ophthalmic forceps to clamp the parasites that have been brushed into the small plate or a paintbrush dipped in fixative to dip them.
Simultaneously, check the parasites attached to the white rat bag and those that have fallen outside the small plate and into the water of the large plate.
After the combing collection is completed, carefully check the parasites remaining in the host’s fur from head to tail and collect them using the hair-turning method, including ticks that may be tightly attached to the host’s skin.
Only larvae of chigger mites (chiggers) parasitize the body surface of rodents and other small mammals. Chiggers are extremely small and typically attach to the thin skin of the host mammals’ bilateral auricles, the base of the external ear, and the opening of the external ear canal. Prioritize these parts for collection. A simple and practical method for collecting chiggers involves using a magnifying glass and a lancet or a scraper such as an ear spoon to scrape off the chiggers and chigger-like debris (suspected chiggers) attached to the bilateral auricles, ear canals, perianal area, perineum, and groin.
Place collected ectoparasites into a lidded centrifuge tube containing 70% or 75% ethanol for fixation and preservation (see step B2).
Figure 4. Collection and fixation of ectoparasites. A toothbrush or comb is used to brush the small mammal host from head to tail 2–3 times, causing ectoparasites to fall into the double-plate device for collection. B. An ophthalmic forceps is used to collect (lightly “clamp”) parasites from the body surface of the host. C. The parasites attached to the white rat bag are checked. D. The collected ectoparasites are placed into a lidded centrifuge tube containing 70% or 75% ethanol for fixation.
Fixation and preservation of ectoparasites
Adhering to the principle of “one host, one vial”, place all collected ectoparasites into a container filled with 70% or 75% ethanol for fixation and preservation. Depending on the specific situation, containers such as standard glass vials, lidded centrifuge tubes, or Eppendorf tubes can be chosen.
Use an oil-based marker to label the container used for fixation and preservation.
Mark a label paper with a number using a pencil and place it inside the container to prevent confusion among ectoparasites from different hosts.
Note: Labels attached to the outside of the container can occasionally fade, and the pencil label here ensures that the number of samples collected can be accurately identified in the case of the fading of the external label, thus achieving double insurance.
Precautions: Gamasid mites and chiggers are small, with chiggers being particularly tiny. It is usually challenging to recognize chiggers with the naked eye, necessitating the use of a magnifying glass during the collection process. When collecting chiggers, also scrape off and collect the chigger-like debris (suspected chiggers) to prevent loss due to their minuscule size. If molecular biological research on the collected ectoparasites is required, or if there is a need to detect and isolate pathogens from the ectoparasites, fix and preserve collected ectoparasites with 95% ethanol or a special fixative in accordance with specific requirements. If long-term preservation is required, place ectoparasites in liquid nitrogen, which can be stored indefinitely.
Specimen-making of flea and sucking lice
Soaking and washing
Transfer the fleas and sucking lice, which are preserved in 70% or 75% ethanol, into a culture dish filled with clean or distilled water.
Allow them to soak and wash for a duration of 30–60 min, then proceed to remove any debris.
Digestion
Move the thoroughly washed flea and sucking lice into a solution of 5% or 10% potassium hydroxide or sodium hydroxide. This will corrode and digest their soft tissues and body contents.
The digestion process typically takes 1–3 days, depending on the size of specimens and ambient temperature. Larger specimens and lower room temperatures will extend the digestion time.
Neutralization
Transfer the digested flea and sucking louse specimens into clean or distilled water and wash them 2–3 times for 30 min each.
Then, move them into a 5% acetic acid solution and let them soak for 30 min. This step neutralizes the strong alkali (potassium hydroxide or sodium hydroxide) present in the fleas and sucking lice, halting the digestion process.
Finally, transfer them into clean or distilled water and soak and wash them 2–3 times for 30 min each.
Dehydration
Transfer the digested and neutralized specimens into a sequence of ethanol gradients with concentrations of 30%, 50%, 70%, 80%, 90%, 95%, and 100% for gradual dehydration.
Let soak with each ethanol gradient for 30–60 min.
Transparency
Move the dehydrated specimens into a mixture of anhydrous ethanol and xylene (mixed in a 1:1 ratio) and pure xylene for transparency.
Soak for 5–10 min each soaking until the specimens become transparent. Avoid excessive soaking time to prevent over-transparency.
Alternatively, directly transfer the dehydrated specimens into clove oil for a 30–60 min soak to achieve transparency.
Mounting
Place 2–3 drops of lipid-soluble adhesives such as Canada balsam, neutral balsam, or fir balsam on the center of a clean glass slide.
Use a dissecting needle to position the transparent fleas and sucking lice in the adhesive.
Under a stereomicroscope (dissecting microscope), adjust the position of the specimens so that the head end faces backward, and the limbs are extended. Then, cover it with a coverslip.
Ideally, only one specimen should be mounted on each glass slide. If two specimens of the same species are available, they can be mounted on one slide.
Baking
Place the mounted specimens flat on a special baking plate or rack and dry them in an oven at 45–60 °C.
Drying and curing generally takes 24–72 h.
Preservation
Observe the dried slide specimens under a microscope.
After completing taxonomic identification, attach a label to each side of the slide. The label should indicate the host animal collected and its Latin scientific name, collection location, collection time, collector, specimen’s species name and Latin scientific name, identification time, and identifier (Figure 5).
Store the labeled specimens individually in a specimen box in a special specimen cabinet.
Figure 5. Specimen-making of fleas and sucking lice for the taxonomic identification under a microscope. A. The glass slide specimen of a flea with the Latin scientific name, collection location, collection time, and collector. B. A microscopic image of the flea Xenopsylla cheopis (Rothschild, 1903) [31], ♂.
Specimen-making of gamasid mites and chigger mites
Gamasid mites and chigger mites, both belonging to the class Arachnida, Arthropoda, share similar specimen-making steps and methods. Given their small size and soft chitinous exoskeletons, particularly in the case of chiggers, specimen-making typically involves the use of Hoyer’s or Berlese’s solution for direct mounting, with Hoyer’s solution being the more common choice. This method bypasses the need for digestion with potassium hydroxide (or sodium hydroxide) solution and gradual dehydration with an ethanol gradient of varying concentrations, followed by the application of water-soluble Arabic gum for mounting. However, for the few mites that are larger and possess thicker chitinous exoskeletons, the preparation method used for fleas and sucking lice is recommended, which involves digestion and gradual dehydration prior to mounting with lipid-soluble adhesives. The following outlines the steps and methods for the specimen-making of gamasid mites and chiggers:
Soaking and washing
Transfer the gamasid mites and chiggers, which have been fixed and preserved in 70% or 75% ethanol, into a Petri dish containing clean or distilled water.
Soak and wash for 30–60 min, depending on the size of the mites, and remove any debris.
Mounting
Place 2–3 drops of mounting fluid (Hoyer’s solution or Berlese’s solution) on the center of a clean slide.
Use a dissecting needle or brush to transfer the washed gamasid mites or chiggers into the mounting fluid.
Adjust the position under the stereomicroscope so that the head end faces backward, the ventral side faces upward, and the limbs are extended.
Cover with a coverslip.
Mount one specimen per slide; however, two specimens of the same species can also be mounted, one with the dorsal side facing up and the other with the ventral side facing up.
Drying and transparency
Lay the mounted slide specimens flat on a specialized baking plate or rack and bake in an oven at 45–60 °C for 7–15 days or longer until the mounting fluid has dried.
Monitor the process closely. If the mounting fluid shrinks, add more mounting fluid from the side of the coverslip using a dissecting needle.
Once dried and transparent, the specimens can be clearly observed under a microscope for morphological structure examination and taxonomic identification.
Preservation
After the dried slide specimens have been identified under a microscope, attach double labels to each side of the slide following the method described in Section C. Write relevant information on the labels.
Specimen-making of ticks
As mentioned above, ticks are also members of the class Arachnida, Arthropoda, which predominantly parasitize the body surface of large and medium-sized mammals. However, a minority of tick larvae and nymphs can be found in small mammals, such as rodents. Given their size, larger ticks do not typically require slide specimen-making and can be directly preserved in a fixed solution. Smaller ticks, including larvae and nymphs of certain species, can be prepared as slide specimens using the method outlined in Section C.
Immersed specimens
For larger ticks, field-collected specimens can be transferred to a wide-mouth bottle containing 70% or 75% ethanol for long-term preservation.
Seal the bottle cap with sealing wax.
Attach a label detailing the host’s name and Latin scientific name, collection location and time, collector, specimen’s name and Latin scientific name, identification time, and identifier.
Slide specimens
Refer to the method outlined in Section C.
Taxonomic identification of ectoparasites
Specimen preparation
Once the collection, fixation, preservation, and specimen-making of ectoparasites are completed, identify each type of ectoparasites as soon as possible.
Ticks, typically larger in size, can be directly placed under a stereomicroscope for taxonomic identification using fixed and preserved specimens. Fleas, sucking lice, gamasid mites, chiggers, and smaller ticks require slide specimen-making before being placed under the stereomicroscope for taxonomic identification.
Literature preparation for taxonomic identification: Prior to formal identification, it is essential to prepare the necessary identification tools, books, and literature related to taxonomic identification, including taxonomic keys.
Morphological observation
Using the taxonomic tools, books, literature, and taxonomic keys, carefully observe, measure, and compare the relevant morphological structures and characteristics.
Each ectoparasite specimen should be gradually identified according to the taxonomic levels of order, family, genus, and species, with an aim to identify it to the “species” level as much as possible.
Some specimens may be unclear due to the arthropod body being wrapped or contaminated by debris, broken limbs, hair loss, or excessive intestinal contents, and cannot be accurately identified. A small number of specimens may be new species that have not yet been recognized and cannot be identified for the time being. In such cases, they can be recorded as “tentative species” or “unidentified species” in the original data.
Data analysis
On-site monitoring indices for ectoparasites and their hosts
Host density indices.
For small mammal hosts of ectoparasites, it is usually necessary to calculate their density indices. The following indices (area density and cage-day density) can be used to reflect their density:
D
a
=
H
A
;
D
m
=
H
M
c
×
D
Where Da represents the area density, A represents the unit area (such as per square meter, etc.), H represents the number of small mammal hosts, Dm represents the cage-day density, MC represents the unit cage number (such as per 100 cages, etc.), H represents the number of hosts, and D represents the number of days. The area density (Da) is the number of hosts captured per unit area, and the cage-day density (Dm) is the number of hosts captured per unit cage number per day.
Ectoparasite composition ratio and infestation indices.
The constituent ratio (Cr) of ectoparasites is the percentage of the number of individuals of a certain parasite species (or a certain parasite group) to the total number of all parasite species (or all parasite groups) examined. The infestation indices include three indices: prevalence (P), mean abundance (MA), and mean intensity (MI). Prevalence is also known as the infestation rate, and mean abundance is also known as the infestation index. The calculation formula is as follows:
C
r
=
N
i
N
×
100
%
;
P
=
H
i
N
×
100
%
;
M
A
=
N
i
H
;
M
I
=
N
i
H
i
Where Ni is the number of individuals of a certain parasite species (or a certain parasite group), N is the total number of all parasite species (or all parasite groups), Hi is the number of host individuals infested with a certain parasite species (or a certain parasite group), and H is the total number of all host individuals examined.
Seasonal fluctuation curve
To dynamically monitor the seasonal fluctuation of a specific type of ectoparasite, a fixed survey point in the area under observation should be selected. A field survey lasting 5–7 days (one week) should be conducted every month for a year, during which small mammal host trapping and ectoparasite collection should be carried out. Following the taxonomic identification of ectoparasites, the number of ectoparasites, constituent ratio, and infestation indices should be counted monthly. Using a rectangular coordinate system, plot the month as the horizontal coordinate and the number of ectoparasites, constituent ratio, and infestation indices as the vertical coordinates to create the seasonal fluctuation curve of the ectoparasites (see Figure 6 and Figure 7) [32].
Figure 6. Seasonal fluctuation of chiggers (Walchia micropelta) on the rat Rattus brunneusculus at Jingha, southern Yunnan of China. April 2016–March 2017. Cited from Lv et al. [32].
Figure 7. Seasonal fluctuation of chiggers (Ascoschoengastia indica) on the rat Rattus brunneusculus at Jingha, southern Yunnan of China. April 2016–March 2017. Cited from Lv et al. [32].
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Song et al. [48]. Potential distribution of Leptotrombidium scutellare in Yunnan and Sichuan Provinces, China, and its association with mite-borne disease transmission. Parasites & Vectors.
General notes and troubleshooting
For taxonomic identification of small mammal hosts, refer to the following literature: Handbook of the Mammals of the World. Volume 6: Lagomorphs and Rodents I [33], Handbook of the Mammals of the World. Volume 7: Rodents II [34], Catalogue of mammals in China [35], and Taxonomy and distribution of mammals in China [36]. Host tissues (e.g., muscle, liver, or spleen) can be collected from deceased hosts for other research purposes, including DNA extraction of hosts and DNA detection of relevant pathogens (viruses, bacteria, etc.).
For taxonomic identification of ectoparasites, together with the relevant reports on new species, refer to the following literature: The Sucking Lice of North America: An Illustrated Manual for Identification [37], Economic Insect Fauna of China Fasc. 40 Acari: Dermanyssoidese [38], Trombiculid mites of China: studies on vector and pathogen of tsutsugamushi disease [39], Classification and retrieval of sucking lice in China [40], Fleas [41], Ticks [42], A Manual of Acarology [43], Biology of Ticks, Volume 2 [44], Hard Ticks (Acari: Ixodida: Ixodidae) Parasitizing Humans [45], Fleas and Ticks in Small Animals [46], and The Flea [47].
With the exception of ticks, most ectoparasites (fleas, sucking lice, chiggers, and gamasid mites) are too small to be directly observed and identified, and slide specimens must be made before the taxonomic identification under a microscope. Fleas and sucking lice fall within the class Insecta of Arthropoda, and their specimen-making steps and methods are similar. For optimal preservation, it is generally recommended to use lipid-soluble adhesives such as Canada balsam, neutral gum, or fir gum to mount the flea and sucking louse onto glass slides, which ensures long-term or permanent preservation. However, when faced with a large number of fleas and sucking lice collected on-site, achieving permanent mounting becomes challenging. As an alternative, Hoyer’s or Berlese’s solution (containing Arabic gum) can be used for mounting, in which each glass slide can accommodate multiple individuals of fleas or sucking lice.
Lysol is an irritating liquid, and arsenic is a highly toxic substance. Technicians and relevant operators must wear protective clothing, gloves, and masks to prevent skin contact, oral ingestion, or inhalation of the reagents. They must strictly comply with the regulations on the use and management of highly toxic substances in the process of purchasing, preparing, using, and storing arsenic.
Acknowledgments
We’d like to express our sincere thanks to the following people for their kind help and suggestions: Rong Fan, Cheng-Fu Zhao, Zhi-Wei Zhang, and Ya-Fei Zhao. The study was supported by the Major Science and Technique Programs from Yunnan Province (Grant No. 202102AA310055-X) to Xian-Guo Guo, and the expert workstation for Dao-Chao Jin in Dali Prefecture.
Competing interests
The authors declare they have no conflict of interest.
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Article Information
Publication history
Received: Mar 17, 2024
Accepted: Sep 2, 2024
Available online: Oct 22, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Environmental science > Parasites
Biological Sciences > Biological techniques
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PEPITA: Parallelized High-Throughput Quantification of Ototoxicity and Otoprotection in Zebrafish Larvae
EN Elizabeth M. Nilles
EB Ethan Bustad
MQ Meng Qin
EM Emma Mudrock
AG Alden Gu
LG Louie Galitan
HO Henry C. Ou
RH Rafael E. Hernandez
SM Shuyi Ma
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5105 Views: 308
Reviewed by: Ivonne SehringAlberto Rissone Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Pharmacology Mar 2024
Abstract
Drug-induced hearing injury (ototoxicity) is a common, debilitating side effect of many antibiotic regimens that can be worsened by adverse drug interactions. Such adverse drug interactions are often not detected until after drugs are already on the market because of the difficulty of measuring all possible drug combinations. While in vivo mammalian assays to screen for ototoxic damage exist, they are currently time-consuming, costly, and limited in throughput, which limits their utility in assessing drug interaction outcomes. To facilitate more rapid quantification of ototoxicity and assessment of adverse drug interactions that impact ototoxicity, we have developed a high-throughput workflow we call parallelized evaluation of protection and injury for toxicity assessment (PEPITA). PEPITA uses zebrafish larvae to quantify ototoxic damage and protection. Previous work has shown that hair cells (HCs) in the zebrafish lateral line are very similar to human inner ear HCs, meaning zebrafish are a viable model to test drug-induced ototoxicity. In PEPITA, we expose zebrafish larvae to different combinations of drugs, fluorescently label the HCs, and subsequently use microscopy to quantify the brightness of the fluorescently labeled HCs as an assay for ototoxic damage and hair-cell viability. PEPITA is a reproducible, low-cost, technically accessible, and high-throughput assay. These advantages allow many experiments to be conducted in parallel, paving the way for systematic evaluation of drug-induced hearing injury and other multidrug interactions.
Key features
• Analysis of drug-induced hair cell damage associated with ototoxicity using Danio rerio
• Ototoxicity assessment performed in vivo
• Uses microscopy to generate images to assay ototoxicity quantitatively
• Enables testing of various combinations of drugs at various doses to determine toxicity-associated drug–drug interaction outcomes (synergy, antagonism)
Keywords: Zebrafish Ototoxicity Otoprotection Microscopy Automated image analysis Drug–drug interaction
Graphical overview
Overview of PEPITA workflow for fish handling. Day 1 (day before fertilization): Set up crossing and incubate overnight. Day 0 (day of fertilization): Execute crossing, bleach the collected embryos for 10 min, transfer embryos into Petri dishes, and incubate overnight. Day 1 (1 day post-fertilization (dpf)): Transfer viable embryos into new Petri dishes under a microscope and incubate overnight. Day 3–4 dpf: Dechorionate the larvae and incubate overnight. Day 5 dpf: Place a 3D-printed multi-well plate-strainer insert into a large Petri dish and transfer the larvae into each well of the insert. Transfer the larvae-containing insert to its respective drug treatment plate for the desired drug exposure time (this varies depending on the drug and the goal of the experiment; we have used treatment durations of 1 min to 24 h). Following this, transfer the inserts into a plate with hair cell staining solution and incubate (this varies depending on the type of stain; we have used staining durations of 1–20 min), then transfer them into a plate with anesthesia solution and incubate until larvae stop moving. Then, transfer the larvae to a 96-well imaging plate and image whole fish in brightfield and relevant fluorescence channels. Imaging time depends on the instrument and settings; our imaging workflow typically takes ~30 min to image a single 96-well plate. Data analysis: The PEPITA software package accepts brightfield (top) and fluorescence (bottom) images, creates and applies a mask to each fluorescence image to focus analysis on the fish. The package next identifies the top 15 brightest local maxima and creates a second mask around ten puncta (blue circles). Then, it applies the second mask and automatically scores each image according to the relative fluorescence intensity of the identified 10 puncta. Relative fluorescence intensities scored from drug-exposed fish can be compared quantitatively to untreated fish to calculate individual dose-response curves or pairwise drug interaction outcome reports. Figure generated in BioRender.
Background
Hearing loss is one of the most common chronic diseases, affecting approximately 30% of individuals aged 65–74 years old and 50% of individuals more than 75 years old (both hearing and balance disorders) [1]. Drug-induced injury to hearing and balance (ototoxicity) is a significant concern, with over 150 medications associated with ototoxicity [2]. Adverse drug–drug interactions (DDIs) pose a challenge in drug treatment, as interactions among medications can amplify the potential for hearing damage. New approaches for preclinical drug screening to assess ototoxicity and identify DDIs are needed for developing safer therapeutic options.
Zebrafish are a well-established vertebrate model for high-throughput profiling and studying ototoxicity. While their tissues and organs share many similarities with those of humans, zebrafish grow rapidly, produce hundreds of embryos per spawn, and are optically transparent as larvae, making them a powerful preclinical tool. Zebrafish have been utilized to assess drug toxicity in many organs and tissues, including cardiovascular, gastrointestinal, and mechanosensory systems [3]. Within the mechanosensory system, zebrafish possess the lateral line, which contains hair cells (HCs) that are homologous to the sensory HCs found in the inner ear of humans. Previous work has demonstrated that ototoxic drugs (e.g., aminoglycoside antibiotics and chemotherapeutics) induce similar HC damage and death in both zebrafish lateral lines and human inner ears [4,5]. Established lateral line damage quantification methods involve confocal microscopy of individual HC clusters (i.e., neuromasts) and manual HC counting. While these approaches provide subcellular resolution on damage phenotypes, they are effort-intensive and thus limited in the number of neuromasts, fish, and conditions that can be profiled per experiment. DDIs quantification would not be tractable with standard neuromast-centric characterization methods.
Here, we describe a high-throughput, semi-automated method for drug screening to assess ototoxicity in vivo using zebrafish, named parallelized evaluation of protection and injury for toxicity assessment (PEPITA). This method is tailored for large-scale drug screening and investigation of DDIs in zebrafish larvae. PEPITA enables efficient quantification of lateral line phenotypes using whole-fish fluorescence microscopy of hundreds of fish per day. Fish can be imaged in multi-well plates after being treated with multiple drugs and fluorescent dyes that report on the phenotype of stained HCs. For example, to measure HC viability, we have used the fluorescent stain oxazole yellow with propidium iodide (YO-PRO-1). Because YO-PRO-1 can only enter through HCs with functioning mechanotransduction channels, it gives a qualitative proxy for HC viability [6,7]. Alternatively, myo6b::gfp fish (transgenic fish that express GFP in their HCs) can be used in place of YO-PRO-1 stained fish as a more cost-effective option for assessing ototoxic damage that also enables longitudinal assessment. PEPITA also includes a software package for quantitatively analyzing images to study the dose-response and DDIs characteristics of individual and pairwise treatment conditions based on the neuromast-specific fluorescence intensity of chemical stains or transgenically expressed fluorescent proteins. Collectively, PEPITA enables semi-automated quantification of lateral line phenotypes measurable with fluorescence readouts. The workflow is amenable to simultaneous quantification of multiple fluorescence channels, and the underlying image analysis pipeline is amenable to adaptation for quantifying toxicity in other zebrafish organs and tissues with fluorescence-based assays.
Materials and reagents
Biological materials
AB strain zebrafish (Danio rerio) (Zebrafish International Resource Center, European Zebrafish Resource Center, China Zebrafish Resource Center)
Tg ; myo6b::gfp zebrafish [8]
Reagents
Note: Azithromycin and neomycin are listed as examples of antibiotics used, but any drugs of interest can be used here.
Azithromycin dihydrate (Thermo Fisher, catalog number: AAJ6674006) (store at 4 °C)
Neomycin sulfate hydrate (Thermo Fisher, catalog number: AAJ6149914)
Texas red-X succinimidyl ester (Thermo Fisher, catalog number: T20175) (store at –20 °C)
Tricaine (3-aminobenzoic acid ethyl ester, methanesulfonate salt) (Pentair, catalog number: TRS5)
YO-PRO-1 Iodide (Thermo Fisher, catalog number: Y3603) (store at –20 °C)
Instant Ocean (Spectrum Brands Product, catalog number: SS15-10)
Bleach (Clorox, catalog number: 30966)
Reverse osmosis (RO) filtered water
Solutions
Antibiotic super-concentrated stock solution (see Recipes for neomycin example)
YO-PRO-1 stain solution (2 μM) (see Recipes)
Tricaine: stock solution (15 mM) (see Recipes)
Tricaine: euthanasia solution (1.35 mM) (see Recipes)
Tricaine: anesthesia solution (0.675 mM) (see Recipes)
ICS water (see Recipes)
Recipes
Neomycin super-concentrated stock solution
Note: This is an example of an antibiotic solution. The super-concentrated stock solution concentrations and solvents used will vary between drugs. Store at -20 °C or -80 °C.
Reagent Final concentration Quantity or Volume
Neomycin sulfate hydrate 200 mM 36 mg
Molecular-grade water 200 μL
Total 200 μL
YO-PRO-1 solution (2 μM)
Note: The recipe is for one 12-well plate, using 3,500 μL of YO-PRO-1 solution per well. Adjust volumes as needed, depending on the size of the multi-well plate being used. Store at -20 °C.
Reagent Stock concentration Final concentration Quantity or Volume
YO-PRO-1 Iodide 1 mM 2 μM 84 μL
ICS water (Recipe 6) N/A 42 mL
Total 42 mL
Tricaine: stock solution (15 mM)
Reagent Stock concentration Final concentration Quantity or Volume
Tricaine powder N/A 15 mM 400 mg
Nano-pure water N/A 97.9 mL
1 M Tris or 1 M sodium bicarbonate 1 M 21 mM 2.1 mL
HCl as needed to reach pH of ~7
Total ~100 mL
Store at -20 °C.
Tricaine: euthanasia solution (1.35 mM)
Reagent Stock concentration Final concentration Quantity or Volume
Tricaine: stock solution (Recipe 3) 15 mM 1.35 mM 4.5 mL
ICS water (Recipe 6) N/A 45.5 mL
Total 50 mL
Tricaine: anesthesia solution (0.675 mM)
Note: The recipe is for one 12-well plate, using 3,500 μL of tricaine solution per well. Adjust volumes as needed, depending on the size of the multi-well plate being used.
Reagent Stock concentration Final concentration Quantity or Volume
Tricaine: euthanasia solution (Recipe 4) 1.35 mM 0.675 mM 25 mL
ICS water (Recipe 6) N/A 25 mL
Total 50 mL
ICS water
Note: Any type of “fish water” or embryo medium safe for use with zebrafish embryos and larvae can be used in place of ICS water. However, bear in mind that differing water composition may cause hair cell sensitivity to damage to vary.
Reagent Final concentration Quantity or Volume
Instant Ocean 0.25 g/L 5.0 g
Calcium chloride 0.05 g/L 1.0 g
Sodium bicarbonate 0.07 g/L 1.3 g
Reverse osmosis (RO) filtered water 20 L
Total (optional) 20 L
Laboratory supplies
Note: Supplies indicated with an asterisk are brand-specific because the Tinkercad design for the multi-well plate-strainer inserts may not fit multi-well plates from other manufacturers.
Breeding tanks with dividers (Carolina Biological Supply, catalog number: 161937)
12-well plates, non-treated* (Corning Costar, catalog number: 07-201-589)
24-well plates, non-treated* (Corning Costar, catalog number: 07-201-590)
96-well plates, non-treated, black, flat-bottom, for imaging (Thermo Fisher, catalog number: 37000-550)
3D printer filament, PLA (Amazon, catalog number: B084XS2TS9)
Nylon mesh, 125 micron (ELKO Filtering Co., catalog number: 03-125/39)
Wide-bore Pasteur pipettes* (Fisherbrand, catalog number: 13-678-30)
Note: Wide-bore Pasteur pipettes are necessary because the embryos will not fit into standard Pasteur pipets.
100 μm cell strainers (Corning, catalog number: 07-201-432)
Petri dishes (100 mm × 15 mm) (Falcon, catalog number: 08-757-100D)
Large Petri dishes (150 mm × 15 mm) (VWR, catalog number: 25384-326)
Compressed air duster cleaner (Inovera, catalog number: IVR10012)
Fine-tipped tweezers (Dumont Tweezer, Style 5, catalog number: 0203-5-PO)
Equipment
All-in-one fluorescence microscope (Keyence, model: BZ-X710)
3D printer (Dremel, model: Digilab 3D45 3-D printer)
Hot plate (Fisher Scientific, Thermix Stirring Hot Plate, model: 210T)
Vortex mixer (VWR, catalog number: 10153-838)
Digital dry bath (BioRad, catalog number: 1160562)
Software and datasets
Tinkercad design for multi-well plate-strainer inserts (freely available at https://www.thingiverse.com/thing:5676024)
PEPITA-tools software package can be found on GitHub at https://github.com/ma-lab-cgidr/PEPITA-tools
ImageJ (version 1.54f, 06/29/2023)
Procedure
Multi-well plate strainer insert preparation
Note: Movie S1 shows how steps 4–11 of this procedure are executed.
3D-print the Tinkercad design for the appropriately sized multi-well plate strainer insert. We used the Dremel Digilab 3D45 3-D printer.
Set a hot plate to medium heat and let it heat up.
While heating, cut rectangles of nylon mesh, approximately 9 cm × 12 cm.
Fold a piece of aluminum foil over itself so that it forms a double-layer piece of foil that is slightly bigger than the nylon mesh pieces.
Once at temperature, place the double layer of aluminum foil on the hot plate. Place a rectangle of nylon mesh on top of the aluminum foil.
Immediately press the 3D-printed multi-well insert into the nylon mesh, applying enough pressure for the bottom of the insert to melt onto the nylon mesh (Figure 1a, b). Apply pressure for 3 s, then quickly press around the edge of each well to form a complete seal between the bottom of the 3D-printed insert and the nylon mesh (Figure 1c). Immediately remove the foil-nylon-insert contraption from the hot plate.
Figure 1. Procedure for adhering the nylon mesh to the 3D-printed multi-well plate insert. (A) Place the 3D-printed insert on top of nylon mesh and aluminum foil. (B) Firmly press the 3D-printed insert onto the mesh for 3 s. (C) Press around the perimeter of each well to form a complete seal around the base of the well with the nylon mesh.
Once off the hot plate, immediately press the insert into the nylon mesh for 3 more seconds on the benchtop to further adhere the 3D-printed insert to the nylon mesh. Press around the edge of each well to further solidify the seal.
Gently peel off the foil from the insert.
Check to ensure the bottom of each well of the insert forms a complete seal with the nylon mesh. If necessary, repeat steps A6–8 several more times until this is achieved.
Note: When repeating steps A6–8, it may only be necessary to selectively apply pressure to just the wells of the insert with incomplete seals.
Once each well is completely sealed by nylon mesh, use small scissors to cut away the excess nylon mesh remaining between wells.
Feel around the bottom of each well to ensure that the seal with the mesh has not been compromised during the trimming process. It may also be helpful to gently poke the bottom of the well with the eraser end of a pencil to ensure the seal is intact.
If necessary, repeat steps A6–8 for the wells that need repair.
Prepare a container of 10% bleach solution and completely submerge the completed multi-well inserts in the bleach solution for at least 20 min and no longer than 24 h.
Prepare a container of distilled water. Transfer the multi-well inserts from the bleach solution to the distilled water and let soak for up to 24 h.
After soaking, remove the multi-well inserts from the distilled water and let air dry. Store in a clean plastic bag until ready to use.
Zebrafish embryo preparation
Day –1: Set up crosses (day before fertilization)
Select healthy zebrafish from a stock population and place into breeding tanks—females on one side of each tank and males on the other.
Day 0: Water change, execute crossing, collect embryos (day of fertilization)
When the tank room lights turn on, promptly refresh the water in the breeding tanks, remove the breeding dividers, and execute crossing. Observe fish until they start spawning, and clear, viable embryos are produced.
When spawning is complete, prepare a bleach solution in a beaker: 100 μL of bleach, 150 mL of ICS water (do this before straining eggs). Bleaching the eggs helps to remove mites and other potential biological contaminants from the water.
Recover all embryos from the tank by straining out embryos with a strainer. Rinse embryos into bleach solution using an ICS water squirt bottle. Add ICS water until the final volume of water is 170 mL.
Leave embryos in the bleach solution for 10 min. Swirl eggs intermittently.
After 10 min, strain the embryos in the bleach solution by pouring the bleach solution with embryos back through the strainer into the sink.
Rinse embryos in the strainer thoroughly with ICS water.
Partition embryos into Petri dishes. Aim for ~100 embryos per Petri dish (for enough dissolved oxygen and enough water to dilute nitrogenous waste being produced). Use as many Petri dishes as needed. Using a dissecting microscope, remove all unviable embryos (usually cloudy or malformed). Incubate overnight at 28.5 °C.
Day 1: Sort embryos (1 day post-fertilization, dpf)
Under a dissecting microscope, find all viable embryos and transfer them with a wide-bore Pasteur pipette into new Petri dishes with new ICS water. Aim for ~50 (no more than 60) embryos per Petri dish.
Place Petri dishes in an incubator overnight at 28.5 °C.
Day 3–4: Dechorionate (3–4 dpf)Note: By 4 dpf, most larvae will be hatched already and will not need to be manually dechorionated.
For larvae that have not yet emerged, remove their chorions using fine-tipped tweezers.
Note: While we use manual dechorionation, alternative methods that do not damage hair cells, such as enzymatic dechorionation, are also viable options and can be used for this purpose.
Place remaining larvae (in Petri dishes) in an incubator overnight at 28.5 °C.
Drug treatment and imaging (5 dpf)
Sanitize lab space with ethanol.
Prepare appropriate working concentrations of the drugs of interest. Transfer appropriate drug treatments into their respective wells in a multi-well plate. Label each plate.
Caution: Make sure to read the Safety Data Sheet for the given drug and follow appropriate PPE and disposal guidelines.
Make sure to check the solubility of the given drug in water before starting this. Some drug classes, like macrolides, are notoriously insoluble in water. So, having a baseline of solubility can help guide what doses to choose.
Pour larvae into a cell strainer and place in a Petri dish (100 mm × 15 mm) with ICS water.
Place the appropriate multi-well plate-strainer insert in a large Petri dish (150 mm × 15 mm) full of water. Transfer the appropriate number of larvae from the cell strainer into each well of the insert. (Maximum 100 larvae per 35 mL [9]; up to 11 larvae per well for a 12-well plate and up to 6 larvae per well for a 24-well plate.)
Look at the number of larvae needed for each well, and add n+1 (an extra) larva to each well to account for accidental pipetting damage (e.g., if 10 larvae are being exposed to a given treatment, add 11 larvae to that well).
Once each well of the insert has the appropriate number of larvae, transfer the larvae-containing insert into its respective drug treatment plate.
Incubate the larvae in the drug treatment for the desired drug exposure time.
While larvae are in drug treatment, warm the ICS water to 28 °C in an incubator and thaw the Tricaine stock.
One hour before the larvae come out of drug treatment, turn off lights and thaw YO-PRO-1 solution (YO-PRO-1 is light sensitive). Make sure the YO-PRO-1 stock is covered with foil while in storage and while using.
Note: If using myo6b::gfp, then skip this step (step C9), as well as steps C11–C12, C16–C17.
Prepare Tricaine: euthanasia solution (see Recipes) and Tricaine: anesthesia solution (see Recipes). Transfer Tricaine: anesthesia solution into each well of a new multi-well plate. Place the multi-well plate with Tricaine: anesthesia solution in a 28 °C incubator until the end of the drug treatment.
Use 3,500 μL per well for a 12-well plate and 1,800 μL per well for a 24-well plate.
Prepare YO-PRO-1 solution (see Recipes) in a glass bottle or 50 mL tube covered in foil.
Transfer YO-PRO-1 solution into each well of a new multi-well plate, cover with foil, and place in a 28 °C incubator until the end of drug treatment.
Use 3,500 µL per well for a 12-well plate and 1,800 μL per well for a 24-well plate.
Fill a tub (any type of water-proof receptacle that can hold > 50 mL of ICS water and with dimensions > 130 mm × 90 mm; a new large Petri dish > 130 mm in diameter can also be used here) with about 1 cm (no higher than the height of each multi-well strainer insert) of fresh ICS water for washing.
After the desired length of drug treatment, promptly remove the drug treatment plate from the incubator.
Dunk each multi-well strainer insert into the same ICS washing tub mentioned in step C13 three times to wash off residual drug treatment, ensuring that the larvae stay in their respective wells. (See Movie S2 for a video showing the transfer of a strainer insert between containers of solution to a reagent-containing multi-well plate. During experiments, each strainer will contain zebrafish larvae.)
Transfer the multi-well strainer inserts into the plate with YO-PRO-1 solution. Cover with foil and incubate for 20 min.
After 20 min, remove the insert from the YO-PRO-1 solution and dunk it into the ICS washing tub 3× to wash off residual YO-PRO-1 stain.
Transfer to the plate with Tricaine: anesthesia solution.
Wait about 5 min or until the larvae stop moving. Then, transfer larvae one by one to an untreated 96-well imaging plate using a P200 with a cut tip set at 75 μL.
Using a regular P200 tip, remove 32 μL of liquid from each well.
Image larvae using a fluorescence microscope with a robotics-assisted stage. For example, we use a Keyence model BZ-X710 microscope.
Use the compressed air duster to clean the bottom of the 96-well plate.
Select High Quality (14-bit) if using Keyence.
Set one point for each larva, making sure to focus on the head of each larva (focus near eye area when in plane, focus near ear area when out of plane).
Capture 2× images.
Note: Microscope settings should be optimized for each individual experiment to ensure capturing clear and well-focused images (refer to Figure 5B for an example). For imaging AB larvae on the Keyence, we use the following reference settings for three fluorescent channels: Brightfield: 1/7500s exposure; 50% light intensity; 80% aperture stop.
Note: PEPITA can effectively quantify lateral line phenotypes from images of larvae from a broad range of orientations. Variations associated with differences in positioning can be accounted for with the inclusion of sufficient replicate numbers to yield technically reproducible results.
After imaging, transfer extra larvae into empty wells of the imaging plate, then add about 200 μL of Tricaine euthanasia solution to each fish-containing well to euthanize.
After at least 10 min in Tricaine: euthanasia solution, put the plates in a 20 °C freezer for secondary euthanasia (please adhere to the specific guidelines outlined in your animal protocols for proper euthanasia, as approved practices may vary).
Data analysis
Image analysis and ototoxicity quantification are performed using the PEPITA software package, implemented in Python, which can be found on GitHub (https://github.com/ma-lab-cgidr/PEPITA-tools). This package contains two primary sets of scripts:
“pipeline.py” and related Python scripts: These scripts process fluorescence microscopy images to generate numeric values and plots, including image scores and dose-response curves. These scripts analyze fluorescence data through the following steps, illustrated in the Graphical Overview:
Accepts brightfield and fluorescence images as input.
Identifies and masks fish larvae in each image by contrast, size, and shape from the brightfield image to exclude noise present in the image away from the immediate proximity of the fish.
Adjusts for autofluorescence using a non-signal-containing channel.
Identifies the top 15 brightest local maxima within the masked region, discards the top 5 to exclude potential noise (e.g., small plastic artifacts that often appear as the brightest points), and creates a second mask around the other ten puncta.
Scores each image based on the sum of the pixel values of the masked image.
Standardizes the image scores based on experimental control condition(s) [negative control(s), e.g., untreated larvae and positive control(s), e.g., larvae treated with a dose of ototoxic drug known to elicit complete hair cell ablation] for each plate. These control conditions should be included in the design and setup of the drug dilution plates and imaging plates.
Fits dose-response curves for single treatments and/or interaction parameters for combination treatments.
ImageJ Macro scripts in ImageJ_scripts/: these can help with the creation of mask files, an optional step in the quantification workflow (see “Mask fish larvae” section below).
A comprehensive introduction to using this package is provided in “README” on GitHub. This method was also described in [10] (Section 2.3). The package was developed to be used as a command-line program or a Python library invoked either in a Notebook or in custom Python scripts. Below is a step-by-step protocol for data analysis using the PEPITA package both with the CLI locally and with a provided Notebook.
Package Installation
For command-line execution:
Clone the “PEPITA-tools” repository in GitHub.
Install Python if not already installed. Visit https://www.python.org/downloads/ for more information.
Ensure script dependencies are installed with requirements.txt. This can be done by running the following command: python -m pip install -r requirements.txt
For Notebook environment:
Note: Here is an example of PEPITA running in Notebook: https://colab.research.google.com/github/ma-lab-cgidr/zebrafish-quantification/blob/master/interactive_pipeline.ipynb. Users are encouraged to download and edit this notebook for ease of use.
Run the code under “Get GitHub repository” in the example notebook to download the latest zebrafish pipeline code from GitHub and print out the script version being used.
Run the following command to ensure all the dependencies listed in the requirement file are installed: !pip install -r /tmp/zebrafish-quantification-master/requirements.txt.
Import pipeline and all necessary packages to run the pipeline.
Getting relevant data
Change configurations: configure necessary settings to match the experiment being analyzed.
For command-line execution: set the log_dir setting in the config-ext.ini file.
i. A default configuration can be found at https://github.com/ma-lab-cgidr/PEPITA-tools/blob/master/config.ini. This file defines both the available configuration settings and their default values.
ii. Create a new config-ext.ini file in the repository directory if one does not yet exist. Set the desired values within it to override default values.
Note: Each configuration setting is explained under the “Configuration” section in “README.”
iii. Set this setting to a location where you have write privileges and where logging information can be conveniently written. If the supplied path does not exist, a new directory will be created.
For Notebook environment:
Edit the downloaded notebook under “Making Configuration Changes,” following the guidance provided in comments.
Prepare image data: Save the brightfield and fluorescent microscopy images in a location accessible to this script.
For command-line execution:
Create a folder locally containing all the images you want to analyze.
For Notebook environment:
i. Connect the Notebook to Google Drive.
ii. Create a folder containing all images that you want to analyze.
[optional] Mask fish larvae
Note: This should only be done if particular images need to be excluded or require custom masking. By default, the PEPITA-tools pipeline will analyze each brightfield image, identifying a fish larva and drawing a mask around it. If a pre-existing mask is provided, the pipeline will use it instead of generating a new one.
Custom masks are commonly necessary in any of the following situations:
i. Excluding an image where a larva is partly off the image or extremely out of plane, such that the object detection algorithm fails to identify it.
ii. Excluding non-lateral line portions of larvae if transgenic strains of zebrafish being used harbor fluorescence outside of the lateral line or if damaged tissues in larvae distal from the lateral line cause non-specific staining with vital dye. For example, we exclude the inner ear for myo6b::gfp strains expressing fluorophores in all HCs.
Using ImageJ macros is a feasible way to mask images. Some are provided in the GitHub repository under ImageJ_scripts/. For detailed information, please refer to the ImageJ section in “README.”
Prepare CSV plate template.
A plate template must be supplied to provide meaningful labels to the images being analyzed by the system. The pipeline will apply labels to images by matching them up in order from the top left to the right and then down.
An example template file is available at raw.githubusercontent.com/ma-lab-cgidr/zebrafish-quantification/master/examples/plate-layout.csv.
Run the pipeline
Run the analysis pipeline with appropriate runtime arguments, providing the package with experiment-specific data and context. Details on how to use these arguments can be found under the “Runtime arguments” section in “README.”
Validation of protocol
This protocol has been used and validated in the following research article:
Bustad et al. [10]. In vivo screening for toxicity-modulating drug interactions identifies antagonism that protects against ototoxicity in zebrafish. Front Pharmacol.
We have validated that the PEPITA workflow produces reproducible and reliable results for single-drug screening and drug interaction studies. Its robust performance ensures consistent outcomes, making it a valuable tool for high-throughput ototoxicity assessments and the investigation of adverse drug–drug interactions.
PEPITA enables the quantification of hundreds of individual fish per experiment. To evaluate the system’s ability to produce granular and reproducible results, we tested the viability of neomycin dose-response curves obtained by PEPITA. YO-PRO-1-fluorescently labeled AB fish and myo6b::gfp fish were treated with neomycin at concentrations ranging from 0 to 128 μM for 4 h, followed by imaging and quantification of relative residual brightness using PEPITA. Figure 2 shows the dose-response measurements of 27 YO-PRO-1-labeled AB fish experiments and 7 myo6b::gfp fish experiments, plotted with estimated log-logistic dose-responses (see Figure S1 for the experimental data plotted with experiment-specific dose-response curves). The 95% confidence intervals of the EC50 (drug dose that elicited 50% HC damage) estimated across each of the fish lines were within a 2-fold dose value (2.526–3.113 μM estimated from YO-PRO-1-stained AB fish and 9.0–19.4 μM estimated from myo6b::gfp fish; EC50 values were estimated from log-logistic curves fitted to the experimental data points). PEPITA enables approximately 80% power to detect 2-fold changes in HC damage with 10 biological replicates (fish) per condition, for experienced users.
Figure 2.Overview of the variability in neomycin dose-response curves obtained by PEPITA. (A) YO-PRO-1-stained AB fish (27 experiments). (B) myo6b::gfp fish (7 experiments). Each point represents a different condition measured in a different experiment. The line and shaded area indicate the log-logistic curve and 95% confidence interval of the estimated dose-response.
To validate the accuracy and reliability of PEPITA, we compared PEPITA’s ototoxic dose-response characterization with the gold-standard approach of counting HCs from individual neuromasts imaged by confocal microscopy. YO-PRO-1-fluorescently labeled AB fish and myo6b::gfp fish were treated with neomycin at concentrations ranging from 0 to 20 μM for 4 h. The neomycin dose response was then quantified using both PEPITA and the standard approach. The neomycin dose-response curve derived by PEPITA, based on relative fluorescence units (RFU), closely matched the estimates calculated by manual HC counting in both fish lines, yielding estimated EC50s that were within 2-fold in value (Figure 3, Figure S2).
Figure 3. Neomycin dose response in YO-PRO-1-stained AB fish. (A) Top: Representative neomycin dose-response curve generated by PEPITA with relative fluorescence units (RFU) (EC50 = 1.4 μM). Middle and bottom: Representative image of a fish exposed to no drug (middle) and exposed to 2.5 μM neomycin (bottom), both used for PEPITA quantification. (B) Top: representative neomycin dose-response curve generated by standard approach of counting HCs from individual neuromasts (EC50 = 2.0 μM). Middle and bottom: Representative image of an individual neuromast from a fish exposed to no drug (middle) and exposed to 2.5 μM neomycin (bottom), used for HC counting. Note that PEPITA uses whole-fish images for quantification; images have been cropped for better visualization of the stained neuromasts. Scale bars: fish images = 300 μm; neuromast images = 10 μm.
After confirming the reliability of single-drug dose response using PEPITA, we further validated the feasibility of investigating drug–drug interactions by screening ototoxic interactions with combinations of drug treatments. Four drug combinations with neomycin were examined and scored using PEPITA to assess the potential ototoxic drug interactions (Figure 4). Negative wEOB scores indicate antagonism between neomycin and the tested drugs. Benzamil showed antagonism with neomycin, which is consistent with its known role as a potent mechanoelectrical transduction channel inhibitor that inhibits neomycin uptake. Neomycin interactions with other aminoglycosides had wEOB scores close to zero, indicating additivity, which is consistent with the drugs of the same chemical class exhibiting shared modalities of HC damage.
Figure 4. Drug interaction scores [represented as windowed excess over Bliss (wEOB), x-axis] between neomycin and other drugs screened for ototoxic drug interactions, indicated in y-axis. wEOB is calculated for each well exposed to both drugs using a similar equation as excess over bliss (EOB) metric [11], with the formula EOB = Ra × Rb -Rab, where Ra and Rb are the responses of the individual drug dose exposures, and Rab is the response of the combined drug dose exposure. Wells with expected and observed responses outside 10%–90% were excluded from the aggregated calculation of the score across dose combinations for a particular drug pair, and the remaining data were averaged to yield the composite wEOB interaction score, intended to represent the overall trend of interaction between two drugs across a range of dose combinations.
General notes and troubleshooting
General notes
Using transgenic fish
Users are encouraged to use transgenic fish in place of or in addition to wild-type counterparts. For example, the myo6b::gfp strain of zebrafish expresses GFP in their HCs and can be used to assess HC viability without YO-PRO-1 staining. If using myo6b::gfp zebrafish, the protocol is the same as for AB wild-type zebrafish, except that any steps involving YO-PRO-1 stain can be omitted (steps C9, C11–12, C16–17). The myo6b:: gfp fish embryos will also need to be screened for fluorophores between 1 and 4 dpf after the larvae begin expressing GFP. To execute fluorophore screening, inspect the embryos under a fluorescent microscope and remove any embryos that appear dim or do not fluoresce at all. It is worth noting that using cohorts of either purely homozygous or purely heterozygous larvae for experiments is recommended to reduce technical variance in fluorescence intensity.
It should also be noted that transgenic fish may have different drug tolerances than their wild-type counterparts, so doses may need to be adjusted accordingly [12]. In addition, transgenic fish may be less fecund than wild-type fish. To compensate for this, try any of the following techniques to increase fecundity: adding plastic plants to breeding tanks, elevating one end of the breeding tank to form a slope, setting up a single, large “group spawn” in a big tank, and outcrossing with wild-type fish. If none of these techniques increase fecundity, a large number of crosses should be set up to ensure that at least some eggs will be produced.
Using fluorescently conjugated drugs
Users are encouraged to use fluorescently labeled antibiotics or drugs to enable the use of PEPITA to quantify lateral line uptake of the drug. Fluorescently conjugated drug stock solutions should be prepared at the same molar concentrations as their non-fluorescing counterparts but should be kept in the dark because they are light-sensitive and will become photo-bleached if stored under direct light. Turn off the lights and cover any drug-containing aliquots, tubes, and plates with foil when working with fluorescently conjugated drugs. When imaging, capture the additional fluorescent channel that targets the conjugated fluorophore, in addition to the channel capturing the fluorophore to assay HC viability (e.g., the GFP channel, when using YO-PRO-1 or a GFP-expressing recombinant strain). For example, if using neomycin labeled with TexasRed, put a TexasRed filter cube in the microscope and add the TexasRed channel when imaging. Make sure the excitation and emission spectra of each assayed fluorophore are sufficiently non-overlapping to facilitate concurrent image capture.
Troubleshooting
Problem 1: Drug solubility.
Possible cause: Low solubility of certain drug classes in water (e.g., macrolides).
Solution: If the drugs are not going into solution and/or the working concentration is cloudy, try preparing the solution in a 5 mL centrifuge tube and vortexing vigorously. If this does not work, try adding heat via a dry bath. As a last resort, titrate nonpolar solvents, such as DMSO, into the working concentration until the drug goes into solution. However, it is important to note that DMSO concentrations above 1% can harm developing zebrafish larvae [13], so caution should be exercised.
Problem 2: “Halo” effect on the imaging plate.
Possible cause: Inappropriate volume of water in the well.
Solution: If the view of the larva is obscured due to a shadow around the inner ring of the well (see Figure 5A), double-check to make sure the 96-well imaging plates are non-treated. Using cell culture or tissue culture-treated plates will cause the water to pull more strongly against the sides of the well and produce a shadow over the top of the larva. If using non-treated plates and the issue persists, try adjusting the volume of water in the well. The protocol states to transfer the larva into the well with 75 μL of water, then remove 32 μL of water, resulting in a final volume of 43 μL. Try adjusting the final volume of water to be slightly more, perhaps around 50 μL. Be mindful of adding too much water, though, as this will increase the risk of larvae not resting flat at the bottom of the well. If the shadowing issue persists, try gently pushing excess droplets of water sticking to the sides of the wells to the bottom of the wells, being careful not to disturb the larvae. Gently tapping the plate two to three times on a benchtop can also help flush down excess water into the bottom of the wells. An example of an ideal larva-containing well is shown in Figure 5B.
Figure 5. Example of improper and proper view of larvae in the 96-well imaging plate. (A) Halo ring effect obscuring view of larva. The volume of water in this well is inappropriate, resulting in a shadow cast toward the inside of the well that hinders the view of the fish. (B) Clear image of the zebrafish. Using an appropriate volume of water in the well results in a clear, unobstructed view of the larva.
Acknowledgments
The authors declare financial support was received for the research, authorship, and publication of this article. This study was supported by the National Institute of Health (Grant: R21 DC018341) and the Lura Cook Hull Trust. The original research paper in which this protocol was described and validated was published in Front. Pharmacol. (2024), DOI: 10.3389/fphar.2024.1363545. We gratefully acknowledge Mackenzie Phan, Natalie Gleason, Rina Yan, Patricia Wu, Andrea McQuate, and David Raible for their technical advice and assistance. The Graphical Overview Figure was generated in BioRender.com.
Competing interests
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Ethical considerations
All animal work was approved by the Seattle Children’s Research Institute Institutional Animal Care and Use Committee (protocol number ACUC00658).
References
Ries, P. W. (1994). Prevalence and characteristics of persons with hearing trouble: United States, 1990–91. Vital Health Stat. 10(188): 1–75. https://pubmed.ncbi.nlm.nih.gov/8165784/
Lanvers-Kaminsky, C., Zehnhoff-Dinnesen, A. A., Parfitt, R. and Ciarimboli, G. (2017). Drug-induced ototoxicity: Mechanisms, Pharmacogenetics, and protective strategies. Clin Pharmacol Ther. 101(4): 491–500. https://doi.org/10.1002/cpt.603.
Raldua, D. and Pina, B. (2014). In vivo zebrafish assays for analyzing drug toxicity. Expert Opin Drug Metab Toxicol. 10(5): 685–697. https://doi.org/10.1517/17425255.2014.896339.
Ton, C. and Parng, C. (2005). The use of zebrafish for assessing ototoxic and otoprotective agents. Hear Res. 208(1–2): 79–88. https://doi.org/10.1016/j.heares.2005.05.005.
Harris, J. A., Cheng, A. G., Cunningham, L. L., MacDonald, G., Raible, D. W. and Rubel, E. W. (2003). Neomycin-induced hair cell death and rapid regeneration in the lateral line of zebrafish (Danio rerio). J Assoc Res Otolaryngol. 4(2): 219–234. https://doi.org/10.1007/s10162-002-3022-x.
Santos, F., MacDonald, G., Rubel, E. W. and Raible, D. W. (2006). Lateral line hair cell maturation is a determinant of aminoglycoside susceptibility in zebrafish (Danio rerio). Hear Res. 213(1–2): 25–33. https://doi.org/10.1016/j.heares.2005.12.009.
Ou, H. C., Santos, F., Raible, D. W., Simon, J. A. and Rubel, E. W. (2010). Drug screening for hearing loss: using the zebrafish lateral line to screen for drugs that prevent and cause hearing loss. Drug Discov Today. 15(7–8): 265–271. https://doi.org/10.1016/j.drudis.2010.01.001.
Hailey, D. W., Esterberg, R., Linbo, T. H., Rubel, E. W. and Raible, D. W. (2017). Fluorescent aminoglycosides reveal intracellular trafficking routes in mechanosensory hair cells. J Clin Invest. 127(2): 472–486. https://doi.org/10.1172/JCI85052.
Aleström, P., D’Angelo, L., Midtlyng, P. J., Schorderet, D. F., Schulte-Merker, S., Sohm, F. and Warner, S. (2020). Zebrafish: Housing and husbandry recommendations. Laboratory Animals. 54(3): 213–224. https://doi.org/10.1177/0023677219869037.
Bustad, E., Mudrock, E., Nilles, E. M., McQuate, A., Bergado, M., Gu, A., Galitan, L., Gleason, N., Ou, H. C., Raible, D. W., et al. (2024). In vivo screening for toxicity-modulating drug interactions identifies antagonism that protects against ototoxicity in zebrafish. Front Pharmacol. 15: 1363545. https://doi.org/10.3389/fphar.2024.1363545.
Berenbaum, M. C. (1989). What is synergy? Pharmacol Rev. 41(2): 93–141. https://pubmed.ncbi.nlm.nih.gov/2692037/
Monroe, J. D., Manning, D. P., Uribe, P. M., Bhandiwad, A., Sisneros, J. A., Smith, M. E. and Coffin, A. B. (2016). Hearing sensitivity differs between zebrafish lines used in auditory research. Hear Res. 341: 220–231. https://doi.org/10.1016/j.heares.2016.09.004.
Hoyberghs, J., Bars, C., Ayuso, M., Van Ginneken, C., Foubert, K. and Van Cruchten, S. (2021). DMSO Concentrations up to 1% are Safe to be Used in the Zebrafish Embryo Developmental Toxicity Assay. Front Toxicol. 3: 804033. https://doi.org/10.3389/ftox.2021.804033.
Supplementary information
The following supporting information can be downloaded here:
Movie S1: Multi-well strainer instruction video
Movie S2: Plate transfer instruction video
Figure S1: Dose response data measuring ototoxicity of neomycin across experiments
Figure S2: Neomycin dose response in myo6b::gfp fish
Article Information
Publication history
Received: Jul 14, 2024
Accepted: Sep 13, 2024
Available online: Oct 12, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,106 | https://bio-protocol.org/en/bpdetail?id=5106&type=0 | # Bio-Protocol Content
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Accurate Measurement of Cell Number–Normalized Differential Gene Expression in Cells Treated With Retinoic Acid
NW Nina Weichert-Leahey *
MZ Mark W. Zimmerman *
AB Alla Berezovskaya
AL A. Thomas Look
BA Brian J. Abraham
(*contributed equally to this work)
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5106 Views: 392
Reviewed by: Shivaprasad H. SathyanarayanaYoshihiro Adachi Anonymous reviewer(s)
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Bio-protocol journal peer-reviewed
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Original Research Article:
The authors used this protocol in Science Advances Oct 2021
Abstract
Genome-wide gene expression analysis is a commonly used method to quantitatively examine the transcriptional signature of any tissue or cell state. Standard bulk cell RNA sequencing (RNA-seq) quantifies RNAs in the cells of the tissue type of interest through massive parallel sequencing of cDNA synthesized from the cellular RNA. The subsequent analysis of global RNA expression and normalization of RNA expression levels between two or more samples generally assumes that cells from all samples produce equivalent amounts of RNA per cell. This assumption may be invalid in cells where MYC or MYCN expression levels are markedly different and thus, overall mRNA expression per cell is altered. Here, we describe an approach for RNA-seq analysis of MYCN-amplified neuroblastoma cells during treatment with retinoic acid, which causes dramatic downregulation of MYCN expression and induces growth arrest and differentiation of the cells. Our procedure employs spiked-in RNA standards added in ratio to the number of cells in each sample prior to RNA extraction. In the analysis of differential gene expression, the expression level of each gene is standardized to the spiked-in RNA standard to accurately assess gene expression levels per cell in conditions of high and low MYCN expression. Our protocol thus provides a step-by-step experimental approach for normalizing RNA-seq expression data on a per-cell-number basis, allowing accurate assessment of differential gene expression in cells expressing markedly different levels of MYC or MYCN.
Key features
• High levels of MYC and MYCN expression in cancer cells cause substantial increases in the levels of overall mRNA expression per cell.
• RNA-seq using control RNAs spiked-in on a per-cell basis more accurately reflects global expression changes, when comparing cell populations with substantially different MYCN expression levels.
• In MYCN-amplified neuroblastoma, retinoic acid dramatically decreases MYCN expression levels, resulting in large changes in overall RNA expression levels per cell.
Keywords: RNA-sequencing Spiked-in RNA Transcriptional amplification MYCN Gene regulation Retinoic acid Neuroblastoma Differentiation
Graphical overview
Spike-in-controlled RNA sequencing in MYCN-amplified neuroblastoma
Background
Transcriptional dysregulation is a driver in the development of malignancy. MYC family members, e.g., MYCN, MYC, and MYCL, are powerful oncogenes that promote tumorigenesis in a wide variety of human tissues [1–6]. Elevated expression of MYC family members can occur through several mechanisms, including amplification of the gene locus, chromosomal translocation with enhancer hijacking, and new enhancer formation through DNA variation [1,6–10]. High expression of MYC family members is associated with rapidly proliferating tumors and poor patient outcomes [11–14]. MYC proteins are helix-loop-helix (bHLH) transcription factors that form heterodimers with MAX and, in normal cells, primarily bind the core promoters of actively transcribed genes [15–17]. Studies in multiple cancer types have demonstrated that high levels of MYC or MYCN increase the binding of MYC proteins to the enhancers and promoters of actively transcribed genes, effectively regulating the entire expressed genome [18–20]. Thus, the predominant effect of high MYC expression levels by cancer cells is increased expression levels of genes actively transcribed by the cells, commonly called “transcriptional amplification” [21].
Transcriptional amplification of the inherent cellular gene expression program through high expression of MYC family members plays a critical role in tumor initiation for many tissue types, including pediatric neuroblastoma. Neuroblastoma is a tumor of the peripheral sympathetic nervous system that originates from neuroblasts of the migratory neural crest. Approximately 20% of high-risk tumors have genomic amplification of MYCN, often as double-minute chromatin bodies, leading to hundreds of copies of MYCN per cell [2]. In an independent subset of high-risk neuroblastoma, MYC is highly expressed via enhancer hijacking [6]. In our previous study [22], we showed that MYCN-amplified neuroblastoma cells undergo growth arrest and differentiation upon treatment with retinoic acid, which leads to a change in the core regulatory transcriptional circuitry with changes in cell state, accompanied by downregulation of MYCN expression. In this study, we assessed differential gene expression between neuroblastoma samples with high and low MYCN expression due to retinoic acid treatment.
To analyze cell states and transcriptional programs, standard RNA sequencing measures the relative abundance of RNAs in the tissue type of interest through massive parallel sequencing of RNA-derived cDNA [23]. Standard analysis of genome-wide RNA expression between two or more samples generally assumes that cells from each sample produce equivalent amounts of RNA per cell. However, previous studies have demonstrated that this analysis algorithm can lead to erroneous interpretations when MYC or MYCN acts as a transcriptional amplifier [18,19,24]. In the case of cancer cells with high levels of expression of MYC or MYCN, each cell produces 2–3× more RNA per expressed gene [18,21,24]. Retinoic acid–treated neuroblastoma cells downregulated MYCN, so we expected 2–3 fold less total RNA per cell [18], potentially confounding genome-wide expression analysis and necessitating a tailored experimental strategy. Here, we describe a step-by-step experimental and computational approach using spiked-in RNA standards from the External RNA Controls Consortium (ERCC) that allowed us to compare each gene's expression in a pool of cells in the context of the effect of downregulated MYCN. Our method can be used for a wide variety of other cancer tissues in which transcriptional amplification through any MYC family member comes into play and profoundly impacts the assessment of cancer cell gene expression signatures.
Materials and reagents
Biological materials
BE2C cells (American Type Culture Collection, CRL-2268)
Reagents
Retinoic acid/ATRA (all-trans retinoic acid) (Sigma-Aldrich, catalog number: R2625)
Dimethylsulfoxide (DMSO) (American Type Culture Collection, catalog number: 4-X)
RPMI medium (Thermo Fisher, catalog number: 11875-135)
External RNA Controls Consortium (ERCC) spike-in mix 1 (Thermo Fisher, catalog number: 4456740)
TRIzol (Invitrogen, catalog number: 15596018)
RNeasy mini kit (Qiagen, catalog number: 74104)
RNase-free DNase kit (Qiagen, catalog number: 79254)
Chloroform (Thermo Fisher, catalog number: J67241.AP)
Isopropanol (Thermo Fisher, catalog number: T036181000CS)
Ethanol (Thermo Fisher, catalog number: T038181000)
DNase (Qiagen, catalog number: 79254)
DMSO-Dimethylsulfoxide tissue culture grade (ATCC, catalog number: 4-X)
Library preparation kit: Illumina® Stranded Total RNA Prep, ligation with Ribo-Zero Plus (catalog number: 20040525)
Library quantification kit: Agilent TapeStation 4200 D1000 ScreenTapes (Agilent, catalog numbers: G2991AA, 5067-5582, 5067-5583, 5067-5602, 5067-5586)
FBS (Sigma-Aldrich, catalog number: F2442)
Trypsin 0.05% EDTA (Thermo Fisher, catalog number: 25300-120)
Trypan blue 0.4% (Gibco, catalog number: 15250061)
Solutions
ATRA, 10 mM (see Recipes)
75% ethanol (see Recipes)
Recipes
ATRA, 10 mM
ATRA was purchased as a dry powder. To achieve a final concentration of 100 mM, dissolve 50 mg of ATRA in 1.6 mL of DMSO. Bring the powder into solution by vortexing for 30 s. To bring to a final concentration of 10 mM, dilute the 100 mM stock solution 1:10 by following the below dilution.
Reagent Final concentration Quantity or Volume
ATRA 100 mM 0.1 mL
DMSO n/a 0.9 mL
Total 10 mM 1 mL
75% ethanol
Reagent Final concentration Quantity or Volume
Ethanol 100% 7.5 mL
H2O n/a 2.5 mL
Total 75% 10 mL
Laboratory supplies
6-well plates, polystyrene microplates (Falcon, catalog number: 353046)
Filtered IsoTipTM tips (universal fit racked pipet tips) (Corning, catalog numbers: 4823, 4808, 4809)
15 mL tubes, high clarity PP centrifuge tube (Falcon, catalog number: 352096)
1.5 mL tubes (Eppendorf, catalog number: 022363204)
Glass pipettes (Pasteur Pipets, Fisherbrand, catalog number: 13-678-20C)
Equipment
Countess 3 automated cell counter (Invitrogen, catalog number AMQAX2000)
Refrigerated centrifuge (Eppendorf, model: 5415R)
Spectrophotometer (used for quantification of extracted RNA) (Nanodrop, model: ND-1000)
Qubit fluorometer (Thermo Fisher, catalog number: Q33238)
TapeStation 4200 (Agilent, model: 4200)
Illumina NovaSeq 6000
Software and datasets
ERCC spike-in sequences:
https://tsapps.nist.gov/srmext/certificates/documents/SRM2374_Sequence_v1.FASTA
Reference gene list as GTF, e.g., here, RefSeq:
ftp://ftp.ensembl.org/pub/grch37/current/gtf/homo_sapiens/Homo_sapiens.GRCh37.87.chr.gtf.gz
Burrows-Wheeler-based alignment tools, e.g., here, hisat2 [25]
https://academic.oup.com/bioinformatics/article/20/3/307/185980?login=false
Relevant genome reference FASTA files, e.g., here:
https://hgdownload.soe.ucsc.edu/goldenPath/hg19/bigZips/hg19.fa.gz
Read quantification software, e.g., here, htseq [26]
R package affy [27]
Procedure
Cell preparation
Seed 0.1 × 106 BE2C cells in 6-well plates in enough wells for each replicate plus at least one additional sample for RNA extraction test run (see section B) (i.e., triplicate plus one extra, n = 4). The same number of cells must be seeded in each well for this protocol to be conducted. Keep cells maintained at 37 °C.
Start treatment of BE2C cells with either 5 μL of DMSO as control or 5 μM ATRA.
Change 3 mL of RPMI with DMSO or 5 μM ATRA every 3 days.
Grow BE2C cells under the required experimental conditions for six days. At this point in the experiment, BE2C cells should be 70%–90% confluent in DMSO control samples, and BE2C cells treated with 5 μM ATRA are now expected to appear differentiated with neurites outgrowing.
At the experimental endpoint (= day 6), aspirate the RPMI, wash 1× with 3 mL of PBS, and add 1 mL of trypsin to each well. Place cells back into the incubator until they detach and collect them by adding 3 mL RPMI. Collect each well into a separate 15 mL collection tube and spin the cells down at 800× g for 5 min. Use these conditions for all centrifugation steps unless otherwise specified. Discard the supernatant and wash the cells one time with 5 mL of PBS. Resuspend cells in 1 mL of PBS into a 1.5 mL tube each.
For each sample, combine 20 μL of the resuspended cells with 20 μL of 0.4% trypan blue. Mix well. Pipette 10 µL of this mix into a Countess chamber slide and insert the slide into the Countess 3 automated cell counter. Document the viable cell count per milliliter for each sample. Calculate the absolute viable cell count by multiplying by the total volume for each sample. Record this absolute viable cell count number for each sample. Cell count should be approximately the same across replicates.
Note: It is absolutely necessary to record the absolute cell count number for each treatment condition. Other approaches to obtaining the absolute cell count per sample, e.g., using a hemocytometer, are appropriate in this step as well.
Pellet the cells, remove the supernatant, and snap freeze in dry ice with ethanol. Alternatively, cell pellets can also be snap-frozen in liquid nitrogen to minimize any unintentional changes to gene expression.
Store samples at -80 °C until ready to proceed. If needed, the protocol can be paused and put on hold here.
RNA extraction test run
For RNA extraction delivering high yields of purified RNA, we adapted a hybrid TRIzol+ RNeasy purification method. In the first step, the phenol-based lysis achieves maximal extraction of cellular mRNA from each sample. In the second step, the column purification using the RNeasy mini kit reduces residual ethanol to a minimum, a contaminant that can have adverse effects on library preparation and high-throughput sequencing.
Begin with only the extra fourth sample collected for each experimental group (save the other three samples from each treatment condition to be processed as outlined in section C). Resuspend the frozen cell pellets in 1 mL of TRIzol reagent and incubate at room temperature for 5 min.
Add 0.2 mL of chloroform using a glass pipette, mix for 15 s, and incubate for 2 min at room temperature.
Centrifuge the samples at 12,000× g for 10 min at 4 °C.
Transfer the aqueous (upper) phase to new tubes and precipitate the RNA from the aqueous phase by adding 0.5 mL of isopropanol (per 1 mL of TRIZOL reagent) and briefly vortexing.
Incubate samples at room temperature for 10 min and centrifuge at 12,000× g for 10 min at 4 °C.
Remove the supernatant, wash the RNA pellet once with 1.0 mL of 75% ethanol, and mix by pipetting up and down.
Centrifuge at 12,000× g for 5 min at 4 °C and remove as much of the supernatant as possible. A small amount of residual ethanol is okay; there is no need to air-dry the pellet.
Dissolve the RNA pellet in 100 µL of RNase-free water, add 350 µL of lysis buffer RLT (RNeasy kit), and mix by pipetting up and down.
Add 250 µL of ethanol (100%) to the diluted RNA and mix by pipetting up and down.
Apply the sample to an RNeasy extraction column placed in a 2 mL collection tube and centrifuge at 12,000× g for 1 min at room temperature.
Prepare DNase (for each sample) by adding 10 µL of DNase to 70 µL of RDD buffer (both from the RNase-free DNase kit) and keeping it on ice. DNase is very sensitive to physical denaturation; therefore, mix gently by pipetting up and down.
Add 350 µL of wash buffer 1 RW1 (RNeasy kit), spin at 12,000× g for 15 s at room temperature, and discard the flowthrough.
Add 80 µL of DNase mix to the column (ensuring all solution reaches the membrane) and incubate at room temperature for 15 min.
Add 350 µL of wash buffer 1 RW1 to each column (with DNase solution still in it), spin the column at 12,000× g for 15 s at room temperature, and discard the flowthrough.
Add 500 µL of wash buffer 2 RPE (RNeasy kit) onto the column and centrifuge at 12,000× g for 15 s at room temperature to wash the column.
Add another 500 µL of wash buffer 2 RPE to the RNeasy column and centrifuge at 12,000× g for 2 min at room temperature.
Transfer the column to a new 1.5 mL collection tube, add 30 µL RNase-free water, and centrifuge at 12,000× g for 1 min at room temperature to elute total RNA.
Quantify and record the total amount of RNA in each sample using Nanodrop (i.e., 100 ng/µL × 30 µL = 3 µg total). If needed, the protocol can be paused and put on hold here.
RNA extraction and ERCC spike-in
Based on the sample with the highest amount of extracted RNA, calculate the volume of ERCC spike-in mix 1 to add to each sample.
First, serial dilute the ERCC spike-in mix 1. It is essential to do this fresh every time (Table 1):
Table 1. Preparation of ERCC spike-in mix 1
Dilution ERCC spike-in mix 1 Nuclease-free water
1:10 1 µL undiluted 9 µL
1:100 1 µL of 1:10 9 µL
1:1,000 1 µL of 1:100 9 µL
1:5,000 2 µL of 1:1,000 8 µL
Next, determine the volume of diluted ERCC spike-in mix 1 to add to the sample with the highest amount of extracted RNA. See Table 2 for calculations.
Table 2. Calculations of ERCC spike-in volume
Total RNA Volume of diluted ERCC spike-in mix 1
10 ng 1 µL (1:5,000 dilution)
100 ng 2 µL (1:1,000 dilution)
1,000 ng 2 µL (1:100 dilution)
5,000 ng 1 µL (1:10 dilution)
Example 1:
BE2C DMSO control sample yields 418 ng of RNA. According to the above table, prepare 1:1,000 ERCC spike-in mix and make the following calculations:
2 µL ERCC (1:1,000 dilution)/100 ng RNA = x µL ERCC (1:1,000 dilution)/418 ng RNA
→ x = 8.36 µL ERCC (1:1,000 dilution) for 418 ng RNA.
Example 2:
BE2C DMSO control sample yields 5,500 ng of RNA. According to the above table, prepare 1:10 ERCC spike-in mix and make the following calculations:
1 µL ERCC (1:10 dilution)/5,000 ng RNA = x µL ERCC (1:10 dilution)/5,500 ng RNA
→ x = 1.1 µL ERCC (1:10 dilution) for 5,500 ng RNA.
Example 3: BE2C DMSO control sample yields 40,000 ng of RNA. According to the above table, prepare 1:10 ERCC spike-in mix and make the following calculations:
1 µL ERCC (1:10 dilution)/5,000 ng RNA = x µL ERCC (1:10 dilution)/ 40,000 ng RNA
→ x = 8 µL ERCC (1:10 dilution) for 40,000 ng RNA.
Given the high costs of the ERCC spike-in mix, one might consider setting up an experiment with fewer cells.
For all other (here, 3) samples, adjust the amount added based on that sample’s cell count (that is, not based on their RNA concentration). For example, DMSO control wells each contained 1.0 × 106 cells at the time of collection and are expected to yield ~1,000 ng of total RNA (based on step B18). ATRA-treated wells each contained 0.8 × 106 cells at the time of collection and are expected to yield ~500 ng of total RNA (based on step B18). Accordingly, add 2 µL of 1:100 diluted ERCC to each DMSO control sample and 1.6 µL of 1:100 diluted ERCC to each ATRA-treated sample (to adjust for 20% fewer cells). If the cell count were the same across treatment groups (i.e., 1.0 × 106 cells collected from both), one would add 2 µL of 1:100 diluted ERCC to every sample. DO NOT adjust the volume of ERCC based on changes in RNA yield since this would mask changes in the global RNA output. The spike-in amounts must be tied to the number of cells. See Note 1.
Resuspend the experimental frozen cell pellets in 1 mL of TRIzol reagent and incubate at room temperature for 5 min.
Add the calculated volume of appropriately diluted ERCC spike-in mix 1 to each sample by adding it directly to the cells resuspended in TRIzol solution. It is critical to add the diluted ERCC spike-in mix 1 early in the purification (i.e., directly to the TRIzol before precipitation) since RNA yield tends to become variable between samples following each subsequent step of the extraction procedure.
Extract and quantify RNA from each sample by repeating steps B2–18 and store spike-in-added RNA samples at -80 °C until ready to proceed. The initial sample used for determining total RNA yield and optimizing ERCC concentration can be discarded or saved for an alternative analysis. See Notes 2 and 3.
Library preparation and sequencing
Note: For this step, samples can be processed using any standard RNA-sequencing method.
Samples should be prepared using commercially available library preparation kits, typically starting with 500 ng of purified total RNA according to the manufacturer’s protocol, e.g., Illumina® Stranded Total RNA Prep, Ligation with Ribo-Zero Plus kit.
The finished dsDNA libraries should be quantified by Qubit fluorometer, TapeStation 4200, and RT-qPCR using the Kapa Biosystems library quantification kit for quality control.
Indexed libraries are then pooled in equimolar ratios and pair-end sequenced on an Illumina NovaSeq 6000 sequencer with the NovaSeq 6000 Reagent Kit v1.5, with 100 cycles at 0.7 nM loading concentration.
Data analysis
Computational analysis
Build a reference genome sequence that contains the sequences of the spike-in probes as additional “chromosomes.” Acquire a set of chromosomal FASTA files (e.g., from the UCSC Genome Browser) for the relevant species (here, human) and the FASTA of ERCC spike-in sequences, e.g., https://tsapps.nist.gov/srmext/certificates/documents/SRM2374_Sequence_v1.FASTA. Process the chromosomal and spike-in FASTA files to create a reference sequence for alignment using your chosen tool, including Burrows-Wheeler-based tools. Because our preferred alignment strategy uses hisat2, we used hisat2-build with default parameters.
E.g., hisat2-build chr1.fa,chr2.fa,chr3.fa,chr4.fa,chr5.fa,chr6.fa,chr7.fa,chr8.fa,chr9.fa, chr10.fa,chr11.fa,chr12.fa,chr13.fa,chr14.fa,chr15.fa,chr16.fa,chr17.fa,chr18.fa,chr19.fa,chr20.fa,chr21.fa,chr22.fa,chrM.fa,chrX.fa,chrY.fa,ERCC92.fa Genome_With_ERCC.
Align FASTQ reads from each separate experiment to the custom reference genome. Our preferred aligner is hisat2 with default parameters. Using SAMtools, convert the aligned reads file to a sorted, indexed BAM file.
E.g., hisat2 -S aligned_reads.sam -x Genome_With_ERCC_wERCC raw_reads.fastq
samtools view -b aligned_reads.sam > aligned_reads.bam
samtools sort -n -o aligned_reads.sorted.bam aligned_reads.bam
Build or acquire a positional gene reference GTF that includes the positions of genes whose expression will be quantified and the ERCC spike-in probes in the custom reference genomes. Begin by acquiring one of many basic reference GTFs of known positions of genes in the reference genome build you initially chose; we used RefSeq. To this GTF, add GTF-formatted positions of ERCC probes. Ensure the attributes field is formatted identically between the gene and probe positions. ERCC probe positions are best left as individual chromosomes, and these chromosomes are referred to in the GTF. For example:
ERCC-00171 ERCC transcript 1 505 0.000000 + . gene_id "ERCC-00171"; transcript_id "DQ854994";; gene_name "ERCC-00171";
ERCC-00171 ERCC exon 1 505 0.000000 + . gene_id "ERCC-00171"; transcript_id "DQ854994";; gene_name "ERCC-00171";
Quantify read coverage of genes and ERCC probes for each sample separately. We used htseq-count to quantify the coverage of all genes using -i gene_id and -m intersection-strict, the sorted BAM file of aligned read positions, and the GTF of gene and probe positions in the reference. This will generate a per-sample file of per-gene read counts.
E.g., htseq-count -i gene_id --stranded=reverse -f bam -m intersection-strict aligned_reads.sorted.bam genes.gtf > aligned_reads.genecounts.txt
(Optional) Normalize read counts using one of many standard strategies that account for per-sample sequencing depth and/or gene length and/or fractions of informative reads, including the transcripts-per-million (TPM) approach. First, generate a file of per-gene total exon sizes by collapsing all exons of each isoform of each gene into a single set of regions using bedtools merge, then quantify the numbers of unique base pairs in these collapsed exons. Use the standard TPM-normalization strategy:
normterm = sum of (readcount * readlength/exonlength) across all genes.
TPM = readcount * readlength/exonlength. * 1e6/normterm [28].
Normalize counts or TPM-normalized expression using the identically calculated ERCC spike-in values. For each gene for each sample, we set the minimum expression value to 0.01 and add a pseudocount of 0.1. Create a table where each row is a gene, each column is a sample, and each cell is count or TPM-normalized count value. Using the affy R package, perform normalize.loess using the ERCC spike-in probe rows of the expression table as the subset.
(Optional) Confirm that the distribution of ERCC probe expression values approximately span the distribution of gene expression values before and after normalization.
Validation of protocol
This protocol has been used to perform spiked-in RNA-seq for several gene expression analyses, including recent publications [22,29]. Durbin et al. faced a similar challenge as we did in our studies [22] when EP300 degradation in neuroblastoma cells led to significantly decreased expression of MYCN. Durbin et al. performed RNA-seq on the MYCN-amplified neuroblastoma cell line KELLY with and without the EP300 degrader JQAD1 and the P300 inhibitor A485 using our External RNA Controls Consortium (ERCC)-controlled spike-in RNA-seq protocol (Durbin et al. [29], Figure 4B and C). In addition, Durbin et al. also used this spiked-in RNA-controlled approach for RNA-seq analysis of KELLY xenografts extracted from nude mice after the treatment with either vehicle control or the EP300 degrader JQAD1 (Durbin et al. [29], Figure 6E).
General notes and troubleshooting
General notes
The overall benefit of using a protocol for RNA sequencing with spiked-in RNA standards is the identification of global, largely unidirectional changes in mRNA levels that might not be detectable through non-normalized mRNA sequencing. In the absence of exogenous normalization, transcript level quantification is represented as a fraction of the total sample (often transcripts per million reads, or TPM), making cross-sample comparison challenging and often impossible if the total RNA per cell is altered by treatment or genetic perturbations. For example, if treatment with a chemical compound such as ATRA decreases the expression of all genes by, e.g., 50% (total mRNA output), but the relative expression of individual genes is not altered, no changes will be detected by traditional mRNA-seq. However, normalizing each sample to the exogenous spike-in control (added relative to cell number) will reveal 50% downregulation of all genes.
We recommend confirming that retinoic acid treatment decreases MYCN/MYC expression levels in the cell line/tissue of interest prior to proceeding with section B of the protocol. Assessment of the MYCN/MYC expression levels could be done via quantitative real-time PCR (qRT-PCR) using the extracted RNA from the fourth sample from each treatment condition and control sample.
The fourth sample from each treatment condition and control sample could also be used for other qRT-PCR experiments analyzing the expression levels of other genes of interest to confirm the results of the RNA sequencing of the triplicate samples.
Troubleshooting
If the total RNA yield of the control sample is very high and would require a large amount of ERCC, consider reducing the number of cells at the beginning of the experiment to decrease the absolute RNA yield in the control sample and seed fewer cells for each treatment condition.
To confirm decreased expression of MYCN/MYC in the retinoic acid–treated cells vs. control cells, consider using the extra fourth sample for a qRT-PCR experiment. We suggest the following primers to detect MYCN expression, as previously published [22]: MYCN-For 5’-CACAGTGACCACGTCGATTT-3’, MYCN-Rev 5’-CACAAGGCCCTCAGTACCTC-3’.
If MYCN/MYC expression levels are not depressed after retinoic acid treatment at day 6 as outlined in this protocol, consider performing a time series experiment, including later time points, e.g., day 8, 10, and day 14.
For low-yield ERCC detection after sequencing across samples or inconsistency between triplicates, consider repeating the experiment with freshly prepared ERCC spike-in dilution mix. ERCC dilutions should always be prepared freshly before each use.
Acknowledgments
Funding: This work was supported by grants from the National Cancer Institute, National Institute of Health, R35 CA210064 (A.T.L), and T32 HL007574-39 (N.W.L). This work was also supported by the Transcription Collaborative of St. Jude Children’s Research Hospital (A.T.L. and B.J.A.). N.W.L. is a Damon Runyon–Physician Scientist Fellow supported by the Damon Runyon Cancer Research Foundation and a recipient of funding from the Rally Foundation for Childhood Cancer Research and the Hyundai Hope on Wheels Young Investigator Award. B.J.A. is supported by the American Lebanese Syrian Associated Charities.
The original research paper in which this protocol was described and validated can be found here: Zimmerman et al. [22] Science Advances (2021), DOI: 10.1126/sciadv.abe0834.
Competing interests
A.T.L. is a founder and shareholder of Light Horse Therapeutics, which is discovering and developing small molecules to disrupt oncogenic protein complexes. M.W.Z. is an employee and shareholder of Foghorn Therapeutics. N.W.L., A.B., and B.J.A. declare no other potential conflicts of interest.
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Article Information
Publication history
Received: Mar 11, 2024
Accepted: Sep 8, 2024
Available online: Oct 13, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Assessment and Quantification of Foam Cells and Lipid Droplet–Accumulating Microglia in Mouse Brain Tissue Using BODIPY Staining
BM Boaz K. Maiyo
SL Sanna H. Loppi
HM Helena W. Morrison
KD Kristian P. Doyle
Published: Vol 14, Iss 21, Nov 5, 2024
DOI: 10.21769/BioProtoc.5107 Views: 313
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Original Research Article:
The authors used this protocol in International Journal of Molecular Sciences Nov 2023
Abstract
This paper presents a refined, user-friendly protocol for using boron-dipyrromethene (BODIPY) to assess and quantify foam cells and lipid droplet–accumulating microglia (LDAM) in mouse brain tissue. The protocol aims to enhance existing methodologies by offering precise and efficient evaluation of foam cells and LDAM burden in various neuropathological conditions linked to lipid metabolism and neuroinflammation. A notable challenge in analyzing tissue from mouse models of these neurodegenerative disorders is the interference caused by the autofluorescent molecule lipofuscin. Our protocol addresses this issue with specific steps that effectively distinguish BODIPY fluorescence from lipofuscin autofluorescence, using advanced imaging techniques and filter settings to ensure accurate and reliable analysis. By providing a straightforward and accessible method, this research aims to facilitate the broader adoption of BODIPY-based techniques for detailed foam cell and LDAM analysis in mouse brain tissue, potentially enhancing diagnostic capabilities and deepening our understanding of how these cells contribute to neurodegenerative disease mechanisms.
Key features
• To induce foam cell/LDAM CNS formation, this protocol was developed using brain tissue from mice subjected to permanent occlusion of the middle cerebral artery.
• The protocol utilizes mouse brain tissue that is fixed in 4% PFA.
• Additional markers, CD68 and Iba1, are incorporated to evaluate myeloid cell lineage.
• The protocol includes a simple method for distinguishing BODIPY fluorescence from autofluorescence.
Keywords: Foam cells Microglia Macrophage Myelin Boron-dipyrromethene BODIPY CNS
Background
Foam cell accumulation, resulting from the overwhelmed processing of myelin-derived lipids by macrophages and microglia, plays a pivotal role in multiple pathological conditions affecting the central nervous system (CNS), including stroke, spinal cord injury, and multiple sclerosis (MS) [1–3]. In the aging brain, lipid droplet–accumulating microglia (LDAM) signify a dysfunctional and pro-inflammatory state [4]. However, myeloid cells that accumulate lipid droplets often exhibit autofluorescence due to the presence of lipofuscin, complicating accurate visualization and quantification. Currently, there is a lack of user-friendly assessment protocols for accurately measuring the burden of foam cells and LDAM in mouse models of neurological disorders. The goal of this paper is to bridge this gap by introducing a comprehensive and accessible boron-dipyrromethene (BODIPY)-based protocol tailored for the analysis of foam cells and LDAM in highly autofluorescent mouse brain tissue.
The BODIPY-based approach outlined in this study leverages the distinctive properties of BODIPY dyes, especially BODIPY 493/503, for specific neutral lipid staining and efficient visualization of foam cells. Particularly well-suited for fluorescence microscopy, this technique offers a level of detail unmatched by alternative methods like Oil Red O or H&E staining. The BODIPY-based staining method we describe is rapid, straightforward, and cost-effective. Additionally, BODIPY staining can be combined with immunostaining for a more comprehensive analysis of myeloid cell phenotype. However, despite the relatively narrow emission spectrum of BODIPY 493/503, peaking at around 503 nm, careful deployment is essential to mitigate interference from the autofluorescent molecule lipofuscin, which has a heterogeneous emission spectrum ranging from 480 to 695 nm [5]. This creates the potential for spectral overlap, even though the emission spectra do not perfectly align.
By presenting a protocol that overcomes the limitations posed by lipofuscin autofluorescence, this study aims to empower researchers with a robust and user-friendly fluorescent tool for studying foam cell and lipid droplet load in mouse brain tissue. The accessibility of this BODIPY-based protocol may encourage broader adoption and implementation, thereby promoting advancements in our understanding of the relationship between foam cells and LDAM accumulation in the aging CNS and various neurological disorders.
Materials and reagents
Biological materials
Tissue sections: 40 µm thick sections from saline perfused C57BL/6 mice (Jackson Laboratories, catalog number: 00064), fixed in 4% paraformaldehyde for 24 h and cryopreserved in 30% sucrose using standard techniques. These mice were sacrificed seven weeks post a distal middle cerebral artery occlusion model of stroke, as described by Doyle et al. [6]
Reagents
Rat anti-mouse CD68 (Bio-Rad, catalog number: MCA1957GA)
Rabbit anti-mouse Iba1 (FUJIFILM Wako Pure Chemical Corporation, catalog number: 019-19741)
Goat anti-rabbit IgG (H+L) cross-adsorbed secondary antibody, Alexa FluorTM 568 (Thermo Fisher Scientific, catalog number: A-11011)
Goat anti-rat IgG (H+L) cross-adsorbed secondary antibody, Alexa FluorTM 647 (Thermo Fisher Scientific, catalog number: A-21247)
BODIPY 493/503 (4,4-Difluoro-1,3,5,7,8-Pentamethyl-4-Bora-3a,4a-Diaza-s-Indacene) (Thermo Fisher Scientific, catalog number: D3922)
Dimethyl sulfoxide (DMSO) (Thermo Fisher Scientific, Gibco®, catalog number:25200-056)
Antifade mounting medium (Vector Laboratories, SKU: H-1400-10)
10× PBS buffer pH 7.4 (Thermo Fisher Scientific, catalog number: AM9625)
Goat serum (Atlanta Biologicals, catalog number: S131x)
Triton X-100 (MilliporeSigma, catalog number: 9002-93-1)
Solutions
0.1 M PBS (see Recipes)
0.3% Triton X-100 in 0.1 M PBS (see Recipes)
Recipes
0.1 M PBS
Reagent Final concentration Amount
10× PBS 10% 100 mL
Double-distilled H2O 90% 900 mL
Total 10% 1,000 mL
0.3% Triton X-100 in 0.1 M PBS
Reagent Final concentration Amount
0.1 M PBS 99.7% 498.5 mL
Triton X-100 0.3% 1.5 mL
Total 0.3% 1,000 mL
Laboratory supplies
12-well tissue culture plate (CELLTREAT Scientific Products, catalog number: 229112)
Brushes for tissue handling (Precisionary, SKU: VF-VM-PB-CANAL)
Hydrophobic barrier pap pen (Thermo Fisher Scientific, catalog number: R3777)
Round cover glass, #1.5 thickness, 12 mm, 100 pack (Fisher Scientific, Thomas Scientific, catalog number: NC1129240)
Superfrost PlusTM microscope slides white tab (Fisher Scientific, Fisherbrand, catalog number: 1255015)
Micro-centrifuge transfer pipette (RPI Research Products International, SKU: 147500)
15 mL conical centrifuge tubes (Fisher Scientific, FalconTM, catalog number: 14-959-53A)
10 μL micropipette tips (USA Scientific, catalog number: 1161-3700)
200 μL micropipette tips (USA Scientific, catalog number: 1163-1700)
1,250 μL micropipette tips (USA Scientific, catalog number: 1161-1820)
Equipment
Digital orbital shaker (LabniqueTM, catalog number: MT-201-BX)
Leica DM6000B microscope (Leica Microsystems, SKU: 14628696)
Zeiss LSM880 NLO upright multiphoton/confocal microscope
Software and datasets
ImageJ (free, https://imagej.net/software/fiji/downloads, Fiji version, 2024)
Procedure
Select infarcted mouse brain tissue sections.
Note: If only staining for BODIPY, skip to step 8.
Place tissue in a 12-well tissue culture plate and wash three times for 10 min each in 0.1 M PBS at room temperature (RT) on a shaker.
Blocking: Block for 1 h at RT with 10% goat serum and 0.3% Triton X-100 in 0.1 M PBS.
Tip: Make sure the blocking solution uses serum from the species in which the secondary antibody was generated.
Primary antibody: Incubate overnight at room temperature on a rotator in 3% goat serum and 0.3% Triton X-100 in 0.1 M PBS containing: 1:500 anti-CD68 made in rat and 1:500 anti-Iba1 made in rabbit.
Tip: If the staining is faint, increase the antibody concentration. If there is excessive background, reduce the antibody concentration. The recommended dilution is 1:500–1,000. Another alternative when staining is too faint is to increase the incubation time to 48–72 h.
Wash three times for 10 min each in 0.1 M PBS at RT on a shaker.
Secondary antibody: Incubate for 2 h at RT in 3% goat serum + 0.3% Triton X-100 in 0.1 M PBS with 1:500 goat anti-rabbit and 1:500 goat anti-rat.
Tip: If the staining is faint, increase the antibody concentration. If there is excessive background, reduce the antibody concentration. The recommended dilution is 1:500–1,000. Another alternative is to increase the incubation time to 4 h.
Note: From this point onward, keep samples protected from light.
Wash three times for 10 min each in 0.1 M PBS at RT on a shaker.
Mounting: Gently place tissue sections in 0.1 M PBS and mount them on slides.
Notes:
Air dry slides for approximately 30 min.
To avoid drying the tissue, do not leave the slides out for more than 1 h.
Draw a hydrophobic barrier around the sections with the hydrophobic pen.
First washing: Wash the sections three times for 10 min each in 0.1 M PBS at RT.
Staining: Incubate the sections in a solution of 1:25 BODIPY 492/515 in DMSO for 30 min, protected from light.
Second washing: Wash the sections three times for 10 min each in 0.1 M PBS at RT, protected from light.
Cover slipping:
Mount the sections with antifade mounting media.
Incubate for 1–2 h at RT, protected from light.
Imaging: Detect the fluorescence emission of BODIPY 493/503-stained samples using a fluorescence microscope equipped with a 493 nm excitation filter and a 503 nm emission filter.
Data analysis
This section outlines the analysis of cellular lipids and lipid droplets stained with BODIPY 493/503 within a specified area using the ImageJ software, a method previously applied by Loppi et al. [7].
Open the program.
Click File, then Open, and select the desired image.
The image appears on the screen.
To select the desired area, click on the Polygon selections on the toolbar and manually outline the area.
• It should appear as a polygon icon with defined sides.
Note: If there is no need to select a smaller area of the image, this step can be omitted.
Select Image, Type, and 8-bit.
• The image should now become black and white.
Continue by clicking Image, Adjust, and Threshold.
A screen will appear that will allow you to select positive staining.
• Adjust the Min and Max threshold values to a level where only the stained areas are colored.
Tip: Set the Max to 255 and slowly lower the min for finer selections.
To quantify the selection, click Analyze and Measure.
• A screen will appear with the name of the image, as well as the Area, Mean, Min, Max, %Area, MinThr, and MaxThr.
Tip: The desired measurements can be selected by clicking Analyze and Set Measurements.
Copy the data to an Excel spreadsheet.
Close the image.
Repeat steps 2–10 until all images are analyzed.
Tip: For accurate comparison between images, keep the Min and Max threshold values the same between images.
Use the %Area value for comparisons.
Calculate the average %Area for each group of brain sections (e.g., control vs. treated).
Perform statistical analysis to determine whether there is a significant difference in lipid accumulation between groups.
Validation of protocol
Autofluorescence control
Background: Autofluorescence, predominantly from lipofuscin, presents a significant challenge for researchers studying injured and diseased brain tissue, as it occurs across a spectrum from 480 to 695 nm [5]. It can be observed by imaging an unstained section through multiple filter cubes with different excitation and emission settings. Lipofuscin autofluorescence is most intense at lower wavelengths and gradually diminishes as wavelengths increase, necessitating longer exposure times for accurate capture (Figure 1A–D). Considering that the BODIPY 493/503 stain emits at 503 nm, it is crucial to ensure that the stained areas accurately represent lipids in the brain rather than artifacts from lipofuscin autofluorescence.
Figure 1. The broad spectrum of lipofuscin autofluorescence. A. Lipofuscin-rich infarcted region of a brain section from a stroked mouse, imaged with an exposure time of 1.2 s, excitation of 355–425 nm, and emission of 470 nm. B. The same section imaged with an exposure time of 1.2 s, excitation of 460–500 nm, and emission of 512–542 nm. C. The same section imaged with an increased exposure time of 3 s, excitation of 540–580 nm, and emission of 590–650 nm. D. The same section imaged with an increased exposure time of 10 s, excitation of 590–650 nm, and emission of 662–738 nm. All images were captured at 10× magnification with the scale bar representing 300 µm.
Experimental setup: To address the issue of autofluorescence, we conducted a control experiment using two brain tissue samples from the same experimental group. One sample was stained with BODIPY according to the described protocol while the other was only incubated in PBS for the same duration instead of the BODIPY solution. Both samples were imaged using a Leica microscope the following day.
Imaging protocol: The BODIPY-stained section was initially imaged at 10× magnification with an exposure time of 750 ms and a gain of 1.5, which provided clear visualization of the stain (Figure 2A). We then increased the magnification to 20×, achieving an even clearer image with an exposure time of 576.07 ms and the same gain (Figure 2B). In contrast, the control section, imaged under identical conditions at both 10× and 20× magnifications, showed no discernible signal (Figure 2C and D). Only when the exposure time was extended to 7 s at 10× and 5 s at 20× did lipofuscin autofluorescence become visible (Figure 2E and F), thereby confirming the specificity of the BODIPY staining over lipofuscin autofluorescence.
Figure 2. Autofluorescence control. A. Whole-brain section stained with BODIPY, imaged at 10× magnification with an exposure time of 750 ms and a gain of 1.5. B. Enlarged view of the infarct from image A, using the same imaging parameters. C. Section processed with the BODIPY protocol but not stained with BODIPY, imaged at 10× magnification with an exposure time of 750 ms and a gain of 1.5. D. Enlarged view of the infarct area from image C. E. The same unstained section as in C, imaged at 10× magnification with an exposure time of 7 s and a gain of 1.5 to reveal autofluorescence. F. Enlarged view of the infarct area from image E. Scale bars: 1 mm for images A, C, and E; 500 μm for images B, D, and F.
Lipid droplet visualization within myeloid cells
When microglia and macrophages are overwhelmed by myelin debris following a stroke, they transform into foam cells. This transformation not only results in a loss of immune function but also activates pathways that may contribute to tissue damage [2]. To validate the use of the BODIPY protocol for visualizing lipid droplets within myeloid cells, stroked brain tissue was immunostained with antibodies specific for IBA1 and CD68 by using standard techniques described by Potts et al. [8]. IBA is a pan surface marker for microglia, while CD68 serves as an intracellular marker predominantly found in the lysosomes of monocytes and macrophages, commonly used to identify activated, phagocytic microglia and macrophages [9,10]. The tissue was subsequently stained with BODIPY and imaged.
Imaging of the infarct revealed successful detection and visualization of BODIPY within IBA1 and CD68 positive cells. The IBA1 stain sharply outlined the myeloid cells, as depicted in Figure 3. BODIPY staining indicated a significant presence of lipid droplets within these cells. CD68 was also observed colocalized with the BODIPY stain within the myeloid cells, confirming the presence of BODIPY within the endosomal-lysosomal system of these cells. These findings validate the effectiveness of BODIPY staining for visualizing lipid-laden myeloid cells, including foam cells and LDAMs, in the context of neurological injury.
Figure 3. Intracellular staining of lipids in myeloid cells. The section depicted was stained with IBA1 (cyan), BODIPY (green), and CD68 (red), and captured using a Zeiss LSM880 NLO upright multiphoton/confocal microscope at 40× magnification with a 2× zoom. The top row images feature a scale bar of 20 μm, and the bottom row images feature a scale bar of 10 μm.
General notes and troubleshooting
Handling fluorescent stains and imaging guidelines
Fluorescent stains gradually lose intensity over time; therefore, it is essential to image all sections within a set on the same day. Be aware that exposure times may vary slightly from day to day. To prevent photobleaching, avoid leaving sections under the microscope for extended periods, as prolonged exposure can cause tissue bleaching, evident as dark spots in the sections. Optimize your imaging process by starting with a lower exposure time to locate the desired area, then increase the exposure time only when ready to capture the image. If possible, perform imaging in a dark room to enhance the quality of your results. Always store slides at room temperature in a dark place to minimize light exposure.
Co-staining protocol
When performing co-staining with other markers using immunohistochemistry (IHC), it is advisable to complete the IHC first, followed by BODIPY staining. For colocalization, higher magnifications provide more precise images. Using 10× or 20× magnification may not yield satisfactory results. Instead, opt for 40× magnification and consider utilizing a Z-stack to achieve optimal colocalization accuracy.
Troubleshooting
Issue: Tissue clumping with the combined IHC and BODIPY staining protocol.
Solution: Pre-mounting instead of free floating.
Complete all IHC steps on pre-mounted slides and, following the washing steps, continue on to BODIPY staining.
Avoid using free-floating methods for IHC followed by BODIPY to avoid tissue clumping.
Rinse all tissue well to remove excess BODIPY staining.
Issue: The scale is unrecognized in the images.
Solution: Manually set the scale in ImageJ by using scale bar images for the magnification used. Follow these steps:
Open the scale bar images in ImageJ.
Use the straight line tool from the toolbar to draw a line over the scale bar.
Go to Analyze and select Set Scale.
• A dialog box will appear.
In the dialog box, change the Known distance to the actual distance of the scale bar.
Set the Unit of length to µm and check the Global box.
• Click OK.
• The dialog box will automatically close.
Keep the scale bar image open in the background for the duration of the analysis. It must remain open as long as ImageJ is running.
Note: Once set, the scale will apply to all images as long as ImageJ remains open, eliminating the need to reset the scale for each new image.
Acknowledgments
We would like to acknowledge MMPC Immunohistochemistry for their foundational BODIPY protocol, upon which this study's protocol was developed [11]. This research was supported by the National Institute of Neurological Disorders and Stroke RF1NS131110, and National Institute on Aging R01AG063808.
Competing interests
The authors declare no conflicting interests.
Ethical considerations
The animal study protocol was approved by the Institutional Animal Care & Use Committee (IACUC) of the University of Arizona.
References
Zierfuss, B., Weinhofer, I., Buda, A., Popitsch, N., Hess, L., Moos, V., Hametner, S., Kemp, S., Köhler, W., Forss‐Petter, S., et al. (2020). Targeting foam cell formation in inflammatory brain diseases by the histone modifier MS‐275. Ann Clin Transl Neurol. 7(11): 2161–2177.
Zbesko, J. C., Stokes, J., Becktel, D. A. and Doyle, K. P. (2023). Targeting foam cell formation to improve recovery from ischemic stroke. Neurobiol Dis. 181: 106130.
Wang, X. X., Li, Z. H., Du, H. Y., Liu, W. B., Zhang, C. J., Xu, X., Ke, H., Peng, R., Yang, D. G., Li, J. J., et al. (2024). The role of foam cells in spinal cord injury: challenges and opportunities for intervention. Front Immunol. 15: e1368203.
Marschallinger, J., Iram, T., Zardeneta, M., Lee, S. E., Lehallier, B., Haney, M. S., Pluvinage, J. V., Mathur, V., Hahn, O., Morgens, D. W., et al. (2020). Lipid-droplet-accumulating microglia represent a dysfunctional and proinflammatory state in the aging brain. Nat Neurosci. 23(2): 194–208.
Mochizuki, Y., Park, M. K., Mori, T. and Kawashima, S. (1995). The Difference in Autofluorescence Features of Lipofuscin between Brain and Adrenal. Zool Sci. 12(3): 283–288.
Doyle, K. P., Fathali, N., Siddiqui, M. R. and Buckwalter, M. S. (2012). Distal hypoxic stroke: A new mouse model of stroke with high throughput, low variability and a quantifiable functional deficit. J Neurosci Methods. 207(1): 31–40.
Loppi, S. H., Tavera-Garcia, M. A., Scholpa, N. E., Maiyo, B. K., Becktel, D. A., Morrison, H. W., Schnellmann, R. G. and Doyle, K. P. (2023). Boosting Mitochondrial Biogenesis Diminishes Foam Cell Formation in the Post-Stroke Brain. Int J Mol Sci. 24(23): 16632.
Potts, E. M., Coppotelli, G. and Ross, J. M. (2020). Histological-Based Stainings using Free-Floating Tissue Sections. J Visualized Exp. (162): e61622.
Perego, C., Fumagalli, S. and De Simoni, M. G. (2011). Temporal pattern of expression and colocalization of microglia/macrophage phenotype markers following brain ischemic injury in mice. J Neuroinflammation. 8(1): 174.
Yu, X., Guo, C., Fisher, P. B., Subjeck, J. R., & Wang, X.-Y. (2015). Chapter Nine - Scavenger Receptors: Emerging Roles in Cancer Biology and Immunology. Adv Cancer Res. 128: 309–364.
Rutkowsky, J. (2019). UC Davis - Immunohistochemistry BODIPY v2. protocols.io: e56rg9d6.
Article Information
Publication history
Received: Jun 28, 2024
Accepted: Sep 9, 2024
Available online: Oct 15, 2024
Published: Nov 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Neuroscience > Cellular mechanisms > Microglia
Cell Biology > Cell imaging > Fluorescence
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5,108 | https://bio-protocol.org/en/bpdetail?id=5108&type=0 | # Bio-Protocol Content
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Multiplex Genome Editing of Human Pluripotent Stem Cells Using Cpf1
HM Haiting Ma
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5108 Views: 462
Reviewed by: Thirupugal GovindarajanVishal Nehru Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in cell reports medicine Sep 2020
Abstract
Targeted genome editing of human pluripotent stem cells (hPSCs) is critical for basic and translational research and can be achieved with site-specific endonucleases. Cpf1 (CRISPR from Prevotella and Francisella) is a programmable DNA endonuclease with AT-rich PAM sequences. In this protocol, we describe procedures for using a single vector system to deliver Cpf1 and CRISPR RNA (crRNA) for genome editing in hPSCs. This protocol enables indel formation and homologous recombination–mediated precise editing at multiple loci. With the delivery of Cpf1 and a single U6 promoter-driven guide RNA array composed of an AAVS1-targeting and a MAFB-targeting crRNA array, efficient multiplex genome editing at the AAVS1 (knockin) and MAFB (knockout) loci in hPSCs could be achieved in a single experiment. The edited hPSCs expressed pluripotency markers and could differentiate into neurons in vitro. This system also generated INS reporter hPSCs with a 6 kb cassette knockin at the INS locus. The INS reporter cells can differentiate into β-cells that express tdTomato and luciferase, permitting fluorescence-activated cell sorting of hPSC-β-cells. By targeted screening of potential off-target sequences that are most homologous to crRNA sequences, no off-target mutations were detected in any of the tested sequences. This work provides an efficient and flexible system for precise genome editing in mammalian cells including hPSCs with the benefits of less off-target effects.
Key features
• A single-vector system to deliver Cpf1 and crRNA enables the sorting of transfected cells
• Efficient and simultaneous multi-modular genome editing exemplified by mutation of MAFB and knockin of AAVS1 loci in a single experiment
• Edited PSCs showed minimal off-target effects and can be differentiated into multiple cell types
Keywords: Human pluripotent stem cells (hPSCs) Genome editing CRISPR Cpf1 Differentiation of hPSCs Regenerative medicine
Graphical overview
Genome editing of human pluripotent stem cells (hPSCs) using Cpf1. The top panel provides a brief overview of the approach; the bottom panel indicates additional details corresponding to the procedures of the protocol. The bottom panel was generated with biorender.com.
Background
Human pluripotent stem cells (hPSC) [1,2] and their differentiated cell types, such as pancreatic β-cells [3], enable cell replacement therapies as a potential treatment for multiple diseases [4]. Genetic modifications of hPSCs can correct genetic lesions and provide functions including increased immune tolerance by overexpressing PDL1 [5]. Multiple programmable nucleases have been developed for efficient genome editing in hPSCs: zinc finger nucleases [6], TALE nucleases [7], and CRISPR-Cas9 [8,9]. Complementary utilization of these nucleases is a critical component for hPSC-based regenerative medicine [10,11].
The CRISPR-Cas9 system enables efficient genome editing applicable to diverse cell types [12]. However, wild-type SpCas9 can generate double-strand DNA breaks in off-target sites partially complementary to guide RNA. The off-target effects of SpCas9 can be reduced by modifying the basic amino acid residues of the DNA binding domain of SpCas9, but the modified SpCas9 could also show reduced activity compared to wild-type SpCas9 [13]. Base-editing [14] and prime-editing [15] technologies enable genome editing without generating double-stranded DNA breaks. However, introducing large DNA fragments to specific loci of cells remains challenging as recent studies showed insertions in the range of hundreds of nucleotides [16]. Furthermore, to achieve effective multiplex editing at the single-cell level with a DNA vector, each guide RNA requires its promoter, complicating multiplex applications [17].
CRISPR-Cpf1 family endonucleases confer efficient genome editing in transformed mammalian cell lines [18] and mouse embryos [19,20]. Additionally, Francisella novicida Cpf1 (FnCpf1) and Acidaminococcus Cpf1 (AsCpf1) have RNase activity to process their cognate CRISPR RNA (crRNA) [18,21], making it possible to deliver multiple crRNAs from a pre-crRNA array driven by a single U6 promoter [22]. Furthermore, Cpf1 has less off-target effects in genome editing experiments with transformed human cell lines [23,24]. These unique features make Cpf1 a system that complements Cas9.
This protocol describes a detailed genome-editing procedure that uses a single vector system that expresses AsCpf1 and its crRNA. With donor constructs, locus-specific knockin alleles of hPSC were generated and could be differentiated into neurons and β-cells. Furthermore, by introducing a pre-crRNA array composed of AAVS1-targeting guide and MAFB-targeting guide, we observed 100% editing of MAFB locus in AAVS1 targeted clones, providing an example for genome editing in multiple loci in hPSCs. This protocol provides a flexible system for genome editing in hPSCs.
Materials and reagents
Biological materials
H1-OCT4-GFP pluripotent stem cells (WiCell Research Institute, catalog number: H1 OCT4-EGFP, NIH Human Embryonic Stem Cell Registry ID: NIHhESC-10-0043)
WIBR3 pluripotent stem cells (Whitehead Institute, NIH Human Embryonic Stem Cell Registry ID: NIHhESC-10-0079)
HEK293T cells (ATCC, catalog number: CRL-3216)
Stbl3 Chemically Competent E. coli (Invitrogen, catalog number: C737303)
DR4 mice [Dnmt1tm3Jae Hprt1b-m3 Tg(pPWL512hyg)1Ems/J] (Jackson Laboratory, catalog number: 003208)
NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice (Jackson Laboratory, catalog number: 005557)
Reagents
DMEM-F12 medium (Life Technologies, catalog number: 11330-057)
KSR (Life Technologies, catalog number: 10828-028)
FGF2 (Life Technologies, catalog number: PHG0261)
2-Mercaptoethanol (2-ME) (Life Technologies, catalog number: 21985-023)
100× L-glutamine (Life Technologies, catalog number: 25030-081)
100× GlutaMAX (Life Technologies, catalog number: 35050-061)
100× MEM-NEAA (Life Technologies, catalog number: 11140-050)
100× penicillin/streptomycin (Life Technologies, catalog number: 15140-122)
100× Insulin-transferrin-selenium-ethanolamine (Life Technologies, catalog number: 51500-056)
Collagenase IV (Life Technologies, catalog number: 17104019)
mTeSR1 medium (STEMCELL Technologies, catalog number: mTeSR1)
Accutase (STEMCELL Technologies, catalog number: 7920)
MCDB131 medium (Life Technologies, catalog number: 10372019)
Y-27632 (Thermo Fisher Scientific, catalog number: 688000-100MG)
Puromycin (Sigma-Aldrich, catalog number: P7255-100MG)
CHIR99021 (Cayman Chemical, catalog number: 252917-06-9)
Matrigel (Thermo Fisher Scientific, catalog number: CB40234)
Activin A (R&D Systems, catalog number: 338-AC-050)
KGF (PeproTech, catalog number: 100-19-250UG)
Vitamin C (Sigma-Aldrich, catalog number: A4544-25G)
Retinoic acid (Sigma-Aldrich, catalog number: R2625-100MG)
SANT-1 (Sigma-Aldrich, catalog number: S4572-5MG)
LDN193189 (Sigma-Aldrich, catalog number: SML0559-5MG)
Heparin (Sigma-Aldrich, catalog number: H3149)
TPB (Tocris, catalog number: 5343)
EGF (PeproTech, catalog number: AF-100-15)
T3 (EMD, catalog number: 64245)
ALK5 inhibitor II (Axxora, catalog number: ALX-270-445-M005)
Gamma secretase inhibitor XX (VWR, catalog number: 82602-300)
Trace Elements A (Corning, catalog number: 25-021-CI)
Trace Elements B (Corning, catalog number: 25-022-CI)
Fatty acid–free BSA (Thermo Fisher Scientific, catalog number: 50412870)
D-Luciferin potassium salt (Perkin Elmer, catalog number: 122799)
RPMI 1640 (Gibco, catalog number: 31800-089)
DMEM high glucose medium (Cytiva, catalog number: SH30022)
FBS (Cytiva, catalog number: SH30396.03HI)
BbsI-HF (New England Biolabs, catalog number: R3539S)
SphI-HF (New England Biolabs, catalog number: R3182L)
CutSmart Buffer (New England Biolabs, catalog number: B6004)
T7 Endonuclease I (New England Biolabs, catalog number: M0302S)
Lipofectamine 2000 transfection reagent (Invitrogen, catalog number: 11668019)
Plasmid Mini Kit (Omega Bio-Tek, catalog number: D6943-02)
DNA Clean & Concentrator (Zymo Research, catalog number: D4033)
T4 PNK (New England Biolabs, catalog number: M0201S)
T4 DNA ligase (New England Biolabs, catalog number: M0202M)
DNeasy Blood & Tissue Kit (Qiagen, catalog number: 69504)
Q5 HiFi 2× Master Mix (New England Biolabs, catalog number: M0492L)
10% TBE PAGE gel (Bio-Rad, catalog number: 3450053)
Mouse anti-OCT3/4 antibody, 1:100 (BD Transduction Laboratories, catalog number: 611203)
Rabbit anti-NANOG antibody, 1:100 (Thermo Fisher Scientific, catalog number: PA1-097)
Mouse anti-TRA-1-60, 1: 100 (Life Technologies, catalog number: 411000)
Mouse anti-SSEA4 antibody, clone MC-813-70, 1:100 (Thermo Fisher Scientific, catalog number: MA1-021)
Mouse anti-Nestin antibody, 1:100 (Santa Cruz, catalog number: SC-23927)
Mouse anti-TUBB3 antibody, 1:1000 (BioLegend, catalog number: 801201)
Alexa 488 conjugated anti-mouse IgG antibodies, used at 1:400 (Life Technologies, catalog number: A21202)
Alexa 488 conjugated anti-rabbit IgG antibodies, used at 1:400 (Life Technologies, catalog number: A21206)
ZymoPURE II Plasmid Midiprep Kit (Zymo Research, catalog number: D4200)
AmpliTaq Gold 360 Master Mix (Thermo Fisher Scientific, catalog number: 4398881)
E.Z.N.A.® Gel Extraction kit (Omega Bio-Tek, catalog number: D2500-02)
HindIII-digested Lamda DNA (New England Biolabs, catalog number: N3012S)
Pyruvate (100 mM) (Corning, catalog number: 25-000-CIR)
Insulin solution (Sigma, catalog number: I9278)
Neurobasal medium (Life Technologies, catalog number: 21103049)
AlbuMAX I Lipid-Rich BSA (Life Technologies, catalog number: 11020021)
N2 NeuroPlex (GeminiBio, catalog number: 400163)
Lactic syrup (Sigma, catalog number: L4263-100ML)
Biotin (Sigma, catalog number: B4639-500MG)
Gem21 without vitamin A (GeminiBio, catalog number: 400-161-010)
Proteinase K (Promega, catalog number: C3021)
Sodium citrate tribasic dihydrate (Sigma, catalog number: C8532)
Primers
F NGS seq primer for MAFB of WIBR3 602-1 ATATGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-2 AGAGGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-3 ACACGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-4 GCGCGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-5 GTGTGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-6 CTCTGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-7 TTAAGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-8 TTGGGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-9 TTCCGGTCAAGTGCGAGAAACTCG
F NGS seq primer for MAFB of WIBR3 602-10 CCTTGGTCAAGTGCGAGAAACTCG
R NGS seq primer for MAFB of WIBR3 GCAGGGACAGGGTCCGGGGTAG
AAVS1 off 1 F CTGCTGAACACTCAGCATCTGCC
AAVS1 off 1 R CAGAGGAGCGAGTGGAGCAGACAG
AAVS1 off 2 F ATTGCAATATCCTCCTATTAGCC
AAVS1 off 2 R CACTAGAGTCACCCTATGGCTCCC
AAVS1 off 3 F CATGGGACAAGTTGATAGCTAAG
AAVS1 off 3 R GAAGTTACTCTGAAACGTATAGCAC
AAVS1 off 4 F GCTGCTCCTGGATTTAGCAAAC
AAVS1 off 4 R GCCCAGCCCAACACTTTTGGTC
AAVS1 off 5 F GTGTGCTAGTATCATTGCAAAAG
AAVS1 off 5 R CACTATTGCGTTCTCTCATTTCTC
MAFB off 1 F TCTGAGGTCCGTCTCACACACTG
MAFB off 1 R CACTGTTCAAAGAGTTTGAACATTCC
MAFB off 1 F CTCCTGACTATTGCAGTTGCTGGTCACC
MAFB off 1 R AACAGAGGAGCGAGTGGAGC
INS off 1 F CGGAGTCTCACTTTGTTGCCATG
INS off 1 R TGGTTAGAACTTCCTGCCCACAG
Gelatin (Sigma, catalog number: G2500)
Mitomycin C (Sigma, catalog number: M4287)
CMV-AsCpf1-2A-GFP-U6-AAVS1-crRNA plasmid (Addgene, #194716)
AVS1-tdT targeting plasmid (Addgene, #194728)
CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #159281)
CMV:AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #194715)
CAGGS-AsCpf1-2A-GFP-U6-tdT-crRNA plasmid (Addgene, #194724)
CAGGS-AsCpf1-2A-GFP-U6-AAVS1-crRNA plasmid (Addgene, #194723)
CAGGS-AsCpf1-2A-GFP-U6-AAVS1-MAFB-crRNA (Addgene, #194725)
CAGGS-AsCpf1-2A-GFP-U6-INS-crRNA plasmid (Addgene, #159283)
INS-2A-luciferase-2A-tdT donor plasmid (Addgene, #159348)
LB broth (Sigma, catalog number: L3522-1KG)
Agar (Fisher Scientific, catalog number: 1423-500)
Na2HPO4·7H2O (Sigma, catalog number: S9390)
Na2HPO4 (Fisher Scientific, catalog number: BP329-500)
NaCl (Sigma, catalog number: S271-3)
KCl (Fisher Scientific, catalog number: BP366-1)
KH2PO4 (Sigma, catalog number: P0662)
CaCl2 (Sigma, catalog number: C8106-500G)
MgCl2·6H2O (Sigma, catalog number: M2670-500G)
Tris (Millipore, catalog number: 648311-1KG)
EDTA (Sigma, catalog number: E6758-500g)
SDS (Fisher Scientific, catalog number: BP166-500)
H3PO4 (85%) (Sigma, catalog number: 345245-500ML)
Ascorbic acid (Sigma, catalog number: A4544-25G)
REDTaq PCR Reaction Mix (Sigma, catalog number: R2523)
Prime-It II Random Primer Labeling Kit (Agilent, catalog number: 300385)
CHROMA SPIN + TE-30 (Takarabio, catalog number: 636093)
Hybond-XL membrane (Cytiva, catalog number: RPN2020S)
BioMax MS film (Carestream, catalog number: 8294958)
NaOH (Sigma, catalog number: S5881-500G)
HCl (Sigma, catalog number: H1758)
Dorsomorphin (R & D Systems, catalog number: 3093/10)
Acetic acid (Sigma, catalog number: A6283)
Ethidium bromide (Sigma, catalog number: E1510)
[a-32P]dCTP (Revvity, catalog number: BLU513H)
Solutions
LB agar (see Recipes)
Gelatin solution (see Recipes)
10× PBS (see Recipes)
Ca2+ and Mg2+ solution (see Recipes)
PBS+ (see Recipes)
MEF medium/HEK293T cells medium (see Recipes)
MEF inactivation medium (see Recipes)
DNA-extraction buffer (see Recipes)
2× phosphate buffer (pH 7.2) (see Recipes)
Hybridization buffer (see Recipes)
20× SSC buffer (pH 7.0) (see Recipes)
hPSC medium (see Recipes)
NGD medium (see Recipes)
50× TAE buffer (see Recipes)
Recipes
LB agar
Reagent Final concentration Quantity or Volume
LB broth 2.5% w/vol 10 g
Agar 1.5% w/vol 6 g
MilliQ water n/a Add to 400 mL
Total (optional) n/a 400 mL
Gelatin solution
Reagent Final concentration Quantity or Volume
Gelatin 0.2% (w/v) 0.8 g
H2O n/a 400 mL
Total n/a 400 mL
10× PBS
Reagent Final concentration Quantity or Volume
Na2HPO4·7H2O 25.6 g
Na2HPO4 13.56 g
NaCl 80 g
KCl 2 g
KH2PO4 2 g
H2O To 1,000 mL
Total 1,000 mL
Adjust pH to 7.4 and autoclave for 15 min at 121 °C.
Ca2+ and Mg2+ solution
Reagent Final concentration Quantity or Volume
CaCl2 5 g
MgCl2·6H2O 5 g
H2O To 500 mL
Total 500 mL
Filter the solution through a 0.22 μm filter and autoclave for a 45 min liquid cycle. After autoclaving, keep the solution at 4 °C.
PBS+
Reagent Final concentration Quantity or Volume
1× PBS (diluted from 10× PBS) 990 mL
Ca2+ and Mg2+ solution (Recipe 4) 10 mL
Total 1000 mL
Stir the solution for approximately 30 min until it clears up. Then, filter through a 0.22 μm filter and keep refrigerated at 4 °C.
MEF medium/HEK293T cells medium
Reagent Final concentration Quantity or Volume
DMEM high glucose 435 mL
FBS 10% 50 mL
Penicillin/streptomycin 1% 5 mL
NEAA 1% 5 mL
Pyruvate (100 mM) 1 mM 5 mL
Total 500 mL
MEF inactivation medium
Reagent Final concentration Quantity or Volume
DMEM high glucose 87 mL
FBS 10% 10 mL
Penicillin/streptomycin 1% 1 mL
NEAA 1% 1 mL
Pyruvate (100 mM) 1 mM 1 mL
Mitomycin C 5 µg/mL 0.5 mg
Total 100 mL
DNA-extraction buffer
Reagent Final concentration Quantity or Volume
1 M Tris (pH 8.0) 50 mM 25 mL
0.5 M EDTA (pH 8.0) 10 mM 10 mL
5 M NaCl 100 mM 10 mL
10% SDS 0.5% 25 mL
MilliQ H2O 430 mL
20 mg/mL Proteinase K 0.5 mg/mL 25 μL for 1 mL, add just before use
2× phosphate buffer (pH 7.2)
Reagent Final concentration Quantity or Volume
Na2HPO4·7H2O 0.5 M 67 g
H3PO4 (85%) 4 mL
MilliQ H2O Add to 1 L
Hybridization buffer
Reagent Final concentration Quantity or Volume
2× phosphate buffer (Recipe 9) 1× 50 mL
0.5 M EDTA (pH 8.0) 1 mM 0.2 mL
5 M NaCl 100 mM 5 mL
20% SDS 7 % 35 mL
MilliQ H2O 15 mL
BSA 1% 1 g, add before use in 100 mL buffer warmed to 60 °C
20× SSC buffer (pH 7.0)
Reagent Final concentration Quantity or Volume
NaCl 175.3 g
Sodium citrate tribasic dihydrate 88.2 g
MilliQ H2O Add to 1 L, adjust pH to 7.0 with a few drops of HCl
hPSC medium
Reagent Final concentration Quantity or Volume
DMEM-F12 385 mL
KSR 5% 25 mL
FBS 15% 75 mL
Penicillin/streptomycin 1% 5 mL
NEAA 1% 5 mL
L-glutamine 1% 5 mL
FGF2 4 ng/mL
2-ME 1 μM
Total (optional) n/a 500 mL
NGD medium
Reagent Final concentration Quantity or Volume
Neurobasal medium 500 mL
Gem21 without vitamin A 1% 5 mL
AlbuMAX I 0.2% w/v 1 g
NeuroPlex N2 0.5% 2.5 mL
NaCl (5 M) 5 mL
Pyruvate (100 mM) 1 mM 5 mL
Penicillin/streptomycin (100×) 1× 5 mL
GlutaMAX (100×) 1× 5 mL
Biotin (5 mg/mL in 1 M NaOH) 0.35 μL
Ascorbic acid (100 mM) 50 μL
Lactic syrup (85%) 100 μL
50× TAE
Reagent Final concentration Quantity or Volume
Tris 2 M 121 g
EDTA (0.5 M, pH 8) 50 mM 50 mL
Acetic acid 1 M 28.55 mL
H2O To 500 mL
Total 500 mL
Laboratory supplies
6-well ultra-low attachment plates (Corning, catalog number: CLS3471-24EA)
AggreWell 400 (StemCell Technologies, catalog number: 34425)
12-well plates (Corning, catalog number: 3512)
6-well plates (Corning, catalog number: 3506)
Cell lifters (Corning, catalog number: 3008)
Electroporation cuvettes, 0.2 cm (Bio-Rad, catalog number: 1652086)
PCR tubes and caps (USA Scientific, catalog number: 1402-2500)
Equipment
SMZ1270 microscope (Nikon, model: SMZ1270)
Eclipse Ti microscope (Nikon, model: Eclipse Ti)
Thermal Cycler (Bio-Rad, model: T100)
Accuspin micro desktop centrifuge (Fisher Scientific, catalog number: 75002461)
ST1 Plus desktop centrifuge (Sorvall ST1 Plus, with =TX-400 rotor and adaptors, catalog number: 75016030)
Heracell 150i CO2 incubator (Thermo Scientific, catalog number: 13998034)
Incushaker (Benchmark Scientific, catalog number: 31-206)
Gene Pulser Xcell Eukaryotic System (Bio-Rad, catalog number: 1652661)
Nanodrop 2000 spectrometer (Thermo Scientific, catalog number: ND2000)
AlphaImager gel documentation system (Alpha Innotech, model: AlphaImager 2200)
Xenogen IVIS Spectrum imager (Xenogen, model: IVIS Spectrum)
FACSAria cell sorter (BD Biosciences)
Software and datasets
Cpf1 crRNA design was performed at https://portals.broadinstitute.org/gppx/crispick/public
Prediction of off-targets was obtained using Cas-OFFinder, http://www.rgenome.net/cas-offinder
Procedure
The following are step-by-step procedures for the process of designing crRNA, validation of crRNA, genome editing of hPSCs (knockout, knockin, or multiplex genome editing), characterizing genome-edited hPSCs, testing potential off-targeting effects, and differentiation of hPSC. Some steps can be skipped based on the experimental design.
Design DNA oligos for Cpf1 crRNAs
Design crRNA. Sequences encoding Cpf1 crRNAs are designed using 23nt crRNA from the website https://portals.broadinstitute.org/gppx/crispick/public. An example of designing Cpf1 crRNA targeting the AAVS1 locus is shown in Figure 1A with the PAM sequence in magenta. The target sequence is in yellow and matches the guide sequence in the Cpf1 crRNA.
Design the DNA oligonucleotide by adding the auxiliary bases (highlighted in cyan in Figure 1B) to the crRNA guide sequence design. The addition of the auxiliary bases generates annealed products with adhesive ends (Figure 1C) to facilitate cloning the annealed products to the Cpf1 crRNA backbone plasmids.
Figure 1. Design gene-specific crRNAs to clone to the bicistronic AsCpf1 plasmids and validation in HEK293T cells.A. Example of Cpf1 crRNA design for the genomic sequence at the AAVS1 locus. Magenta denotes the TTTN PAM sequence and yellow denotes gene-specific target sequence. B. Example of oligo design for the targeted genomic sequence at the AAVS1 locus. C. Annealed products from the oligos in B. Cyan denotes the sequence for adhesive ends to facilitate cloning and yellow denotes a target-specific sequence. D. BbsI cloning sites of Cpf1 plasmids. Cyan denotes spacer sequence, gray denotes BbsI cognitive sites, and arrows denote cleavage sites upon BbsI digestion. E. Design of a bicistronic vector delivering both guide RNA (AAVS1 sg RNA shown as nucleotides in green) and AsCpf1 followed by 2A-GFP. F. Representative FACS scatter plots of control HEK293T cells, AsCpf1-2A-GFP expressing, AsCpf1-2A-GFP-AAVS1 expressing HEK293T cells. G. T7 endonuclease I (T7EI) assay to detect mutations in the AAVS1 locus from unsorted and sorted cells. H. Diagram showing the AsCpf1 MAFB crRNA in yellow. The PAM sequence for AsCpf1 was highlighted in magenta. I. T7EI assay of mutations in the MAFB locus from transfected cells. J. Design of an AAVS1-targeting (green) and MAFB-targeting (red) crRNA array driven by a U6 promoter. K. T7EI assay to detect mutations in the AAVS1 and MAFB locus from transfected cells.
Clone crRNA encoding oligos to AsCpf1-2A-GFP-U6-crRNA-cloning vector
The CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector and the CMV:AsCpf1-2A-GFP-U6-crRNA-cloning vector can be obtained from Addgene (159281 and 194715, respectively). Use the samples from Addgene to streak LB agar plates supplemented with 100 μg/mL ampicillin and incubate the agar plates at 30 °C overnight. Pick a single colony to inoculate 5 mL of LB medium with 100 μg/mL ampicillin and shake at 220 rpm at 30 °C overnight.
Purify plasmids with the Omega Miniprep kit and measure concentration with a Nanodrop spectrometer.
Set up the following restriction digestion reaction (Table 1).
Table 1. Components for the restriction endonuclease reaction
Reagent Amount
CutSmart Buffer 3 μL
BbsI-HF 0.5 μL (10 units)
Plasmid 3–5 μg
Water Fill up to 30 μL
Incubate the enzymatic reaction at 37 °C for 3 h.
Purify the DNA from the enzymatic digestion using the Zymo DNA Clean & Concentrator and elute in about 25 μL of elution buffer. Measure the concentration using a Nanodrop spectrometer. The DNA sequences between the arrows of Figure 1D will be excised, leaving adhesive ends compatible with the annealed DNA oligo in Figure 1C.
Dilute the DNA oligos to 100 μM and anneal the oligos by setting up the following reaction (Table 2).
Table 2. Components for the oligo annealing reaction
Reagent Amount
Forward oligo (100 μM) 1 μL
Reverse oligo (100 μM) 1 μL
10× T4 DNA ligase buffer 1 μL
T4 PNK 0.5 μL
Water 4.5 μL
Incubate the reaction on a thermocycler at 37 °C for 1 h. Then, anneal the reaction by using the following conditions:
1. 95 °C, 10 min
2. 95–85 °C, −2 °C/s
3. 85 °C, 1 min
4. 85–75 °C, −0.3 °C/s
5. 75 °C, 1 min
6. 75–65 °C, −0.3 °C/s
7. 65 °C, 1 min
8. 65–55 °C, −0.3 °C/s
9. 55 °C, 1 min
10. 55–45 °C, −0.3 °C/s
11. 45 °C, 1 min
12. 45–35 °C, −0.3 °C/s
13. 35 °C, 1 min
14. 35–25 °C, −0.3 °C/s
15. 25 °C, 1 min
16. 25–4 °C, −0.3 °C/s
17. 4 °C, hold
Perform a 10× dilution of the annealed oligos and ligate the annealed oligo with BbsI-digested plasmid by setting up the following reaction (Table 3).
Table 3. Components for the ligation reaction
Reagent Amount
Purified, BbsI-linearized plasmid (30 ng/μL) 2 μL
Annealed oligo (1:10 diluted) 1 μL
10× T4 DNA ligase buffer 0.5 μL
T4 DNA ligase 0.5 μL
Water 1 μL
Ligate at room temperature or 16 °C overnight. Use 1 μL of ligation product to transform Stbl3 chemically competent E. coli based on the manufacturer’s protocol and incubate the LB plates at 30 °C overnight.
Pick 3–6 individual colonies, inoculate 5 mL of LB medium with 100 μg/mL ampicillin, and shake at 220 rpm at 30 °C overnight. Make miniprep from the samples and sequence with U6 primer (5'-GAGGGCCTATTTCCCATGATTCC-3') to determine clones with the correct crRNA sequence insertion.
Make midi-prep of the correct plasmids using ZymoPURE II Plasmid Midiprep kit.
Validation of crRNA by transfecting Cpf1 plasmids to HEK293T cells for genome editing
Culture HEK293T cells in DMEM high glucose medium supplemented with 10% FBS, 1× L-glutamine, 1× MEM-NEAA, and 1× penicillin/streptomycin. Passage the cells 1:5 with Trypsin-EDTA upon reaching near confluency.
Plate HEK293T cells in 12-well plates. Upon 60%–80% confluence, transfect the cells with 0.5 μg of CMV-AsCpf1-2A-GFP-U6-AAVS1-crRNA plasmid (Addgene #194716, Figure 1E) with Lipofectamine 2000 transfection reagent (Invitrogen) according to manufacturer’s guideline.
Three days after transfection, dissociate cells with trypsin, followed by DNA purification with DNeasy Blood & Tissue Kit. Optionally, GFP-expressing cells could be sorted on a FACSAria (Figure 1F) as starting materials for DNA purification with the DNeasy Blood & Tissue Kit.
Set up PCR reactions as below (Table 4).
Table 4. Components for the genotyping PCR
Reagent Amount
Genomic DNA 0.5 μg
AmpliTaq Gold 360 Master Mix 10 μL
Forward primer (5'-CCCCTTACCTCTCTAGTCTGTGC-3') 2 μL 2 μM solution (0.1 μM final concentration)
Reverse primer (5'-CTCAGGTTCTGGGAGAGGGTAG-3') 2 μL 2 μM solution (0.1 μM final concentration)
Water Fill up to 20 μL
The PCR condition is:
One cycle of 95 °C 5 min
35 cycles of 95 °C 30 s
56 °C 30 s
72 °C50 s
One cycle of 72 °C 5 min
Purify the PCR products using a PCR purification kit. Anneal a total of 200 ng PCR products in NEB buffer 2 using the annealing conditions stated above and digest with 5 units of T7EI (NEB) for 1 h at 37 °C before resolving on 10% TBE PAGE gels (Bio-Rad) run at 100 V for 2 h. Observe DNA fragments with staining with ethidium bromide and take images with an AlphaImager gel documentation system (Figure 1G).
Similarly, MAFB-targeting crRNA containing Cpf1 plasmids can be generated with the procedures described above by annealing the following primers: MAFB-crRNA F: 5'-GTAGATTGTGAGTCGTGGCCGGTCCTGGC-3', MAFB-crRNA R: 5'- AAAAGCCAGGACCGGCCACGACTCACAAT-3'. The validation results are in Figure 1H–I using the following T7EI primers: MAFB-T7EI F: 5'-GGGGCTACGCCCAGTCTTGCAGGTATAAACG-3', MAFB-T7EI R: 5'- TCTCTCTCTCCGGCTCTGCTCGAGTCTAGGAGG-3'.
Multiple crRNAs could be generated from a single U6 promoter-driven guide RNA array separated by a spacer. In the example in Figure 1J, the crRNA sequence of the AAVS1 locus (green) and the MAFB locus (red) are separated by a space sequence (black). T7EI assay showed that transfection of the plasmids with the 2crRNAs showed similar indel efficiency compared to each crRNA individually (Figure 1K).
Knockin of the AAVS1 locus HEK293T cells by using the validated crRNA and a targeting construct
An AAVS1-tdT targeting plasmid can be obtained from Addgene (#194728) and purified following the previous procedures.
Transfect HEK293T cells from one well of a 12-well plate with 0.8 μg of targeting vector AAVS1-CAGGS-tdTomato (Figure 2A) and 0.4 μg of AsCpf1-AAVS1 plasmid using Lipofectamine 2000 transfection reagent.
Figure 2. AsCpf1-mediated knockin of a tdTomato-expressing cassette to the AAVS1 locus in HEK293T cells. A. Schematic illustrating the targeting strategy with AsCpf1. B. Experimental timeline to generate HEK293T cells with CAGGS-tdTomato knockin at the AAVS1 locus. C. Expression of tdTomato in AsCpf1-targeted HEK293T cells (bottom panel) but not in the control cells (top panel). Scale bar: 20 µm. D. Representative genotyping PCR results with primer 495 and 489 in A to identify HEK293T cell clones with the CAGGS-tdTomato cassette integrated at the AAVS1 locus. E. Representative Sanger sequencing chromatogram by sequencing the junction region of the PCR amplicon from targeted cells. Sequences inside and outside of the homology arm are shown in red and black, respectively.
Sort GFP and tdTomato (tdT) double-positive cells 3 days post-transfection on a FACSAria. Plate sorted cells at a density of approximately 2,000 cells per 10 cm culture plate and subject to selection with 2 μg/mL puromycin until colonies are formed (Figure 2B).
Mechanically passage individual tdT-expressing colonies to individual wells of a 12-well plate for expansion (Figure 2C).
To genotype HEK293T clones with correct knockin, use 0.5 μg of genomic DNA for a PCR reaction with primer 495 (5'- TCTCTCTCCTGAGTCCGGACC-3') and primer 489 (5'-ACTGAGCTCTCAGGCACCGGGCTTGCGG-3') (Figure 2A) with REDTaq PCR Reaction Mix with the following PCR condition: 95 °C 5 min, followed by 35 cycles of 95 °C 30 s, 57 °C 30 s, 72 °C 1 min 40 s, then 72 °C for 5 min. Resolve PCR products with 1% agarose gels (Figure 2D). Extract DNA from bands of the expected size with a gel purification kit (Omega Bio-tek) and subject to Sanger sequencing with primer 495 (Figure 2E).
Knockout of tdT in HEK293T-tdT cells by a tdT-crRNA
We use the CAGGS-AsCpf1 construct for introducing a crRNA targeting the tdTomato-coding sequence (Figure 3A–3B) and transfected HEK293T cells that stably express tdTomato (HEK293-tdT) (Figure 2C). As expected, we observed expression of GFP and increased tdTomato-low and negative cell proportion in the GFP-positive population compared to control cells (Figure 3C). At 4 days post-transfection, tdTomato-low cells can be observed in the HEK293T-tdTomato cells, particularly in cells with high expression of GFP (Figure 3C). The loss of expression of tdTomato becomes clearer after the propagation of sorted cells with low tdTomato expression (Figure 3D). The following are the experimental procedures.
Design a tdT crRNA to target the 5’ region of the tdT-coding sequence (Figure 3A). Design the oligos for annealing according to Section A.
Anneal the tdT-targeting crRNA oligos and ligate to CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #159281) based on the procedures described in section B, giving rise to the CAGGS-AsCpf1-2A-GFP-U6-tdT-crRNA plasmid (Addgene, #194724).
Transfect the CAGGS-AsCpf1-2A-GFP-U6-tdT-crRNA plasmid to HEK293T-tdT cells based on the procedures described in section C. GFP-expressing cells showed reduced expression of tdT starting from 4 days post-transfection (Figure 3C).
The cells with reduced tdT expression can be isolated with FACS sorting, followed by expansion and isolation of individual clones.
Purify DNA from isolated clones and use it for PCR with the following primers. tdT-forward primer: 5’-GCAACGTGCTGGTTATTGTGCTG-3’; tdT-reverse primer: 5’-TCGCCCTCGCCCTCGATCTCGAA-3’. Purify and sequence PCR amplicons with the tdT-forward primer to identify mutations in the tdT coding sequence.
Figure 3. Disruption of tdTomato-expression in HEK293T-tdT cells with CAGGS-Cpf1 construct that expresses a tdT crRNA. A. Diagram showing the crRNA targeting the tdTomato-coding sequence. The PAM sequence is highlighted in magenta, and the guide sequence is in yellow. B. Diagram of the CAGGS-Cpf1-tdT-crRNA construct. C. Representative micrographs showing the expression of Cpf1 and the tdT crRNA in HEK293T cells reduced tdTomato fluorescence signals 4 days after transfection. Scale bar: 50 µm. D. Flow cytometry analyses showing control HEK293T-tdT cells (left panel), HEK293T-tdT cells expressing AsCpf1 driven by CAGGS promoter (middle panel), and HEK293T-tdT cells expressing AsCpf1 and the tdT crRNA.
Genome editing of hPSCs by ablating expression of tdTomato
The following procedures describe the ablation of tdTomato expression of H1-OCT4-GFP hPSCs with heterozygous integration of CAGGS-tdTomato expression cassette at the AAVS1 locus [25] by using the CAGGS-AsCpf1-2A-GFP-tdT crRNA construct.
Coat plates with gelatin. Prepare the 0.2% gelatin solution by autoclaving 0.8 g of gelatin in 400 mL of MilliQ H2O for 15 min. To coat 6-well plates, add 2 mL of cooled gelatin solution to each well of a 6-well plate and coat overnight. Coating of other tissue culture vessels could be performed by using the gelatin solution proportional to the surface area of the wells.
Generation of mouse embryonic fibroblast cells: All animal experiments were performed following the protocols approved by the Animal Research Regulation Committee at the Whitehead Institute and guidelines from the Department of Comparative Medicine at the Massachusetts Institute of Technology. Mating pairs of DR4 mice [26] were set up, and timed pregnancies were determined by detection of vaginal plugs (E0.5). At E13.5, pregnant DR4 female mice were euthanized by carbon dioxide (CO2) inhalation. After hair clipping of the abdomen and cleaning of exposed skin with 1% Wescodyne solution followed by 70% ethanol, uteri with embryos were dissected using autoclaved surgical tools and put in PBS with penicillin/streptomycin in a biosafety cabinet. Each uterus was transferred to a 10 cm tissue culture plate and the embryo was dissected with autoclaved surgical tools. After removing internal organs and extra-embryonic tissues, the remaining embryonic tissue was minced into small pieces with a scalpel. Then, 0.5 mL of 0.25% trypsin/EDTA was added to each embryo and incubated for 15 min in a 37 °C CO2 incubator. DNaseI solution (10 μL of 10 mg/mL dissolved in PBS) was added, and the tissue was mechanically dissociated by pipetting with a 1 mL pipette until no sticky components remained. Then, cells were transferred to a 15 cm tissue culture plate with 40 mL of MEF medium and incubated in a CO2 incubator as P0 cells. Upon confluency in approximately 5 days, the cells were passaged by 1:5 as P1 and then 1:5 as P2.
Remove the medium from confluent P2 MEF plates and add MEF medium with 5 μg/mL mitomycin C (13 mL for a 15 cm dish). After incubation in a CO2 incubator for 2 h, dispose of the mitomycin-containing medium in a mitomycin C waste container, rinse the cells with PBS+ 3 times, and dispose of the wash buffer in the mitomycin C container. Then, wash the cells with PBS once and harvest cells after trypsinization.
Plate mitomycin C-inactivated MEF cells at a density of 0.4 million cells per well of a gelatin-coated 6-well plate after removing the gelatin solution and culture overnight.
Maintain human pluripotent stem cell line H1-OCT4-GFP with a heterozygous knockin of a CAGGS-tdTomato expression cassette H1-tdT [25] in DMEM-F12 supplemented with 20% KSR, 4 ng/mL FGF2, 1× 2-mercaptoethanol, 1× L-glutamine, 1× MEM-NEAA, and 1× penicillin/streptomycin. Passage cells with 1 mg/mL collagenase IV every 4–6 days on mitomycin C-inactivated MEF feeders cultured in DMEM medium supplemented with 10% FBS, MEM-NEAA, 1× penicillin/streptomycin, and 1× L-glutamine.
Electroporation of plasmids to hPSC: Electroporate the CAGGS-AsCpf1-2A-GFP-U6-tdT-crRNA plasmid (Addgene, #194724) to H1-tdT cells to test the editing of the tdTomato-encoding sequence. Culture hPSCs in the medium supplemented with 10 μM Rho kinase (ROCK) inhibitor Y-27632 overnight before dissociation and electroporation. Also, prepare MEF plates during the day before electroporation. Dissociate approximately 10 million hPSCs into single cells with Accutase, filter with a 40 μm cell strainer, wash with medium containing Y-27632, centrifuge, and resuspend in 800 μL of PBS with 100 μg of Cpf1 plasmid. Load the mixture to a 0.4 cm gap Gene Pulser electroporation cuvette and incubate on ice for 5 min before electroporation with a Gene Pulser Xcell System with 1 pulse of 250 V, 500 μF. Then, incubate the cuvette on ice for 5 min before plating cells on two 6-well plates of MEF. GFP-expressing cells showed reduced expression of tdT starting from 4 days post-transfection (Figure 4A).
Figure 4. AsCpf1-mediated mutation formation in hPSCs. A. Representative fluorescent micrographs of hPSCs electroporated with the corresponding constructs on the left. B. Disruption of tdTomato expression in H1-tdT cells by CAGGS promoter driven AsCpf1 with a tdT crRNA. A 4-nucleotide deletion was identified in cells that lost tdTomato expression but not in control cells. Scale bar: 100 μm.
Manually passage individual colonies onto MEF-coated 12-well plate wells.
To test if the loss of tdTomato expression in cells was caused by mutations of the tdTomato-coding sequence, tdT_Fwd (5'- GCAACGTGCTGGTTATTGTGCTG-3') and tdT_Rev (5'- TCGCCCTCGCCCTCGATCTCGAA-3') primers were used to amplify the potentially edited regions of tdT with PCR. Sanger sequencing showed out-of-frame mutation in electroporated cells but not in control (Figure 4B).
Knockin of hPSC and multiplex genome editing
Maintain WIBR3 hPSCs [27] in DMEM-F12 supplemented with 15% FBS, 5% KSR, 4 ng/mL FGF2, 1× 2-mercaptoethanol, 1× L-glutamine, 1× MEM-NEAA, and 1× penicillin/streptomycin. Passage cells with 1 mg/mL collagenase IV every 4–6 days on mitomycin C-inactivated MEF feeders cultured in DMEM medium supplemented with 10% FBS, MEM-NEAA, 1× penicillin/streptomycin, and 1× L-glutamine.
Electroporation of plasmids to WIBR3 hPSCs: One day before electroporation, culture cells in the medium supplemented with 10 μM Rho kinase (ROCK) inhibitor Y-27632. Dissociate approximately 10 million hPSCs into single cells with Accutase, filter with a 40 μm cell strainer, wash with medium containing 10 μM Y-27632, centrifuge, and resuspend in 800 μL of PBS with 100 μg of CAGGS-AsCpf1-2A-GFP-U6-AAVS1-crRNA plasmid (Addgene, #194723) and 40 μg of AAVS1-tdTomato donor construct (Figure 5A) (Addgene, #194728). Load the mixture to a 0.4 cm gap Gene Pulser electroporation cuvette and incubate on ice for 5 min before electroporation with a Gene Pulser Xcell System with 1 pulse of 250 V, 500 μF. Then, incubate the cuvette on ice for 5 min before plating cells on two 6-well plates of DR4 MEF. Add puromycin-containing medium (0.5 μg/mL) 3–4 days after electroporation (Figure 5B). Manually passage individual colonies onto MEF-coated 12-well plate wells (Figure 5C), expand, and subject to DNA purification.
Purify genomic DNA by isopropanol precipitation. Lyse cells from puromycin-resistant clones in DNA-extraction buffer with freshly added 0.5 mg/mL proteinase K and incubate on a 55 °C rocking platform overnight. Then, add an equivalent volume of isopropanol. Precipitate DNA samples by centrifugation at 4 °C at 16,000× g for 10 min, followed by 70% ethanol wash and centrifugation at the same condition. Dry DNA samples and dissolve in 10 mM Tris (pH 8.5) overnight on a 55 °C rocking platform.
Test the purified genomic DNA with PCR using the primers described in step D5.
Perform Southern blots by digesting 15 μg of purified DNA with 50 units of SphI-HF for 8–10 h at 37 °C in 50 μL. Resolve digested samples in a 0.8% agarose 1× TAE gel with ethidium bromide (EtBr) using HindIII-digested Lamda DNA for approximately 10 h at 75 V. Image the gel with AlphaImager and mark ladder bands with Indian ink. Soak the gel in 0.25 M HCl for 15 min at room temperature on an orbital shaking platform, followed by incubation with 0.4 M NaOH for 15 min at room temperature. Then, transfer the DNA of the gel to the Hybond-XL membrane presoaked in 0.4 M NaOH for 3–4 h or overnight at room temperature by botting with tissue stacks with weight on top.
After the transfer, wash the membrane with 0.2 M Tris buffer (pH 7) at room temperature for 5 min, followed by 2× SSC buffer at room temperature for 5 min. Dry the membrane before prehybridization in hybridization buffer in a hybridization tube placed in a hybridization oven set to 60 °C with constant rotation of the tubes.
Generate AAVS1 internal probe by PCR using AAVS1-tdTomato plasmid as a template and the following primers: 5'- GAATTCGCCCTTTGCTTTCTCTGAC-3', and 5'-TGAGCTCTCGGACCCCTGGAAGAT-3'. Generate AAVS1 external probe by PCR with H1 genomic DNA and the following primer: 5'-ACAGGTACCATGTGGGGTTC-3', and 5'-CCCTTGCCTCACCTGGC GAT-3'. The PCR products were gel purified with E.Z.N.A. Gel Extraction Kit after separation on an agarose gel.
Use Prime-It II Random Primer Labeling Kit to generate radioactive probes and add 100 ng of purified probe DNA to 10 μL of random 9-mer primer, 5 μL of OLB buffer from the kit, and DNase- and RNase-free water to 34 μL. Denature the mixture at a 98 °C heating block for 5 min before chilling on ice for 2 min. Then, add 10 μL of Klenow buffer containing dATP, dGTP, dTTP, but no dCTP, 1 μL of Klenow-Exo- from the kit, and 5 μL (about 50–100 μCi) fresh [a-32P]dCTP (Revvity), mix well, and spin down before incubation at 37 °C for 10–20 min. Purify the labeled probe using CHROMA SPIN + TE-30 Columns based on the product manual.
Denature the purified probe at 95–100 °C for 3 min before putting it on ice for 2 min. Then, add the probe to the pre-hybridized membrane to hybridize with internal or external probes overnight at 60 °C.
Wash the hybridized membrane in 60 °C 2× SSC with 0.2% SDS for 5 min in a water bath shaker.
Then, wash the membrane in 60 °C 0.2× SSC with 0.2% SDS for 15 min in a water bath shaker.
Lastly, wash the membrane in 60 °C 0.2× SSC with 0.5% SDS for 45 min in a water bath shaker.
Dry and wrap the membrane with Saran wrap and expose to autoradiography films such as BioMax MS film or compatible films in an autoradiography cassette in a -80 °C freezer overnight before developing the film (Figure 5D). The first 4 clones are correctly targeted with heterozygous knockin of CAGGS-tdT in the AAVS1 locus.
Immunofluorescence analyses of pluripotency markers in targeted WIBR3-tdT cells are performed according to previous methods [28] with the following antibodies: mouse anti-OCT3/4, 1:100 (BD Transduction Laboratories, 611203); rabbit anti-NANOG, 1:100 (Thermo Fisher Scientific); mouse anti-TRA-1–60, 1: 100, Life Technologies); mouse anti-SSEA4 antibody, 1:100 (Thermo Fisher Scientific, clone MC-813-70). Alexa 488 conjugated secondary antibodies (Life Technologies) were used at 1:400. Targeted hPSC clones expressed the pluripotency markers as control WIBR3 cells (WIBR3 CTL, Figure 5E).
Multiplex genome editing experiments targeting AAVS1 and MAFB are performed in the same way, except using the CAGGS-AsCpf1-2A-GFP-U6-AAVS1-MAFB-crRNA (Addgene, #194725) instead of #194723. The correctly targeted clones are identified with PCR and sequencing validation of the correct junction sequences (Figure 5F-5G). Then, correct clones are further verified with Southern blotting, and the correctly targeted clones are labeled in green (Figure 5H).
Next-generation sequencing is used to examine gene editing of MAFB in AAVS1 and MAFB multiplex experiments by using the same reverse primer (R NGS seq primer for MAFB of WIBR3: 5’- GCAGGGACAGGGTCCGGGGTAG-3’) and differentially barcoded forward primers (primers in the reagent lists). Set up PCR reactions based on the conditions described in step C4 and 30 s of amplification time. Resolve amplicons on a 1.5% agarose gel followed by gel purification. Dissolve purified PCR products in 35 µL of 10 mM Tris buffer (pH 8.0) at a concentration of 10–20 ng/µL. Samples were pooled and submitted to CCIB DNA Core Facility at Massachusetts General Hospital (Cambridge, MA) for CRISPR-sequencing. Briefly, partial Illumina adaptor sequences and unique identifiers (barcodes) were attached by ligation to both the 5’ and 3’ ends of the PCR amplicons. A subsequent low-cycle PCR amplification step was used to complete the addition of full-length adaptor sequences. Paired-end sequencing (2 × 150 bp) of the resulting TruSeq-compatible paired-end Illumina libraries was performed on an Illumina MiSeq platform, using V2 chemistry. The NGS data are in Figure 5I–5J.
Teratoma assays are performed to test if targeted hPSCs could form teratoma with three germ layers. Single-plex and multiplex targeted hPSCs, each from two near-confluent six-well plates, are dissociated with collagenase and resuspended in DMEM without serum. The cells are injected intramuscularly into NSG mice between 8 and 12 weeks of age using 1 mL syringes with 23-gauge needles. The formation of teratomas is monitored twice weekly for about 2 months. Palpable tumors reaching between 7 and 10 mm in diameter are dissected and fixed in buffer-neutralized formalin solution. Paraffin embedding and histological analysis using hematoxylin and eosin (H&E) staining are performed with standard protocols. Targeted hPSCs from both methods generated teratomas with three germ layers including pigmented ectodermal cells, cartilage-like mesodermal cells, and glandular endodermal cells (Figure 5K). All animal experiments were performed following the protocols approved by the Animal Research Regulation Committee.
Figure 5. Multiplex genome editing in hPSCs with AsCpf1. A. Schematic illustrating the strategy for knockin at the AAVS1 locus in hPSCs with Cpf1. B. Timeline for the experiments. C. Expression of tdTomato in a representative knockin clone but not in control. D. Verification of correct knockin with Southern blot with internal probe (top panel) and external probe (bottom panel). Clones showing correct knockin are labeled in green. E. Expression of pluripotency markers OCT4, NANOG, TRA-1-60, and SSEA4 in targeted hPSC clones. F. PCR-based characterization of knockin at the AAVS1 locus in WIRB3 with the guide RNA array shown in Figure 1J. G. Example of Sanger sequencing chromatogram showing the correct junction of the sequence not in the homologous arm of the targeting construct (black letters) and sequence in the homologous arm (red letters). H. Southern blotting analyses of knocking of the AAVS1 locus with the guide RNA array shown in Figure 1J. I. Diagram showing the distribution of knockin and MAFB mutations in isolated hPSC clones. Each vertical bar pair represents an isolated hPSC clone. J. Distribution of different mutation alleles in the MAFB gene upon AsCpf1 mediated editing with the guide array in Figure 1J. K. H&E staining of teratoma tissues formed upon injection of WIBR3-tdTomato cells (top panels) or WIBR3-602-1 cells into immunocompromised mice. Scale bars in C, E, and K: 100 μm.
Differentiation to neural precursors neurons
Neuronal differentiation is performed based on the published protocol [29].
For neural progenitor cells (NP) differentiation, plate a total of 5 million dissociated WIBR3 or WIBR3-tdT cells to one Matrigel-coated 6-well plate well in NGD medium supplemented with 10 ng/mL FGF2, 1:500 insulin, and 2.5 μM dorsomorphin (R & D Systems) for 14 days with daily medium changes to keep medium from turning very yellow. Passage NP cells every 3 days in NGD medium with 10 ng/mL FGF2, 1:500 insulin.
For neuronal differentiation, plate about 0.2 million NP cells in each 12-well plate well in 1 mL of NGD medium and change the medium every 2–3 days. Fix NP and neurons after 4–6 weeks of differentiation using the immunofluorescence protocols described above in G14. Nestin expression showed proper NP differentiation of WIBR3-tdT cells (Figure 6A), and neuronal maker TUBB3 was expressed similarly in unedited WIBR3 and Cpf1 edited WIBR3-tdT cells (Figure 6A), indicating Cpf1 editing did not affect neural differentiation of hPSCs.
Figure 6. Differentiation of Cpf1-edited WIBR3-tdT cells to neural precursor cells and neurons. A. WIBR3-tdT cells were differentiated into neural precursor (NP) cells for 14 days. Then, NP cells were passaged, fixed, and stained with an anti-Nestin antibody (1:100, SC-23927). The formation of tdT-expressing (red) neural rosette structures (green) was shown. B. The WIBR3-tdT-differentiated NP cells were differentiated toward neurons with growth factor withdrawal for four weeks. Neurite structures were shown by anti-TUBB3 (1:1,000) staining. Scale bars in A and B: 100 μm.
Generating INS reporter hPSC cells
The INS crRNA (Figure 7A) was designed, generated, and validated in a similar way as described before. Electroporation was performed as described before. The generated CAGGS-AsCpf1-2A-GFP-U6-INS-crRNA plasmid is available from Addgene (159283). The primers 417-1 (5'- TGGGGCAGGTGGAGCTGGGCGG-3') and primer 418-1 (5'-ACCACCCCTGGCCCCTCAGAGACC-3') were used for T7EI assays.
The H1-OCT4-GFP hPSCs [30] were cultured in the same way as H1-tdT described before.
The INS-2A-luciferase-2A-tdT donor plasmid (Figure 7B) was described before [5] and is available from Addgene (159348). To target the INS locus in H1-OCT4-GFP, electroporation was performed using the procedures described before by using 100 μg of CAGGS-AsCpf1-2A-GFP-U6-INS-crRNA and 40 μg of donor construct INS-2A-luciferase-2A-tdT. Add the puromycin-containing medium (0.5 μg/mL) 3–4 days after electroporation. Manually passage individual colonies onto MEF-coated 12-well plate wells, expand, and genotype with PCR using the primer 662 (5'-GGCCTTTGGTGCAGTGACCAGAGTGTCAGG-3') and primer 663 (5'- GATTCTCCTCGACGTCACCGCATGTTAGC-3'). Subject clones with correct knockin to Southern blotting analysis.
For the INS targeted cells, amplify the internal probe with PCR with the following primers and firefly luciferase gene as a template: forward: 5'-ATGGAAGACGCCAAAAACATAAAGAAAGGCCC-3'; reverse: 5'-CACGGCGATCTTTCCGCCCTT-3'. Amplify the external probe with PCR with the following primers and genomic DNA from H1-OCT4-GFP hPSCs: forward: 5'-GACTCCCCACTTCCTGCCCATCT-3'; reverse: 5'-TCTTCTCCCAGCCCCGTCCTCAC-3'. The Southern blotting process is based on the procedures described in section G. Clones with correct heterozygous knockin were 1, 2, 7, 8, 13, and 15 in Figure 7C. Note that the control H1-OCT4-GFP has two different bands for the external probe, possibly because of the different sequences of the two alleles at the INS locus.
Figure 7. AsCpf1-mediated targeting in the INS locus in hPSCs enables monitoring of INS expression. A. The design of an INS targeting crRNA. The PAM sequence is highlighted in magenta, and the guide sequence is highlighted in yellow. B. The targeting strategy for generating INS luciferase and tdTomato reporter INS-luciferase-tdT allele by knocking the 2A-luciferase-2A-tdt cassette to the 3' end of the INS coding sequence. C. Southern blots with internal probe (left panel) and external probe (right panel) to identify clones with a single copy of the 2A-luciferase-2A-tdt cassette inserted into the INS locus. Correct clones were labeled in green. The double bands in the control samples for the external probe (right panels) correspond to two wild-type INS alleles of the H1-OCT4-GFP cells. D. Expression of tdTomato in cells during the differentiation of the INS-luciferase-tdT reporter hPSCs toward pancreatic β-like cells. E. Representative confocal micrographs of INS-luciferase-tdT hPSC differentiated β-like cells with anti-tdTomato staining (red) and anti-C-peptide staining (green). F. Quantitative RT-PCR analysis of sorted tdTomato positive cells (red bars) showed significantly higher expression of INS and luciferase compared to sorted tdTomato negative cells (grey bars) (* P < 0.05, t-test, n = 3 experiments). G. β-like cells specific luciferase signal at stage 6, day 7. Scale bars in D and E: 50 μm.
Differentiation of β-cells
Differentiation of β-cells was based on a published protocol [5] with the following steps.
Adapt H1-OCT4-GFP-INS-tdT hPSC cells for feeder-free conditions by culturing in Matrigel-coated tissue culture plates in mTeSR. For coating plates, dilute Matrigel 1:100 in cold DMEM-F12 medium. Then, add 2 mL of diluted Matrigel solution to one well of a 6-well plate well and coat overnight at room temperature before pre-warming in a 37 °C incubator for 1 h before using. Culture cells in mTeSR medium in 6-well plates and passage every 3–4 days with a 1:4–6 passage ratio. To prevent differentiation, passage cells before complete confluency.
Treat PSCs with mTeSR medium supplemented with 10 μM Rho kinase (ROCK) inhibitor Y-27632 overnight. Then, dissociate the cells into single cells with Accutase. Dislodge cells by tapping the sides of the plates and add RI-supplemented mTeSR to dissociate cells into single cells. After cell counting with Countess, plate about 1.8 million live cells to each well of a 6-well Matrigel-coated plate in mTeSR medium supplemented with 10 μM ROCK inhibitor Y-27632.
After overnight culture, replace the Y-27632-containing mTeSR medium with the mTeSR medium. Differentiation is initiated when cells reach around 90% confluency (usually between 1–2 days after plating cells). Wash cells with PBS- and then perform daily medium changes as follows.
S1 (3 days) base medium: MCDB131 with 0.5% FAF-BSA, 1.5 g/L NaHCO3, 10 mM glucose, 1× GlutaMAX, 1× penicillin/streptomycin. Freshly supplemented factors: 100 ng/mL Activin A and 3 μM CHIR99021 (day 1); 100 ng/mL Activin A, and 0.3 μM CHIR99021 (day 2); 100 ng/mL Activin A (day 3).
S2 (2 days) base medium: MCDB131 with 0.5% FAF-BSA, 1.5 g/L NaHCO3, 10 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, and 0.25 mM Vc. Freshly supplemented factors: 50 ng/mL KGF.
S3 (2 days) base medium: MCDB131 with 2% FAF-BSA, 1.5 g/L NaHCO3, 10 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, 0.25 mM Vc, and 0.5× insulin-transferrin-selenium-ethanolamine. Freshly supplemented factors: 50 ng/mL KGF, 0.25 μM SANT-1, 1 μM retinoic acid, 100 nM LDN193189, and 200 nM TPB.
S4 (4 days) base medium: MCDB131 with 2% FAF-BSA, 1.5 g/L NaHCO3, 10 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, 0.25 mM Vc, and 0.5× insulin-transferrin-selenium-ethanolamine. Freshly supplemented factors: 50 ng/mL KGF, 50 ng/mL EGF, 0.25 μM SANT-1, 0.1 μM retinoic acid, 200 nM LDN193189, and 100 nM TPB. For the last medium change, 10 μM Y-27632 was added to the S4 medium.
S5 (3 days) starts with a transition from 2D culture to 3D culture. Cells are washed with PBS-, dissociated with Accutase, washed with H1 S5 base medium (see below), and about 6 million live cells are plated in 3 mL of S5 medium (see below) supplemented with 10 μM Y-27632 in each well of a 6-well AggreWell 400 plates. After overnight culture, move the clusters to 6-well ultra-low adherent culture plates placed on an orbital shaker set at 95–100 rpm for 2 more days in S5 medium. The rest of the differentiation is carried out in ultra-low attachment plates placed on an orbital shaker with the same setting.
S5 base medium: MCDB131 with 2% FAF-BSA, 1.5 g/L NaHCO3, 20 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, 0.25 mM Vc, 0.5× insulin-transferrin-selenium-ethanolamine, 10 µg/mL heparin, and 10 μM ZnSO4. Freshly supplemented factors: 10 μM ALK5 inhibitor II, 0.25 μM SANT-1, 0.05 μM retinoic acid, 100 nM LDN193189, and 1 μM T3.
S6 (7 days) base medium: MCDB131 with 2% FAF-BSA, 1.5 g/L NaHCO3, 20 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, 0.25 mM Vc, 0.5× 100× insulin-transferrin-selenium-ethanolamine, 10 µg/mL heparin, 10 μM ZnSO4. Freshly supplemented factors: 10 μM ALK5 inhibitor II, 100 nM gamma-secretase inhibitor XX, 100 nM LDN193189, and 1 μM T3.
S7 (7–14 days) medium: MCDB131 with 2% FAF-BSA, 1.5 g/L NaHCO3, 8 mM glucose, 1× penicillin/streptomycin, 1× GlutaMAX, 1× MEM-NEAA, 10 μg/mL heparin, 10 μM ZnSO4, 1× trace element A, and 1× trace element B.
The INS-tdT-targeted hPSCs are tdTomato negative at the pluripotent stem cell state. During the differentiation of the targeted INS reporter hPSCs toward β-cells, tdTomato signal emerges at day 2 of stage 5 (Figure 7D, S5D2), and more tdTomato expressing cells are detected during later stages (Figure 7D). Immunofluorescence analyses show that C-peptide expression cells also expressed tdTomato (Figure 7E). Quantitative RT-PCR analysis of cells sorted based on tdTomato expression levels show high levels of INS and luciferase expression in tdTomato-positive cells compared to tdTomato-negative or lowly expressing cells (Figure 7F). To test the expression of luciferase, 15 mg/mL D-luciferin is added to undifferentiated hPSC or INS reporter cells differentiated to β-cells. After mixing for 1 min, luciferase signals are acquired with an IVIS Spectrum imager (Xenogen) with an exposure of 1 min and mid-bin setting. Imaging quantification is performed with Living Image (Version 4.5.4). A control well is used to subtract the background signal from the measurements of samples. Consistent with the qPCR results (Figure 7F), specific luciferase signals were identified in differentiated β-like cells but not in undifferentiated pluripotent stem cells (Figure 7G).
Test potential off-targeting effects
The predicted off-target sites can be obtained from http://www.rgenome.net/cas-offinder with the following guide sequences (PAM sequences in italic font, position allowing for variable bases in bold). Sequences showing most similarities are shown with differences in lower case. No less than 4-nucleotide differences off-target sites were identified.
AAVS1: TTTTATCTGTCCCCTCCACCCCACAGT
MAFB: TTTCTGTGAGTCGTGGCCGGTCCTGGC
INS: CTCCTGCACCGAGAGAGATGGAATAAA
PCR reactions with primers (AAVS1 off 1 to INS off 1 in the primer list) were performed with 0.5 μg of genomic DNA from parental and targeted pluripotent stem cell clones. PCR products were resolved with 1% agarose gel followed by purification with a gel purification kit and Sanger sequencing with the sequencing primers listed in Table 5.
Table 5. Experimental evaluation of off-targeting effects of Cpf1-mediated genome editing in hPSCs. The nucleotides in lower case denote the differences between the potential off-target site and the corresponding crRNA.
Putative off target sequence Genomic position* (direction) Mutation/tested
AAVS1 crRNA: TTTNTGTGAGTCGTGGCCGGTCCTGGC
DNA: TTTCTGTGAGTCcTGGCCctTCCTGGa
7: 131811-131837 (-) 0/5
AAVS1 crRNA: TTTNTGTGAGTCGTGGCCGGTCCTGGC
DNA: TTTTATCTcTCCCCTCCACCCaAaAcT
13: 91233988-91234014 (+) 0/5
AAVS1 crRNA: TTTNATCTGTCCCCTCCACCCCACAGT
DNA: TTTCATCTcTCCCCTaCtCCCCACtGT
2: 18590645-18590671 (+) 0/5
AAVS1 crRNA: TTTNATCTGTCCCCTCCACCCCACAGT
DNA: TTTTATCTtcCCgCTgCACCCCACAGT
2: 235878805-235878831 (-) 0/5
AAVS1 crRNA: TTTNATCTGTCCCCTCCACCCCACAGT
DNA: TTTAATtTtgCCaCTCCACCCCACAGT
6: 123920988-123921014 (-) 0/5
MAFB crRNA: TTTNTGTGAGTCGTGGCCGGTCCTGGC
DNA: TTTATGTGAaTCaTGGCCaGTCCTGtC
11: 134335239-134335265 (+) 0/10
MAFB crRNA: TTTNTGTGAGTCGTGGCCGGTCCTGGC
DNA: TTTCTGTGAGTCcTGGCCctTCCTGGa
7: 131811-131837 (-) 0/10
INScrRNA: TTTNTTCCATCTCTCTCGGTGCAGGAG
DNA: TTTATTCCATCTCTCTgctTcCAGGAG
11: 62432165-62432191 (+) 0/6
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Ma et al. [5]. Human T-cells expressing a CD19 CAR-T receptor provide insights into mechanisms of human CD19 positive b-like cell destruction. Cell Rep Med. (Figure 1 and Figure S1).
General notes and troubleshooting
General notes
Working with hPSCs could require intuitional approval and appropriate MTA. Research needs to adhere to approved protocols that follow the ISSCR guidelines regarding hPSC research. Keeping hPSC cell density from reaching complete confluency reduces the chance of differentiation.
All work involving vertebrate animals including mice requires Institutional Animal Care and Use Committee (IACUC) approval.
Appropriate licenses for working with radioactive isotopes, comprehensive training, protective equipment, and users’ dosimeters are needed to work with radioactive Southern blotting probes. Digoxigenin-labeled probes could be an alternative approach for radioactive probes.
For efficient editing of hPSCs, the CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #159281) is recommended. If the transfected cells do not express GFP, it is helpful to FACS sort GFP-expressing cells to enrich cells that express Cpf1.
For cell lines such as HEK293T cells, both CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #159281) and CMV:AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #194715) can be used. Lower editing efficiency could be observed compared to Cas9-mediated gene editing using T7EI assays.
The AAVS1-tdT targeting construct (Addgene, #194728) was modified based on AAVS1-tdT (Addgene, #159275) by modifying a segment with high sequence similarity to the Cpf1 AAVS1 crRNA to avoid potential digestion of the targeting construct by Cpf1.
The starting cell density for differentiation of β-cells would need optimization. For H1-OCT-GFP cells, Figure 8A shows a suitable density to start differentiation, whereas Figure 8B and Figure 8C show a lower and higher density than the optimal condition, respectively. The cellular morphologies of the first two steps were published before (Figure 1A, first three images of [31]). A representative image after pancreatic precursor 2 (PP2) is shown in Figure 8D.
Figure 8. Cell density optimization for starting β-cell differentiation. A. Suitable cellular density for differentiation. B. Representative cellular density that is lower than optimal densities. C. Representative cell density that is higher than optimal densities. E. Representative image of PP2 cells before dissociation to form spheroids. Scale bar: 50 μm.
Troubleshooting
Problem 1: Low editing efficiency of hPSCs.
Possible cause: Insufficient expression of Cpf1.
Solution: Use the CAGGS promoter to drive Cpf1 expression by cloning crRNA to the CAGGS-AsCpf1-2A-GFP-U6-crRNA-cloning vector (Addgene, #159281). Increase the amount of plasmids used in electroporation or transfection. Furthermore, including the crRNA targeting your gene of interest in a crRNA assay together with AAVS1 crRNA (illustrated in Figure 1J) and performing simultaneous targeting of the AAVS1 locus using a donor plasmid that allows puromycin selection (Figure 5A) can assist the identification of edited hPSC clones.
Problem 2: Poor survival rate of hPSCs after electroporation.
Possible cause: Insufficient ROCK inhibitor treatment and non-optimal MEF plates.
Solution: Before dissociation and electroporation, hPSCs are treated with a final concentration of 10 µM ROCK inhibitor-supplemented hPSC medium overnight. After electroporation, the hPSCs should be plated in the medium with 10 µM ROCK inhibitor for overnight attachment to MEF feeder cells. The plates should have a confluent monolayer of MEF cells. It is important to wash off mitomycin C after MEF inactivation. MEF inactivated with irradiation (3000 rad) could also be used as feeder cells, but they tend to last shorter than mitomycin C-inactivated MEF.
Acknowledgments
The authors wish to thank Prof. Rudolf Jaenisch and members of Prof. Jaenisch’s laboratory for advice and discussion, Prof. Feng Zhang’s laboratory for sharing pX458 and pY010 plasmid through Addgene, Dr. Huan Yang of Prof. Hongkui Deng’s laboratory for sharing INS targeting construct, Patrick Autissier, Eleanor Kincaid, and Hanna Aharonov of FACS core laboratory of Whitehead Institute for sorting cells, Gary Rogers of the Whitehead Institute for assistant with medium preparation, the Center for Computational and Integrative Biology (CCIB) at Massachusetts General Hospital for the use of the CCIB DNA Core Facility (Cambridge, MA), which provided NGS and data analysis service, and Arend Vischer of the Ma’s laboratory for feedback on this work. This study was supported by a generous gift from Liliana and Hillel Bachrach and in part by the NIH (RO1-CA084198) to R.J., and the University of Illinois start-up funding to H.M.
Competing interests
H.M. declares no competing interests.
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Article Information
Publication history
Received: Jun 27, 2024
Accepted: Sep 6, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Stem Cell > Pluripotent stem cell > Regenerative medicine
Cell Biology > Cell engineering > CRISPR-cas9
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Preparation of Polarity-Marked Microtubules Using a Plus-End Capping DARPin
GH Gil Henkin
CB Cláudia Brito
AP Andreas Plückthun
TS Thomas Surrey
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5109 Views: 440
Reviewed by: Manish ChamoliMarcus Braun Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Aug 2023
Abstract
The eukaryotic cytoskeleton is formed in part by microtubules, which are relatively rigid filaments with inherent structural polarity. One consequence of this polarity is that the two ends of a microtubule have different properties with important consequences for their cellular roles. These differences are often challenging to probe within the crowded environment of the cell. Fluorescence microscopy–based in vitro assays with purified proteins and stabilized microtubules have been used to characterize polarity-dependent and end-specific behaviors. These assays require ways to visualize the polarity of the microtubules, which has previously been achieved either by the addition of fluorescently tagged motor proteins with known directionality or by fluorescently polarity marking the microtubules themselves. However, classical polarity-marking protocols require a particular chemically modified tubulin and generate microtubules with chemically different plus and minus segments. These chemical differences in the segments may affect the behavior of interacting proteins of interest in an undesirable manner. We present here a new protocol that uses a previously characterized, reversibly binding microtubule plus-end capping protein, a designed ankyrin repeat protein (DARPin), to efficiently produce polarity-marked microtubules with different fluorescently labeled, but otherwise biochemically identical, plus- and minus-end segments.
Key features
• Produces polarity-marked microtubules with biochemically identical segments.
• Allows analysis of end-specific and polarity-dependent activities of purified microtubule-associated proteins
• Requires purified microtubule plus-end capping DARPin (D1)2
• Concentrations optimized for porcine brain tubulin
Keywords: Microtubules In vitro reconstitution Fluorescence microscopy Microtubule polarity Microtubule-associated proteins (MAPs) Molecular motors DARPins
Graphical overview
Background
Microtubule filaments have a structural polarity determined by the orientation of their constituent α/β-heterodimers. When grown in vitro from purified tubulin, microtubules stochastically switch between growing and shrinking at both ends with distinct dynamics [1,2]. Plus-ends, where β-tubulin is exposed, typically grow faster and switch to the shrinking state more frequently than the minus-ends [2,3]. In cells, where a growing list of proteins has been shown to preferentially interact with plus- or minus-microtubule ends [4–6], the minus-end is often, but not always, stabilized by the microtubule nucleating γ-tubulin ring complex [7–9], whereas the plus-end is subject to control by various proteins that control dynamic instability rates [4]. However, the extent to which microtubule-associated proteins selectively interact with one or the other microtubule end is often hard to determine. This is due to the high density of microtubules in structures such as the mitotic spindle or in neuronal axons relative to the resolution of fluorescence microscopy typically used to observe proteins in live cells.
Biochemical assessment in vitro, with purified proteins of interest, can readily reveal end-specific behaviors, amenable to straightforward kinetic and dynamic analyses that are often impossible in cells [4,10–17]. Determining this behavior requires being able to determine unambiguously the polarity of individual microtubules, which is often done through differential fluorescent marking [18–20]. However, the majority of protocols published for selective marking involve use of tubulin chemically modified with N-ethylmaleimide (NEM) [18,19,21], which blocks microtubule growth from the minus-end but is also inhibitory to growth at the plus-end, even if less so, requiring careful optimization of modified and unmodified tubulin concentrations and resulting in microtubules that are not fully polymerization-competent in subsequent assays. Furthermore, the protocols involve two steps with two different stabilizing agents, with highly distinct mechanisms of stabilization—the slowly-hydrolyzing GTP analog guanosine-5'-[(α,β)-methyleno]triphosphate (GMPCPP) and the microtubule-binding drug paclitaxel (taxol). This means that the two segments of the microtubule lattice, as well as the ends, are also distinct—one segment contains only GMPCPP-tubulin, while the other contains only taxol-stabilized, partially NEM-modified GDP-tubulin, with differences in chemistry, structure, and mechanical properties [22–27]. These differences can confound the analysis of protein binding to the different microtubule segments and the behavior of potentially end-specific microtubule-binding proteins. Indeed, various microtubule-associated proteins are known to exhibit strikingly different behaviors depending on the nucleotide state of the microtubule lattice [28–31].
We were interested in updating this protocol in such a way that we could produce polarity-marked microtubules where both segments were otherwise biochemically identical. Our procedure is based on recent work that has identified de novo–designed proteins that reversibly cap the growth of distinct microtubule ends [32–34]. We use one of these proteins, a designed ankyrin repeat protein (DARPin) in a tandem arrangement, termed (D1)2, to provide a new method for polarity marking stable microtubules that are grown with GMPCPP-only throughout. Furthermore, we show how to confirm the polarity of the microtubules, using a simple gliding assay with a microtubule motor protein of known directionality.
Materials and reagents
Reagents
Porcine brain tubulin, prepared as previously reported [35,36]
Purified tandem DARPin D1 protein [(D1)2], expressed in E. coli and purified according to standard DARPin expression and purification protocols [37,38]. A detailed description of the (D1)2 expression on a 500 mL scale and purification via metal affinity chromatography and size exclusion chromatography can be found here [14,32]. The bacterial (D1)2 expression plasmid can be requested from A. Pluckthun ([email protected])
Fluorescently labeled porcine brain tubulin (i.e., AlexaFluor647 or Atto647-labeled) prepared as previously reported [21,35]
Note: Fluorescence labeling ratio should be determined by Nanodrop prior to starting the protocol.
Optional for neutravidin-biotin-based immobilization in, e.g., TIRF microscopy assays: biotinylated porcine brain tubulin, prepared as previously reported [21,35]
Piperazine-N,N'-bis(2-ethanesulfonic acid) (PIPES) (Sigma-Aldrich, catalog number: P6757)
Magnesium chloride (MgCl2), 1 M (Sigma-Aldrich, catalog number: 63069)
Ethylene glycol-bis(β-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) (Sigma-Aldrich, catalog number: E3889)
Guanosine-5'-[(α,β)-methyleno]triphosphate (GMPCPP), 10 mM (Jena bioscience, catalog number: NU-405)
High-quality purified water, e.g., Millipore Milli-Q
For gliding assay:
An active, directional microtubule motor protein; we had the minus-end-directed HSET motor protein available in the laboratory, prepared as previously published [39]. Many other microtubule motors with established polarity, such as the plus-end directed kinesin-1 or its motor-domain truncations, can be used instead
Poly(L-lysine)-polyethylene glycol (PLL-PEG) [SuSoS, catalog number: PLL(20)-g[3.5]- PEG(2)]
β-Mercaptoethanol (β-ME), 14.3 M (Sigma-Aldrich, catalog number: M6250)
β-casein (Sigma-Aldrich, catalog number: C6905)
ATP (Sigma-Aldrich, catalog number: A6419)
Solutions
EGTA, 0.5 M, pH 8 (see Recipes)
BRB80, 5× stock (see Recipes)
BRB80, 1× (see Recipes)
For gliding assay:
PLL-PEG, 2 mg/mL (see Recipes)
β-casein, 25 mg/mL (see Recipes)
ATP, 100 mM, pH 7 (see Recipes)
Gliding buffer (see Recipes)
Recipes
Note: All solutions should be prepared using high-quality purified water, e.g., Millipore Milli-Q.
EGTA, 0.5 M, pH 8
Add solid KOH to dissolve EGTA powder and adjust pH only by adding KOH, being careful not to overshoot; store at room temperature.
BRB80, 5× (50 mL)
Reagent Final concentration Quantity or Volume
PIPES 400 mM 6.06 g
EGTA (0.5 M, pH 8.0) 5 mM 0.5 mL
MgCl2 (1 M) 5 mM 0.25 mL
Total 50 mL, pH 6.8
Filter (0.22 μm pore size) and store for up to 4 weeks at 4 °C.
Note: PIPES dissolves upon the addition of solid KOH. MgCl2 and EGTA should be added only after PIPES is dissolved (requiring ~1.35 g of KOH for these amounts). The final pH should be adjusted only by addition of KOH, being careful not to overshoot, to control the total ionic strength of the solution.
BRB80, 1×
Diluted from the above recipe on the day of use; keep on ice.
PLL-PEG, 2 mg/mL, in water
Aliquot and store at -20 °C.
β-casein, 25 mg/mL, in BRB80
Ultracentrifuge, aliquot, snap-freeze in liquid nitrogen, and store at -80 °C.
ATP, 100 mM, pH 7
Filter, aliquot, and store at -80 °C.
Gliding buffer
BRB80 with 5 mM β-ME, 1 mM ATP, 1 mg/mL β-casein; prepare on the day of the experiments and keep at room temperature.
Laboratory supplies
1.5 mL centrifuge tubes (e.g., Eppendorf, catalog number: 0030120086)
Ice bucket
Pipettes
Pipette tips
For imaging/ gliding assay:
Cover glasses, thickness #1.5, 22 × 22 mm (VWR, catalog number: 630-1843)
Standard microscopy slides, 76 × 26 mm (e.g., VWR, catalog number: 631-1550P)
Double-sided adhesive fleece transparent tape, 10 m × 15 mm (Tesa®, catalog number: 053380000001)
Filter paper 3 mm CHR, 46 cm × 57 cm (Whatman®, catalog number: 3030-917)
KorasilonTM Paste, medium viscosity (Sigma-Aldrich, catalog number: 769681-1EA)
Diamond scribing pen (e.g., Ted Pella, catalog number: 54468)
Forceps (e.g., Sigma-Aldrich, catalog number: Z680214-1EA)
Equipment
Microcentrifuge (e.g., Eppendorf, model: 5418 R)
Heat block set to 37 °C (e.g., Eppendorf, model: ThermoStat C)
For imaging/ gliding assay:
Fluorescence microscope (e.g., Nikon Instruments, model: E600)
Software and datasets
ImageJ/FIJI (e.g., ImageJ version 1.54, https://imagej.net/ij/index.html)
Procedure
Dim, GMPCPP-stabilized microtubules
Thaw aliquots of frozen unlabeled tubulin, fluorescent tubulin, and (optionally) biotinylated tubulin. On ice, prepare a mixture of 3 μM total tubulin in BRB80 1×, supplemented with 0.5 mM GMPCPP, including fluorescent tubulin at a final labeling ratio of 3%–5% and (optionally) 18% biotinylated tubulin, with a total volume of 200 μL in a microcentrifuge tube.
Incubate mixture on ice for 5 min.
Transfer the tube to 37 °C and incubate for 1 h. Warm 3 mL of BRB80 1× to 37 °C for subsequent steps.
After incubation, dilute seeds with 170 μL of warm BRB80 1× and centrifuge at 17,000× g for 10 min at room temperature in the tabletop microcentrifuge.
Remove the supernatant.
Note: You should be able to see a pellet with color depending on the fluorescent dye used.
Wash the pellet by adding 400 μL of warm BRB80 1× and centrifuging again at 17,000× g for 10 min at room temperature.
Remove the supernatant and resuspend the pellet in 50 μL of warm BRB80 1×. Keep microtubules at room temperature.
Bright GMPCPP-stabilized minus-end extensions using DARPin (D1)2
Thaw an aliquot of DARPin (D1)2 and dilute to 2.5 μM in BRB80 1× on ice.
From freshly thawed aliquots of unlabeled tubulin, fluorescent tubulin, and (optionally) biotinylated tubulin, prepare a new “bright” mixture of 3 μM total tubulin in BRB80 1×, supplemented with 0.5 mM GMPCPP, including fluorescent tubulin at a final labeling ratio of 10%–15% and (optionally) 18% biotinylated tubulin, with a total volume of 200 μL in a microcentrifuge tube.
Incubate the “bright” mixture on ice for 5 min.
During the incubation, add 2 μL of the DARPin (D1)2 dilution to 8 μL of the previously prepared dimly-labeled microtubules at room temperature. The protein will bind the microtubule plus-ends, ensuring that the subsequent bright extensions are exclusively from the microtubule minus-ends.
Dilute the “bright” mixture with 200 μL of cold BRB80 1×.
Transfer 90 μL of the diluted “bright” mixture to 37 °C and incubate for 1 min.
Add the 10 μL DARPin (D1)2/dim seed mixture directly to the warmed “bright” mixture [resulting in final concentrations of 50 nM DARPin (D1)2, 1.35 μM tubulin, and 0.23 mM GMPCPP].
Incubate the mixture at 37 °C for 1 h. Brightly labeled, stabilized extensions will grow from microtubule minus-ends, as the plus-ends are capped, resulting in microtubules with two distinctly labeled segments.
After incubation, dilute seeds with 170 μL of warm BRB80 1× and centrifuge at 17,000× g for 10 min at room temperature in the tabletop microcentrifuge.
Remove the supernatant, including unpolymerized tubulin, GMPCPP, and the capping protein, DARPin (D1)2.
Note: There are fewer seeds at this point, so the pellet may be challenging to see.
Wash the pellet by adding 400 μL of warm BRB80 1× and centrifuging again at 17,000× g for 10 min at room temperature.
Remove the supernatant and resuspend the pellet in 50 μL of warm BRB80 1×. Keep microtubules at room temperature. Polarity-marked microtubules should be used within the day.
Quick check: properly segmented microtubules
Using a fluorescent microscope, the success of the protocol can be checked by quickly preparing a 1:6 μL dilution of the microtubules and “squashing” the dilution between a coverslip and slide. Fluorescence imaging will readily show how many of the dim stabilized microtubules have a single bright extension. (Figure 1).
Figure 1. AlexaFluor647-labeled, polarity-marked microtubules on a glass slide. Imaged with a 60× objective on a spinning disk confocal fluorescence microscope.
Gliding assay to determine correct polarity labeling
We use a simple gliding assay adapted from previous work to determine the polarity of our labeled microtubules.
Place two strips of the double-sided tape on a standard microscopy slide (the counter glass) about 5 mm apart. Press the tape down forcefully with forceps or the end of a microcentrifuge tube. Leave the protective backing.
Leave for at least 15 min at room temperature to allow the tape to fully adhere.
Pipette 10 μL of 2 mg/mL PLL-PEG between the two strips of tape and spread to all the edges using a pipette tip.
Allow PLL-PEG solution to dry.
Rinse thoroughly with MilliQ water and dry using pressurized air.
Using a diamond scribe pen, cut a #1.5 22 × 22 mm coverslip into four equal squares. Any debris from the cutting can be removed with pressurized air.
Remove the protective tape backing and place one of the coverslip pieces on top, pressing down firmly over the tape to ensure proper adhesion.
Use 50 μL of gliding buffer to wash the chamber, using filter paper to pull the solution through, and incubate with gliding buffer for 3 min at room temperature.
Dilute the motor protein, e.g., the minus-end-directed kinesin-14 HSET, to 100 nM in gliding buffer, and wash 50 μL through the chamber, followed by 3 min incubation with the motor dilution at room temperature.
Dilute the polarity-marked seeds 1:6 with room temperature gliding buffer and flow 50 μL into the chamber, again pulling through with the filter paper.
Seal the chamber ends with KoralisonTM paste.
Image fluorescent microtubules at the microscope, using a 5 s interval between frames to capture gliding events over the course of 20 min (Figure 2, Video. 1). Seeds should glide directionally according to the motor, e.g., the minus-end-directed HSET will drive microtubules with the plus/dim end leading. We found that 100% of motile microtubules with one dim and one bright segment glided in the appropriate direction. It can be difficult to achieve such a high percentage of correctly labeled polarity-marked microtubules using the NEM-tubulin method, which typically requires defining an acceptable threshold for successful labeling that is sufficiently high for the specific downstream assay.
Figure 2. AlexaFluor647-labeled, polarity-marked microtubules gliding on glass-adsorbed HSET motor proteins. As a minus-end-directed motor, HSET will drive microtubules according to their polarity with their plus-ends leading, indicating that the bright and dim segments are the minus- and plus-ends of the microtubule, respectively. Imaged with a 60× objective on a spinning disk confocal microscope. Arrows indicate the position of the bright end in the first frame. See also Video 1.
Video 1. Polarity-marked microtubules gliding on glass-adsorbed minus-end-directed HSET motor proteins. Microtubules attach to the surface over time and glide in the direction of their dim segments, i.e., with their plus-ends leading. Imaged with a 60× objective on a spinning disk confocal fluorescence microscope.
Data analysis
Data can be analyzed relatively simply by inspection of the fluorescent images and videos. Counting the number of appropriately segmented microtubules (one bright and one dim segment) vs. the overall number of microtubules gives a measure of efficiency. Proper gliding can be inferred by the directionality of the motor protein. In the case of surface-bound kinesin-14, a minus-end-directed motor, microtubules will be driven to glide by their plus-ends first, which should be the dim end according to this protocol.
Validation of protocol
This protocol has been used and validated in the following research article:
Henkin, Brito et al. [14]. The minus-end depolymerase KIF2A drives flux-like treadmilling of γTuRC-uncapped microtubules. J Cell Biol (Figure 1, panels C, D, F-K; Figure S2, panels B, C)
General notes and troubleshooting
General notes
In our hands, using the gliding assay, we found that 100% of microtubules with appropriate segmentation (one dim section, one bright section) were indeed correctly polarity labeled: n = 37 microtubules across two experiments, as we previously published [14]. As further validation, we showed that KIF2A, a microtubule depolymerase with a preference for minus-ends (independently confirmed by other groups [13,40]), depolymerized 98% of polarity-labeled microtubules predominantly from the bright minus-end (n = 271 microtubules across three different experimental sets) [14].
These seeds, if biotinylated, can be used in assays with microtubule-associated proteins, as in our previously published manuscript [14].
More details on the procedure for preparing these experiments using biotinylated, passivated glass and neutravidin links can be found in previous work [35].
Troubleshooting
Problem 1: Microtubule extensions do not grow during the DARPin (D1)2 step.
Possible cause: Tubulin concentration is too low; dynamic properties of tubulin may vary according to source, preparation methods, or means of storage.
Solution: Test in parallel a range of tubulin concentrations to grow the bright extensions. We tested a range between 0.2 and 1.5 μM to find a good condition that promotes relatively fast growth without nucleating too many new microtubules.
Possible cause: DARPin (D1)2 concentration too high; the protein acts by capping exposed β-tubulin, not only at the microtubule plus-end but also in solution, resulting in suppression of minus-end growth as well at high concentrations.
Solution: Test in parallel a range of DARPin (D1)2 concentrations. We tested a range from 50 to 150 nM to identify a condition that blocks plus-end but allows minus-end growth.
Problem 2: Microtubules extend from both directions during the DARPin (D1)2 step.
Possible cause: Tubulin concentration is too high.
Solution: Test in parallel a range of tubulin concentrations (see above).
Possible cause: DARPin (D1)2 concentration too low.
Solution: Test in parallel a range of DARPin (D1)2 concentrations (see above).
Problem 3: Microtubules show a repeated dim-bright-dim-bright patterning.
Possible cause: Microtubules can anneal end-to-end during storage at room temperature.
Solutions: Dilute to slow further annealing. Resuspend in the final step in 100 or 200 μL of warm BRB80 1× instead of 50 μL. Ensure that microtubules are prepared directly before experiments.
Problem 4: Microtubules do not glide in the gliding assay.
Possible cause: Motor protein not active in given concentrations.
Solutions: Try a different motor protein; try higher concentrations of motor protein; try lower concentrations of motor protein (in case of jamming).
Acknowledgments
We thank Raquel Garcia-Castellanos for technical support and Felix Ruhnow for microscopy support. We acknowledge support of the Spanish Ministry of Science and Innovation through the Centro de Excelencia Severo Ochoa (CEX2020‐001049‐S, MCIN/AEI/10.13039/501100011033), and the Generalitat de Catalunya through the CERCA programme. We are grateful to the CRG Core Technologies Programme for their support and assistance. T.S. acknowledges funding from the European Research Council (ERC) under the European Union's Horizon 2020 research and innovation programme (grant agreement No 951430) and from the Spanish Ministry of Science and Innovation (grants PID2019-108415GB-I00/AEI/10.13039/501100011033, and PID2022-142927NB-I00/AEI /10.13039/501100011033/FEDER, UE). C.B. was supported by EMBO long-term fellowship ALTF-883-2020 and Marie Curie fellowship TuRCReg.
Competing interests
The authors declare no competing interests.
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Plückthun, A. (2015). Designed ankyrin repeat proteins (DARPins): binding proteins for research, diagnostics, and therapy. Annu Rev Pharmacol Toxicol. 55: 489–511. https://doi.org/10.1146/annurev-pharmtox-010611-134654.
Campanacci, V., Urvoas, A., Consolati, T., Cantos-Fernandes, S., Aumont-Nicaise, M., Valerio-Lepiniec, M., Surrey, T., Minard, P. and Gigant, B. (2019). Selection and Characterization of Artificial Proteins Targeting the Tubulin α Subunit. Structure. 27(3): 497–506.e494. https://doi.org/10.1016/j.str.2018.12.001.
Consolati, T., Henkin, G., Roostalu, J. and Surrey, T. (2022). Real-Time Imaging of Single gammaTuRC-Mediated Microtubule Nucleation Events In Vitro by TIRF Microscopy. Methods Mol Biol. 2430: 315–336. https://doi.org/10.1007/978-1-0716-1983-4_21.
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Binz, H. K., Stumpp, M. T., Forrer, P., Amstutz, P. and Plückthun, A. (2003). Designing repeat proteins: well-expressed, soluble and stable proteins from combinatorial libraries of consensus ankyrin repeat proteins. J Mol Biol. 332(2): 489–503. https://doi.org/10.1016/s0022-2836(03)00896-9.
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Article Information
Publication history
Received: Jul 2, 2024
Accepted: Sep 13, 2024
Available online: Oct 17, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cell Biology > Cell structure > Microtubule
Biochemistry > Protein > Labeling
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5,110 | https://bio-protocol.org/en/bpdetail?id=5110&type=0 | # Bio-Protocol Content
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Fast and High-Resolution Imaging of Pollinated Stigmatic Cells by Tabletop Scanning Electron Microscopy
LR Lucie Riglet
IF Isabelle Fobis-Loisy
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5110 Views: 410
Reviewed by: Samik BhattacharyaPablo Bolanos-VillegasKristin L. Shingler
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Bio-protocol journal peer-reviewed
Nov 20, 2024 | This version
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Oct 14, 2021
Original Research Article:
The authors used this protocol in eLIFE Sep 2020
Abstract
In plants, the first interaction between the pollen grain and the epidermal cells of the stigma is crucial for successful reproduction. When the pollen is accepted, it germinates, producing a tube that transports the two sperm cells to the ovules for fertilization. Confocal microscopy has been used to characterize the behavior of stigmatic cells post-pollination [1], but it is time-consuming since it requires the development of a range of fluorescent marker lines. Here, we propose a quick, high-resolution imaging protocol using tabletop scanning electron microscopy. This technique does not require prior sample fixation or fluorescent marker lines. It effectively captures pollen grain behavior from early hydration (a few minutes after pollination) to pollen tube growth within the stigma (1 h after pollination) and is particularly efficient for tracking pollen tube paths.
Key features
• Analysis of the pollen behavior in stigmatic cells of Arabidopsis thaliana but can be broadly used for other species.
• Rapid and high-resolution imaging method.
• Allows testing pollen grain hydration states, pollen tube paths on stigmatic cells from various genetic backgrounds, and also pollen tube phenotypes.
Keywords: Scanning electron microscopy Stigmatic cell Pollen grain Pollen tube path Pollination Hydration state
Graphical overview
Background
In plants, successful fertilization hinges on the early communication between the male pollen grain and the stigmatic epidermis of the female organ. The upper part of the pistil ends with a specialized tissue, the stigma, composed of hundreds of unicellular elongated epidermal cells, called stigmatic cells or papillae [2]. When a pollen grain lands on a papilla, it adheres and undergoes hydration. Subsequently, it germinates, producing a pollen tube that grows at the stigma surface and through the transmitting tract of the style and the ovary to transport male gametes toward the ovules for fertilization. Confocal microscopy has been used to visualize the early step of pollen–stigma interaction and better characterize the behavior of both partners post-pollination [1]. However, this method is time-consuming because it requires the production of fluorescent marker lines specific for both pollination partners. Furthermore, the commonly used ubiquitous promoters, such as the CAMV35S or ubiquitin 10 promoters, typically employed to drive the expression of fluorescent proteins in plant tissues, exhibit low activity in papillae and pollen. Therefore, it is required to utilize specific promoters to conduct the expression of fusion proteins in transgenic stigma and pollen.
Here, we share a rapid, high-resolution imaging protocol using tabletop scanning electron microscopy, used in Riglet et al. [3], that does not require prior sample fixation or fluorescent marker lines. It enables effective visualization of pollen–stigma interaction and characterization of the pollen behavior, capturing events from early hydration of the pollen grain (a few minutes after pollination) to pollen tube growth within the stigma (1 h after pollination). This method proves particularly efficient for assessing hydration states and tracking pollen tube paths; it is particularly suitable to compare the pollen tube path on a wild-type stigma with mutated stigmatic cells but can also be used in reverse, comparing mutated pollen grains and tube paths on wild-type stigmatic cells.
Materials and reagents
Biological materials
Arabidopsis thaliana, ecotype Columbia Col-0 plants with flowers at stages 12–15 [4] in individual pots
Other materials
Straight fine tweezers (Dumont, model: Inox 8)
Double-sided black tape (Synergie4, model: DEN-77816)
Wood sticks and soft plant ties to stack the plants
Sewing thread to tag the emasculated pistils
Equipment
Tabletop scanning electron microscope (Hirox, model: SH-3000)
Stereomicroscope (Nikon, model: SMZ645, total magnification from 8× to 50×; eyepieces 10)
Software and datasets
ImageJ software (1.54c, 2024)]
Procedure
Plant preparation
A few Arabidopsis thaliana seeds, accession Col0, were germinated in 0.25 L plastic pots containing a peat-based compost with 60 kg/m3 of clay. After one week, we remove plantlets to keep one plant per pot to facilitate manipulation during emasculation and pollination. Plants are maintained in a growth chamber under long-day conditions (16 h light, 120–150 µmol m-2·s-1/8 h dark at 21 °C/19 °C) with a relative humidity of around 60%. The plants are watered throughout their growth cycle. To prevent pest attacks, it is important to avoid overwatering, especially when the plants are young. Fertilizer (N/P/K: 18/10/18) is added to the watering water (4 mL/L) once a week.
In our growth conditions, plants are ready for emasculation four weeks after germination; they have three or four robust stems. Avoid using old plants, which have more fragile stems and flowers and are more prone to dehydration during emasculation.
Water the plant thoroughly the day before emasculation to ensure turgid stigmas and pollen.
Flower emasculation should be done the day before pollination and imaging. Perform all processes at room temperature (around 22 °C).
Stigma preparation (emasculation procedure, the day before)
Flower emasculation should be done the day before pollination and imaging. See Video 1.
Position the plant in its culture pot under a stereomicroscope.
Bend the plant's inflorescences and gently clamp a flower bud at the end of stage 12 (Figure 1, Smyth et al. [4]) between the thumb and the forefinger to prevent bud movement during emasculation.
Video 1. Emasculation and pollination procedures
Figure 1. Arabidopsis thaliana developmental stages, focused on the stigma and pollen maturity. Inspired from Smyth et al. [4]. Stage duration is specified. A sepal from a stage-12 flower bud was removed to better visualize internal organs.
Using straight fine tweezers, remove flowers older than stage 12 [4] to minimize the risk of cross-pollination and younger buds to facilitate access to the stage-12 bud.
Using fine tweezers, carefully open the flower bud to expose the stigma (Figure 2A, B). Verify that anthers are not dehiscent, without any pollen grain released.
Note: Emasculate in the morning when the anthers are less dehiscent, typically when the growth chamber lights are just starting, to minimize the risk of pollen contamination from other stigmas.
Carefully remove the anthers (pale yellow, Figure 2C), petals, and sepals.
Note: Instead of completely removing organs, cut them with the fine tweezers (as indicated by arrows in Figure 2C) to retain part of the tissues (Figure 2D) and prevent excess dehydration of the bare pistil.
Ensure that no pollen grains remain on the stigma papillae (Figure 2E).
Caution: If any pollen grains are present, discard the bud and select another stage-12 bud.
Note: Limit emasculation to no more than five buds per plant to prevent pistil dehydration or damage to the emasculated buds while manipulating the plant.
Figure 2. Arabidopsis thaliana stigma at the end of stage 12 and emasculation prior to pollination. (A) Overview of the entire flower bud at the end of stage 12 with sepals and petals pushed back to better visualize the pistil. Orange arrows indicating the site for emasculation. (B) Close-up of the upper part of the flower. (C) Flower (manually opened) with orange arrows indicating the site for emasculation by anther removal. (D) Complete view of the emasculated flower. (E) Detailed close-up of the unpollinated stigma, free of pollen grains. Scale bars, 500 μm. (F) Staking of the plants using wood sticks and soft plant ties to support their stems.
Post-emasculation care
To label the emasculated pistil, a sewing thread can be attached at the base of the bare pistil.
Stake the plants (Figure 2F). We use wood sticks and soft plant ties to support their stems (see Graphical overview, step 1).
Caution: This step is important to prevent the bare pistils from getting in contact with pollen from other flower buds.
Return the plants to the culture room. If imaging stage-13 stigmas, wait approximately 18 h after emasculation (Figure 1; Smyth et al. [4]).
Note: Ensure the plant is well-watered to maintain turgescence in the stigma and pollen.
Pollination procedure (the day of imaging)
Under a stereomicroscope, hand-pollinate each emasculated flower with mature wild-type pollen grains (e.g., flowers at stage 13, Figure 1 and 3A).
To do so, collect a single dehiscent anther from a non-emasculated flower bud with tweezers and carefully brush the stigma with it. For easier quantification of pollen tube paths, deposit only a small amount of pollen grains (as shown in Figure 3B, C). To achieve this, gently brush the dehiscent anther on a tissue to remove excess pollen grains before applying it to the stigma.
Note: Collect a dehiscent anther from stage 13 to early stage 14 with pollen grains not too old to ensure the highest germination rate.
Note on pollen germination: Temperature and humidity influence pollen germination. Therefore, prepare and store samples in a room at a moderate temperature (around 20 °C) and ensure it is not too dry (at least 50% humidity). This environment helps maintain optimal conditions for pollen germination.
Figure 3. Pollination of Arabidopsis thaliana stigma at stage 13, stereomicroscope view. (A) Anther at maturity a few hours after dehiscence with yellow pollen grains. (B) Stigma shortly after pollination, displaying non-hydrated oval pollen grains (rugby balloon–like shape). (C) Stigma 30 min post-pollination, exhibiting well-hydrated roundish pollen grains (basket balloon–like shape). Scale bars, 100 μm.
Post-pollination handling: Leave the pollinated plants at room temperature on the bench. Depending on the focus of the imaging:
Wait 10–15 min to image pollen hydration.
Wait 30–60 min to image long pollen tubes.
Sample preparation
Cut the pollinated pistils transversally in the middle of the ovary using fine tweezers at the designated time post-pollination (see Figure 4).
Caution: Ensure that the sample height does not exceed 0.5 cm to fit into the microscope chamber.
Figure 4. Preparation of the sample before deposition on the scanning electron microscopy platform. (A–B) Position of the transversal cut of the pistil in the middle of the ovary.
Image setup procedure
Sample mounting: Gently deposit two or three cut pistils vertically onto a 4 × 2 cm double-sided tape positioned on the SEM platform (see Figure 5). Ensure that the pistils are not too close to each other, maintaining a minimum distance of 1 cm between both.
Figure 5. Scanning electron microscopy platform preparation. (A) Schematic view showing the method of mounting the stigmas on the SEM platform. (B) Platform displaying the cut pollinated pistils stuck on the double-sided tape before imaging. Scale bar, 1,000 μm. (C) Close-up view of two pistils mounted on the platform. Scale bar, 500 μm.
Scanning electron microscopy setup: Place the platform in the Hirox SEM SH-3000 (Figure 6), ensuring the door of the SEM is well sealed.
Note: The same protocol can be used with other tabletop SEMs.
Figure 6. Scanning electron microscopy Hirox SH-3000
Select temperature (-20 °C) and wait until that temperature is reached.
Select accelerating voltage: 15 kV.
Turn on the vacuum.
Imaging procedure
Observation: Observe samples within the first 15 min to prevent cells from collapsing during imaging.
Caution: Fast action is crucial as cells can collapse quickly within 15 min (see Figure 7A), making quantification of the pollen tube path impossible.
For a quick initial screening, use the AutoFocus button.
Tip: To obtain a sharp image of the stigma surface and adjust the focus, initially zoom in at high magnification on pollen grains located in the center of the stigma. Adjust the focus, contrast, and brightness. Then, zoom out to capture a broad view of the pistil.
Image capture: Acquire the images (Figure 7A–E). This method allows for imaging of the pollen hydration stage. Figure 7B displays non-hydrated pollen grains, while Figure 7C presents well-hydrated ones (Figure 7C). Pollen tube germination and tube paths can also be imaged and later quantified (Figure 7D–E) (See Data analysis section).
Figure 7. Visualization of pollinated stigmas at the scanning electron microscope. (A–C) Top view of a pollinated stigma observed at 300× magnification, showing (A) deflated stigmatic cells (collapsed), (B) fully turgid stigmatic cells with non-hydrated pollen grains (oval) a few minutes after pollination, and (C) stigmatic cells with well-hydrated pollen grains (round) and pollen tubes growing 30 min after pollination. (A–C) Scale bar, 50 μm. (D) Detail of (C) at 700× magnification. Scale bar, 50 μm. (E) Pollen–stigmatic cell interaction, viewed at 700× magnification. Scale bar, 20 μm.
This method can be applied to compare the pollen tube paths on wild-type and mutated stigmatic cells. Additionally, it can be used reversely, comparing wild-type and mutated pollen grain hydration and pollen tube paths on wild-type stigmatic cells. To compare the behavior associated with a wild type and a mutant, ensure that the stigmatic cells are pollinated at the same development stage and that pollination is done simultaneously. Ideally, mount a wild type and a mutant on the same platform to ensure imaging is performed under identical conditions.
Alternatively, single-stigma pollination can be conducted by applying two different pollen genotypes on each half of the stigma, as previously described [5].
Data analysis
Data processing and analysis: Quantification of pollen tube turn numbers using ImageJ software
Image processing: Open the images in ImageJ software and use the Cell Counter plugin to record the number of turns made by pollen tubes on the stigmatic epidermis.
Turn quantification: Count the number of turns made by the pollen tubes up to the base of the stigmatic cell.
Notes:
Select only papillae that received a single pollen grain to facilitate tracking of the pollen tube path. Indeed, if multiple pollen tubes are growing on the same papilla, their path may be affected by another one, deviating from the initial direction. This property, which we defined as self-avoidance, is illustrated in Figure 8A [6].
Select papillae for which the entire cell is visible from the top to the base.
Direction categorization: Use the Cell Counter tool in ImageJ to categorize the direction taken by the pollen tube. Navigate through Plugins → Analyze → Cell Counter. Classify the turns into several categories (Figure 8B):
Type 1: Straight—the pollen grain germinates on one side of the papilla and the tube stays on the same side.
Type 2: Half a loop—the pollen grain germinates on one side of the papilla and the tube grows toward the other side.
Type 3: One full loop—the pollen grain germinates on one side of the papilla and the tube returns to the same side before reaching the stigmatic base.
Type 4: Two loops.
Type 5: More than two loops.
An example of calculation and analysis is presented in Figure 8C, D.
Figure 8. Pollen tube paths on Arabidopsis thaliana stigmatic cells. (A) SEM images of a wild-type stigmatic cell pollinated by two wild-type pollen grains illustrating self-avoidance properties. Each pollen grain and tube have been colored to better visualize their paths. (B) Different configurations of pollen tube paths on Arabidopsis thaliana stigmas. (C) Example of the quantification of the number of loops made by the pollen tube on papillae at stage 13 (data from Riglet et al. [3]). Data are expressed as mean ± standard error of the mean (SEM). (D) Example of a histogram illustrating the quantification of the number of loops formed by the pollen tube on papillae at stage 13 (data from Riglet et al. [3]).
Statistical analysis
At least three independent experiments (different dates and plant batches) have to be performed, with at least 50 pollinated papillae analyzed in each experiment.
Data are expressed as mean ± SEM.
Use an adjusted chi-square test for homogeneity to analyze differences between two genotypes. Employ a chi-square test for independence to compare the same genotype at different stages.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Riglet et al. [3]. KATANIN-dependent mechanical properties of the stigmatic cell wall mediate the pollen tube path in Arabidopsis. eLife (Figure 2 panels A, B; Figure 4 panels B, C; Figure 5 panels B, D; Figure 5, supplement 1 panels A–D; Figure 5, supplement 2 panel C, Figure 5, supplement 3 panels A–G).
Acknowledgments
This protocol was originally described in and adapted from Riglet et al. [3]. This work was supported by a fellowship from the French Ministry of Higher Education and Research and by Grant ANR-14-CE11-0021.
Competing interests
The authors declare no competing interests.
References
Rozier, F., Riglet, L., Kodera, C., Bayle, V., Durand, E., Schnabel, J., Gaude, T. and Fobis-Loisy, I. (2020). Live-cell imaging of early events following pollen perception in self-incompatible Arabidopsis thaliana. J Exp Bot. 71(9): 2513–2526.
Heslop-Harrison, Y. and Shivanna, K. R. (1977). The Receptive Surface of the Angiosperm Stigma. Ann Bot. 41(6): 1233–1258.
Riglet, L., Rozier, F., Kodera, C., Bovio, S., Sechet, J., Fobis-Loisy, I. and Gaude, T. (2020). KATANIN-dependent mechanical properties of the stigmatic cell wall mediate the pollen tube path in Arabidopsis. eLife. 9: e57282.
Smyth, D. R., Bowman, J. L. and Meyerowitz, E. M. (1990). Early flower development in Arabidopsis. Plant Cell. 2(8): 755–767.
Nasrallah, M. E., Liu, P. and Nasrallah, J. (2002). Generation of Self-Incompatible Arabidopsis thaliana by Transfer of Two S Locus Genes from A. lyrata. Science. 297(5579): 247–249.
Riglet, L., Quilliet, C., Godin, C., John, K. and Fobis-Loisy, I. (2024). Geometry and cell wall mechanics guide early pollen tube growth in Arabidopsis thaliana. bioRxiv. doi.org/10.1101/2024.02.05.578915.
Article Information
Publication history
Received: May 16, 2024
Accepted: Sep 25, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Plant Science > Plant cell biology > Cell imaging
Cell Biology > Cell imaging > Electron microscopy
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Optimized Isolation of Lysosome-Related Organelles from Stationary Phase and Iron-Overloaded Chlamydomonas reinhardtii Cells
JL Jiling Li
HL Huan Long
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5111 Views: 179
Reviewed by: Ritu Gupta Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant Physiology Mar 2023
Abstract
Lysosome-related organelles (LROs) are a class of heterogeneous subcellular organelles conserved in eukaryotes, performing various functions. An important function of LROs is to mediate phosphorus and metal homeostasis. Chlamydomonas reinhardtii serves as a model organism for investigating metal ion metabolism. Considering that LROs contain polyphosphate and various metal elements, the purification strategy is based on their higher density by fractionating cell lysate through OptiPrep density gradient ultracentrifugation. Here, we optimized a method for purifying LROs from C. reinhardtii cells that have reached stationary phase (sta-LROs) or are overloaded with iron (Fe-LROs). Our protocol provides technical support for further investigations on the biogenesis and function of LROs in C. reinhardtii.
Key features
• This protocol purifies LROs from C. reinhardtii without disrupting the structure of chloroplasts.
• Following the purification of sta-LROs, these can be further fractionated into subgroups with distinct densities through the second iodixanol gradient.
• This protocol is applicable for the purification of LROs from the cell wall–deficient C. reinhardtii strain cw15.
Keywords: sta-LROs Fe-LROs C. reinhardtii OptiPrep density gradient Ultracentrifugation
Graphical overview
Background
Eukaryotic cells have evolved sophisticated mechanisms to counteract imbalances in environmental metal ions by sequestering excess metal ions (such as iron, copper, manganese, calcium, or zinc) within subcellular compartments to alleviate the toxicity of excessive metals [1–5]. These stored metal ions are subsequently utilized under conditions of deficiency to sustain intracellular metal ion homeostasis. Lysosome-related organelles (LROs) are a group of subcellular organelles involved in the storage of metal ions, including vacuoles in plants and yeast, acidocalcisomes in trypanosomes and algae, and other LROs [2,3,6].
Chlamydomonas reinhardtii is a unicellular green alga, serving as a eukaryotic model organism for studying trace element homeostasis, owing to its ability to grow under conditions of both excess and deficiency of metal elements [7,8]. Moreover, C. reinhardtii exhibits the capability to absorb and transform toxic metals from the environment [9–13], making it a primary producer model for investigating the mechanisms of toxic metal uptake and conversion. Docampo and colleagues isolated polyphosphate bodies from C. reinhardtii and reported that they were similar to acidocalcisomes in terms of their chemical composition and the presence of proton pumps [14]. When there is an excess of iron, copper, and manganese ions in the cultured medium, C. reinhardtii cells sequester the absorbed excess metal ions into LROs, maintaining the homeostasis of metal ions [1,4,5].
Our previous work has demonstrated that C. reinhardtii LROs exhibit heterogeneity in their morphology, elemental content, and protein composition. These LROs are crucial for the storage and homeostasis of trace metals, calcium, and phosphorus [15]. Sta-LROs, which are generated in cells during the stationary phase [15], resemble the protein lytic vacuoles in plants [16] and the digestive vacuoles of the malaria parasite [17]. Additionally, Fe-LROs, which arise in cells overloaded with iron, share similarities with acidocalcisomes [15]. Mg, Ca, Fe, Cu, Zn, and Mn are localized within polyphosphate granules in both sta-LROs and Fe-LROs, which are crucial organelles for the storage of phosphorus and metal ions within cells [15]. We have developed a method to quantify the storage capacity of LROs for phosphorus and metal ions, enabling the assessment of the sequestration of specific trace metals into the LROs under varying conditions [15]. Furthermore, 21 metal transporters have been identified in sta-LROs and Fe-LROs, suggesting their potential roles in mediating the transport of trace metal ions into or out of LROs [15].
Establishing a purification protocol for C. reinhardtii LROs is crucial for studying the formation and function of LROs, including their roles in metal ion metabolism and the transformation of environmentally toxic metals. Here, we introduce the purification steps for sta-LROs and Fe-LROs. The isolation principle relies on the high density of LROs, due to their rich content of phosphorus and metal ions. The cells are squeezed by using a syringe with a needle (diameter of 25G), which preserves the chloroplast structure; sta-LROs and Fe-LROs are then purified via OptiPrep density gradient ultracentrifugation. The limitation of this protocol is that it can only purify LROs from the cell wall–deficient C. reinhardtii strain, as the syringe compression method is insufficient to break wild-type C. reinhardtii cells. In future studies, it would be worth exploring the use of commercial Subtilisin (Alcalase) [18], to remove the cell wall of C. reinhardtii without disrupting other subcellular structures, followed by utilizing this protocol to purify LROs from wild-type C. reinhardtii cells.
Materials and reagents
Reagents
NH4Cl (Sinopharm Chemical Reagent, CAS number: 12125-02-9)
MgSO4·7H2O (Sinopharm Chemical Reagent, CAS number: 10034-99-8)
CaCl2·H2O (Sinopharm Chemical Reagent, CAS number: 10035-04-8)
Tris base (Genview, CAS number: BT350-500G)
KH2PO4 (Sinopharm Chemical Reagent, CAS number: 7778-77-0)
K2HPO4 (Sinopharm Chemical Reagent, CAS number: 7758-11-4)
Na2EDTA·2H2O (Sinopharm Chemical Reagent, CAS number: 6381-92-6)
(NH4)6Mo7O24 (Sinopharm Chemical Reagent, CAS number: 12054-85-2)
ZnSO4·7H2O (Sinopharm Chemical Reagent, CAS number: 7446-20-0)
H3BO3 (Sinopharm Chemical Reagent, CAS number: 10043-35-3)
MnCl2·4H2O (Sinopharm Chemical Reagent, CAS number: 13446-34-9)
CoCl2·6H2O (Sinopharm Chemical Reagent, CAS number: 7791-13-1)
CuSO4·5H2O (Sinopharm Chemical Reagent, CAS number: 7758-99-8)
FeSO4·7H2O (Sinopharm Chemical Reagent, CAS number: 7782-63-0)
KOH (Sinopharm Chemical Reagent, CAS number: 1310-58-3)
Glacial acetic acid (Sinopharm Chemical Reagent, CAS number: 64-19-7)
NH4Cl (MACKLIN, CAS number: 12125-02-9)
CaCl2·2H2O (MACKLIN, CAS number: 10035-04-8)
MgSO4·7H2O (MACKLIN, CAS number: 10034-99-8)
K2HPO4 (MACKLIN, CAS number: 7758-11-4)
KH2PO4 (MACKLIN, CAS number: 7778-77-0)
Acetic acid (MACKLIN, CAS number: 64-19-7)
EDTA Na2·2H2O (Genview, CAS number: 6381-92-6)
(NH4)6Mo7O24·4H2O (MACKLIN, CAS number: 12027-67-7)
ZnSO4·7H2O (MACKLIN, CAS number: 7446-20-0)
MnCl2·4H2O (MACKLIN, CAS number: 13446-34-9)
FeCl3·6H2O (MACKLIN, CAS number: 10025-77-1)
Na2CO3 (MACKLIN, CAS number: 497-19-8)
CuCl2·2H2O (MACKLIN, CAS number: 10125-13-0)
HCl (Sinopharm Chemical Reagent, CAS number: 7647-01-0)
EGTA (Genview, CAS number: 67-42-5)
KCl (Sinopharm Chemical Reagent, CAS number: 7447-40-7)
MgCl2 (Sinopharm Chemical Reagent, CAS number: 7786-30-3)
Sucrose (Biofroxx, CAS number: 57-50-1)
OptiPrep density gradient medium (Sigma-Aldrich, CAS number: 92339-11-2)
DTT (Blotopped, CAS number: 3483-12-3)
Protease inhibitor cocktail (Sigma, CAS number: P9599)
Solutions
TAP medium (see Recipes)
Revised TAP (see Recipes)
IB buffer (see Recipes)
OptiPrep gradient (see Recipes)
2 M DTT (see Recipes)
Recipes
Tris-acetate-phosphate (TAP) medium
Reagent Final concentration Quantity or Volume
TAP salts (100×) 1% (v/v) 10 mL
Tris base (100×) 1% (v/v) 10 mL
Phosphate buffer (1,000×) 0.1% (v/v) 1 mL
Hutner trace elements (1,000×) 0.1% (v/v) 1 mL
Glacial acetic acid 0.1% (v/v) 1 mL
Milli-Q Water Dilute with H2O to a final volume of 1 L
Total 1 L
Stocks for TAP medium:
TAP salts (100×)
Reagent Final concentration Quantity or Volume
NH4Cl 0.7 M 37.5 g
MgSO4·7H2O 40 mM 10 g
CaCl2·H2O 30 mM 5 g
Milli-Q H2O Dilute with H2O to a final volume of 1 L
Total 1 L
Tris base (100×)
Reagent Final concentration Quantity or Volume
Tris base 2 M 242 g
Milli-Q H2O Dilute with H2O to a final volume of 1 L
Total 1 L
Phosphate buffer (1,000×)
Reagent Final concentration Quantity or Volume
KH2PO4 0.4 M 27 g
K2HPO4 0.62 M 54 g
Milli-Q H2O Dilute with H2O to a final volume of 500 mL
Total 500 mL
Hutner trace elements
Note: Store at 4 °C.
Reagent Quantity or Volume
Na2EDTA·2H2O 55.36 g
(NH4)6Mo7O24·4H2O 1.108 g
ZnSO4·7H2O 22 g
H3BO3 11.4 g
MnCl2·4H2O 5.06 g
CoCl2·6H2O 1.61 g
CuSO4·5H2O 1.57 g
FeSO4·7H2O 4.99 g
Milli-Q H2O Dilute with H2O to a final volume of 1 L
Total 1 L
Dissolve 55.36 g Na2EDTA·2H2O in 250 mL of H2O and gradually add KOH with heating to accelerate the dissolution. Dissolve 1.108 g of (NH4)6Mo7O24·4H2O in 50 mL of H2O. Dissolve 22 g of ZnSO4·7H2O in 100 mL of H2O. Dissolve 11.4 g of H3BO3 in 200 mL of H2O with heating. Dissolve 5.06 g of MnCl2·4H2O in 50 mL of H2O. Dissolve 1.61 g of CoCl2·6H2O in 50 mL of H2O. Dissolve 1.57 g of CuSO4·5H2O in 50 mL of H2O. Dissolve 4.99 g of FeSO4·7H2O in 50 mL of H2O before mixing to avoid oxidation.
Once all the solutions mentioned above are prepared, mix them, excluding Na2EDTA. Boil the mixture, then add the Na2EDTA solution. The mixture will turn green. Ensure all reagents are completely dissolved, then cool the solution to 70 °C and maintain this temperature. Adjust the pH to 6.7 using KOH (calibrate the pH meter at 70 °C; NaOH is unsuitable for this purpose). After adjusting the pH, dilute the solution to a final volume of 1 L, seal the conical flask with cotton, and let it stand for 1–2 weeks, shaking daily. The solution will eventually turn purple and form a rusty brown precipitate. Filter out the precipitate and store the resulting solution at 4 °C for future use.
Revised TAP medium
Note: Reagents used in the revised TAP medium must be trace element grade. Reagents purchased from MACKLIN are only used in revised TAP.
Reagent Final concentration Quantity or Volume
Beijerinck’s solution 1% (v/v) 10 mL
Phosphate solution 0.833% (v/v) 8.33 mL
Tris-Acetate stock solution 1% (v/v) 10 mL
Na2EDTA stock solution 25 µM 1 mL
(NH4)6Mo7O24 stock solution 28.5 nM 1 mL
Na2SeO3 stock solution 0.1 µM 1 mL
ZnEDTA stock solution 2.5 µM 1 mL
MnEDTA stock solution 6 µM 1 mL
FeEDTA stock solution 20 µM 1 mL
CuEDTA stock solution 2 µM 1 mL
Milli-Q Water Dilute with H2O to a final volume of 1 L
Total 1 L
For iron-limited TAP medium, add 0.01 mL of FeEDTA stock solution. For iron-overloading TAP medium, add 10 mL FeEDTA stock solution.
Stocks for revised TAP medium:
Beijerinck’s solutions (100×)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
NH4Cl 750 mM 40 g
CaCl2·2H2O 34 mM 5 g
MgSO4·7H2O 40 mM 10 g
Milli-Q Water Dilute with H2O to a final volume of 1 L
Total 1 L
Phosphate solution (120×)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
K2HPO4 82 mM 14.34 g
KH2PO4 54 mM 7.26 g
Milli-Q Water Dilute with H2O to a final volume of 1 L
Total 1 L
Tris-Acetate stock solution (100×)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
Tris base 2 M 242 g
Acetic acid 1.7 M 100 mL
Milli-Q Water Dilute with H2O to a final volume of 1 L
Total 1 L
Na2EDTA concentrate (Pre 1)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
Na2EDTA·2H2O 125 mM 13.959 g
Milli-Q Water Dilute with H2O to a final volume of 300 mL
Total 300 mL
Dissolve 13.959 g of Na2EDTA·2H2O in about 250 mL of H2O and titrate to pH 8.0 with trace element grade KOH (about 1.7 g).
(NH4)6Mo7O24 concentrate (Pre 2)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
(NH4)6Mo7O24·4H2O 285 µM 0.088 g
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
Na2SeO3 concentrate (Pre 3)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
Na2SeO3 1 mM 0.043 g
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
Individual stock solutions (1,000×) of trace elements
Note: Store at 4 °C.
i. Na2EDTA stock solution
Reagent Final concentration Quantity or Volume
Na2EDTA 25 mM 50 mL of Pre 1
Milli-Q Water 200 mL
Total 250 mL
ii. (NH4)6Mo7O24 stock solution
Reagent Final concentration Quantity or Volume
(NH4)6Mo7O24 28.5 µM 25 mL of Pre 2
Milli-Q Water 225 mL
Total 250 mL
iii. Na2SeO3 stock solution
Reagent Final concentration Quantity or Volume
Na2SeO3 0.1 mM 25 mL of Pre 3
Milli-Q Water 225 mL
Total 250 mL
iv. ZnEDTA stock solution
Reagent Final concentration Quantity or Volume
ZnSO4·7H2O 2.5 mM 0.18 g
Na2EDTA 2.75 mM 5.5 mL of Pre1
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
v. MnEDTA stock solution
Reagent Final concentration Quantity or Volume
MnCl2·4H2O 6 mM 0.297 g
Na2EDTA 6 mM 12 mL of Pre1
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
vi. FeEDTA stock solution
Reagent Final concentration Quantity or Volume
FeCl3·6H2O 20 mM 1.35 g
Na2EDTA 22 mM 2.05 g
Na2CO3 22 mM 0.58 g
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
Note: Mix Na2EDTA with Na2CO3 in Milli-Q water. Add FeCl3·6H2O after the first two components are dissolved. Do not use Pre 1.
vii. CuEDTA stock solution
Reagent Final concentration Quantity or Volume
CuCl2·2H2O 2 mM 0.085 g
Na2EDTA 2 mM 4 mL of Pre1
Milli-Q Water Dilute with H2O to a final volume of 250 mL
Total 250 mL
Isolation buffer (IB buffer)
Note: Store at 4 °C.
Reagent Final concentration Quantity or Volume
1 M Tris-HCl 20 mM 4 mL
0.5 M EGTA 5 mM 2 mL
0.5 M KCl 5 mM 2 mL
0.2 M MgCl2 2 mM 2 mL
Sucrose 6% (W/V) 12 g
Milli-Q Water Dilute with H2O to a final volume of 200 mL
Total 200 mL
Stocks for IB buffer:
1 M Tris-Cl
Reagent Final concentration Quantity or Volume
Tris base 1 M 12.114 g
Milli-Q Water Dilute with H2O to a final volume of 100 mL
Total 100 mL
Dissolve 12.114 g of Tris base in 50 mL of H2O, adjust the pH to 7.6 with HCl, and adjust to 100 mL.
0.5 M EGTA
Reagent Final concentration Quantity or Volume
EGTA 0.5 M 19 g
Milli-Q Water Dilute with H2O to a final volume of 100 mL
Total 100 mL
0.5 M KCl
Reagent Final concentration Quantity or Volume
KCl 0.5 M 3.725 g
Milli-Q Water Dilute with H2O to a final volume of 100 mL
Total 100 mL
0.2 M MgCl2
Reagent Final concentration Quantity or Volume
MgCl2 0.2 M 1.9 g
Milli-Q Water Dilute with H2O to a final volume of 100 mL
Total 100 mL
OptiPrep gradient
Note: OptiPrep density gradient medium needs to be dried into iodixanol powder at 50 °C before use. Store at 4 °C.
Reagent Final concentration Quantity or Volume
OptiPrep density gradient 80% (w/v) 3.2 g of iodixanol powder in 4 mL of IB buffer
60% (w/v) 2.4 g of iodixanol powder in 4 mL of IB buffer
40% (w/v) 1.6 g of iodixanol powder in 4 mL of IB buffer
37% (w/v) 1.48 g of iodixanol powder in 4 mL of IB buffer
34% (w/v) 1.36 g of iodixanol powder in 4 mL of IB buffer
27% (w/v) 1.08 g of iodixanol powder in 4 mL of IB buffer
24% (w/v) 0.96 g of iodixanol powder in 4 mL of IB buffer
15% (w/v) 0.6 g of iodixanol powder in 4 mL of IB buffer
2 M DTT
Note: Store at -20 °C.
Reagent Final concentration Quantity or Volume
DTT 2 M 0.3085 g
Milli-Q Water Dilute with H2O to a final volume of 1 mL
Total 1 mL
Laboratory supplies
Blue cap bottles, 1 L (Beyotime, catalog number: FBT008), 500 mL (Beyotime, catalog number: FBT006), 250 mL (Beyotime, catalog number: FBT002)
1 L beaker, 500 mL beaker, 250 mL beaker
500 mL glass conical flask
Volumetric flasks of different volumes
50 mL centrifuge tube
BD Precision Glide needle 25G×5/8 (Becton Dickinson Medical, catalog number: 301805)
Open-top thin wall ultra-clear tube, 5 mL (Beckman Coulter, catalog number: 344057)
1.5 mL Eppendorf tubes
Equipment
Avanti® J-E centrifuge (Beckman Coulter, model: 369001)
96 mm diameter polypropylene conical bottle adapter (Beckman Coulter, model: 392078)
JS-5.3 AllSpin swinging-bucket rotor and buckets (Beckman Coulter, model: 368690)
JA-20 fixed-angle aluminum rotor (Beckman Coulter, model: 334831)
Desktop micro speed freezing centrifuge Optima MAX-XP (Beckman Coulter, model: 393315)
MLS-50 swinging-bucket rotor (Beckman Coulter, model: 367280)
Microcentrifuge (Eppendorf, model: 5424R)
Procedure
Cell cultures
Inoculate cell wall–deficient strain cw15 cells grown on a TAP plate into two flasks containing 300 mL of TAP medium. Culture the cells at 24 °C with shaking (180 rpm) under continuous illumination (25 E·m-2·s-1) for 6 days. These cells can then be used to purify sta-LROs.
For the iron overloading experiment, inoculate 5–6 mL of 2-day-old cw15 cells, cultured in revised TAP medium, into two flasks containing 300 mL of iron-limited TAP medium (0.2 µM Fe). Culture the cells under continuous light for 5 days with shaking (180 rpm) until the cell density reaches 3–5 × 106 cells mL−1. Subsequently, culture cells for another 26–28 h in iron-overloading TAP medium (200 µM Fe). These cells can then be used to isolate Fe-LROs.
Note: It is recommended to purify LROs from freshly cultured cells. Storing cell pellets at -80 °C is not advised, as freeze-thaw cycles can cause partial cell lysis and chloroplast disruption, leading to impure LROs extraction.
Cell disruption
Collect the cells from each flask (600 mL in total) by centrifugation at 3,500× g for 3 min.
Discard the supernatant and wash the pellets once with 100 mL of IB buffer by resuspending with a pipette.
Discard the supernatant again and resuspend the total pellets in 15 mL of IB buffer supplemented with a complete protease inhibitor cocktail and 2 mM DTT.
Disrupt the cells by squeezing them through a syringe with a 25 G needle three times, ensuring most cells are opened without disrupting the chloroplast structure. During the squeeze process, ensure that the cell suspension flows out of the syringe needle in a continuous stream rather than intermittently dripping.
First OptiPrep gradient ultracentrifugation
Adjust the volume of each sample to 70 mL with IB buffer.
Centrifuge the samples at 10,000× g for 10 min at 4 °C.
Discard the supernatant and resuspend the pellets in 4 mL of 34% OptiPrep (add 8 μL of DTT and 40 μL of protein inhibitor cocktail before use).
Prepare a discontinuous OptiPrep gradient with 1 mL each of 15%, 24%, 34% (with samples), 37%, and 40% OptiPrep. It is recommended to prepare the OptiPrep gradient solution fresh for each use. The gradient should be constructed from the bottom up, starting with the highest density gradient at the bottom of the ultracentrifuge tube. Using a 1 mL pipette, carefully layer each subsequent gradient along the inner wall of the centrifuge tube at the slowest possible speed. During the gradient preparation process, distinct boundaries between different density layers can be clearly observed when held up to the light. There is no need for pause time between the different density layers.
Centrifuge the gradient at 50,000× g in a Beckman MLS-50 rotor for 60 min at 4 °C, setting the acceleration to the maximum level and the deceleration to coast. The pellet below the 40% layer (P1) is the purified total LROs. Purified LROs can be stored at -80 °C for subsequent protein identification and related experiments, with a recommended storage duration of no more than three months. For localization experiments such as staining or immunofluorescence, it is recommended to use the purified LROs immediately after preparation.
Note: The above steps are applicable for the purification of both sta-LROs and Fe-LROs from cw15 cells.
Second OptiPrep gradient ultracentrifugation
Discard the supernatant and resuspend the total sta-LROs with 2 mL of 40% OptiPrep (add 4 μL of DTT and 20 μL of protein inhibitor cocktail before use).
Prepare a discontinuous OptiPrep gradient with 1 mL each of IB buffer, 27%, 40% (with samples), 60%, and 80% OptiPrep.
Centrifuge the gradient at 50,000× g in a Beckman MLS-50 rotor for 60 min at 4 °C, setting the acceleration to the maximum level and the deceleration to coast.
Collect each layer (L1: layer between 40% and 60%, L2: layer between 60% and 80%) and the pellet below 80% (P2) in 1.5 mL Eppendorf tubes, fill to 1.5 mL with IB buffer, and mix by repeated pipetting.
Centrifuge at 15,000× g for 10 min at 4 °C.
Discard the supernatant; pellets are subgroups of sta-LROs separated by different densities.
Note: The second OptiPrep gradient ultracentrifugation is used to separate total sta-LROs into subgroups, as most Fe-LROs accumulate in the pellet below 80% OptiPrep due to their high density.
Validation of protocol
This protocol has been used and validated in the following research article:
Long et al. [15]. Structural and functional regulation of Chlamydomonas lysosome-related organelles during environmental changes. Plant Physiology. DOI: 10.1093/plphys/kiad189
General notes and troubleshooting
General notes
Set the acceleration to the maximum level and the deceleration to coast during ultracentrifugation.
Vacuum must be applied prior to ultracentrifugation.
Acknowledgments
This study was supported by the National Key R&D Program of China (2020YFA0907400). This protocol was described and validated in the following research article: Long et al. [15]. Structural and functional regulation of Chlamydomonas lysosome-related organelles during environmental changes. Plant Physiology. DOI: 10.1093/plphys/kiad189.
Competing interests
The authors declare that the research was conducted without any commercial or financial relationships that could be construed as potential conflicts of interest.
References
Hong-Hermesdorf, A., Miethke, M., Gallaher, S. D., Kropat, J., Dodani, S. C., Chan, J., Barupala, D., Domaille, D. W., Shirasaki, D. I. and Loo, J. A. (2014). Subcellular metal imaging identifies dynamic sites of Cu accumulation in Chlamydomonas. Nat Chem Biol. 10(12): 1034–1042.
Docampo, R. and Huang, G. (2016). Acidocalcisomes of eukaryotes. Curr Opin Cell Biol. 4166–72.
Docampo, R., (2024). Advances in the cellular biology, biochemistry, and molecular biology of acidocalcisomes. Microbiol Mol Biol Rev. 88(1): e00042–23.
Tsednee, M., Castruita, M., Salomé, P. A., Sharma, A., Lewis, B. E., Schmollinger, S. R., Strenkert, D., Holbrook, K., Otegui, M. S. and Khatua, K. (2019). Manganese co-localizes with calcium and phosphorus in Chlamydomonas acidocalcisomes and is mobilized in manganese-deficient conditions. J Biol Chem. 294(46): 17626–17641.
Schmollinger, S., Chen, S., Strenkert, D., Hui, C., Ralle, M. and Merchant, S. S. (2021). Single-cell visualization and quantification of trace metals in Chlamydomonas lysosome-related organelles. Proc Natl Acad Sci USA. 118(16): e2026811118.
Blaby-Haas, C. E. and Merchant, S. S. (2014). Lysosome-related Organelles as Mediators of Metal Homeostasis. J Biol Chem. 289(41): 28129–28136.
Blaby-Haas, C. E. and Merchant, S. S. (2017). Regulating cellular trace metal economy in algae. Curr Opin Plant Biol. 3988–96.
Salomé, P. A. and Merchant, S. S. (2019). A Series of Fortunate Events: Introducing Chlamydomonas as a Reference Organism. Plant Cell. 31(8): 1682–1707.
Hanikenne, M., (2003). Chlamydomonas reinhardtii as a eukaryotic photosynthetic model for studies of heavy metal homeostasis and tolerance. New Phytol. 159(2): 331–340.
Beauvais-Flück, R., Slaveykova, V. I. and Cosio, C. (2017). Cellular toxicity pathways of inorganic and methyl mercury in the green microalga Chlamydomonas reinhardtii. Sci Rep. 7(1): 8034.
Samadani, M., Perreault, F., Oukarroum, A. and Dewez, D. (2018). Effect of cadmium accumulation on green algae Chlamydomonas reinhardtii and acid-tolerant Chlamydomonas CPCC 121. Chemosphere. 191174–182.
Samadani, M. and Dewez, D. (2018). Cadmium accumulation and toxicity affect the extracytoplasmic polyphosphate level in Chlamydomonas reinhardtii. Ecotoxicol Environ Saf. 166200–206.
Thiriet-Rupert, S., Gain, G., Jadoul, A., Vigneron, A., Bosman, B., Carnol, M., Motte, P., Cardol, P., Nouet, C. and Hanikenne, M. (2021). Long-term acclimation to cadmium exposure reveals extensive phenotypic plasticity in Chlamydomonas. Plant Physiol. 187(3): 1653–1678.
Ruiz, F. A., Marchesini, N., Seufferheld, M. and Govindjee. and Docampo, R. (2001). The Polyphosphate Bodies of Chlamydomonas reinhardtii Possess a Proton-pumping Pyrophosphatase and Are Similar to Acidocalcisomes. J Biol Chem. 276(49): 46196–46203.
Long, H., Fang, J., Ye, L., Zhang, B., Hui, C., Deng, X., Merchant, S. S. and Huang, K. (2023). Structural and functional regulation of Chlamydomonas lysosome-related organelles during environmental changes. Plant Physiol. 192(2): 927–944.
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Article Information
Publication history
Received: Jul 9, 2024
Accepted: Sep 13, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Plant Science > Plant cell biology > Organelle isolation
Cell Biology > Organelle isolation > Lysosome
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Peer-reviewed
Small-Molecule Probe for Imaging Oxidative Stress–Induced Carbonylation in Live Cells
OD Ozlem Dilek
DT Dilek Telci
HE Hazel Erkan-Candag
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5112 Views: 396
Reviewed by: Komuraiah MyakalaThomas Linto Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Scientific Reports May 2020
Abstract
Protein carbonylation has been known as the major form of irreversible protein modifications and is also widely used as an indicator of oxidative stress in the biological environment. In the presence of oxidative stress, biological systems tend to produce large amounts of carbonyl moieties; these carbonyl groups do not have particular UV-Vis and fluorescence spectroscopic characteristics that we can differentiate, observe, and detect. Thus, their detection and quantification can only be performed using specific chemical probes. Commercially available fluorescent probes to detect specific carbonylation in biological systems have been used, but their chemical portfolio is still very limited. This protocol outlines the methods and procedures employed to synthesize a probe, (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol (2Hzin5NP), and assess its impact on carbonylation in human cells. The synthesis involves several steps, including the preparation of its hydrazone compounds mimicking cell carbonyls, 2-Hydrazinyl 5-nitrophenol, (E,Z)-2-(2-ethylidenehydrazonyl)-5-nitrophenol, and the final product (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol. The evaluation of fluorescence quantum yield and subsequent cell culture experiments are detailed for the investigation of 2Hzin5NP effects on cell proliferation and carbonylation.
Key features
• This protocol builds upon probe development using click chemistry method by Dilek et al. [1], and its biolabeling application in renal cancer cell lines.
• The non-fluorescent probe has a fast reaction with carbonyl moieties at neutral pH to form a stable fluorescent product leading to a spectroscopic alteration.
• Microscopic and fluorometric analyses can distinguish the exogenous and endogenous ROS-induced carbonylation profile in human dermal fibroblasts along with renal cell carcinoma.
• Carbonylation level that differs in response to exogenous and endogenous stress in healthy and cancer cells can be detected by the newly synthesized fluorescent probe.
Keywords: Bioorthogonal chemistry Click chemistry Fluorescent probes Fluorescence imaging Diagnostics Cancer cells
Graphical overview
Background
Fluorescent probes play a crucial role in visualizing cellular events; the development of probes for imaging oxidative stress-induced carbonylation in live cells represents a significant area of research [1–4]. Oxidative stress–induced protein carbonylation is a key marker of cellular damage and is implicated in various pathological conditions, including neurodegenerative diseases and cancer [5–7]. This area of study aims to enhance our understanding of cellular responses to oxidative stress [8] and provides potential targets for therapeutic interventions [9,10]. Several methodologies have been employed to study protein carbonylation, including immunoblotting and mass spectrometry [11]. However, these techniques often lack spatiotemporal resolution and may require cell fixation, preventing real-time observation. Fluorescent probes, therefore, offer the advantage of live-cell imaging, allowing dynamic monitoring of carbonylation processes [12,13]. To monitor carbonylation in live-cell imaging, the click chemistry method [14,15] is often used as a fast conjugation method, which can be characterized by high efficiency, selectivity, and compatibility with biological systems [16], exemplified by the hydrazine-hydrazone chemistry, where the reactive hydrazine group reacts with an aldehyde or ketone moiety to form a stable hydrazone linkage [17]. This chemistry has been widely employed in the development of fluorescent probes, such as utilizing a fluorescent dye conjugated with an aldehyde group that selectively reacts with a hydrazine-functionalized biomolecule, enabling precise visualization and tracking within biological systems. Our protocol employs a fluorescent probe, (E,Z)-2-(2-(2-hydroxybenzylidene)hydrazonyl)-5-nitrophenol (2Hzin5NP), facilitating real-time observation of oxidative stress–induced carbonylation in live cells. The protocol includes a WST-1 cell proliferation assay and quantification of carbonylation in cell lysate, enabling quantitative assessment of cellular responses. One particular limitation of using this probe for carbonylation imaging is its specificity for responding to an oxidative stress environment; this should be validated against other cellular processes. The protocol involves detecting carbonylation levels in renal carcinoma cell lines A-498 and ACHN, as well as in human dermal fibroblasts (HDF), using these as cell models [18]. However, its applicability to other cell types should also be investigated. Beyond imaging oxidative stress–induced carbonylation, this protocol may find applications in drug discovery and cellular signaling such as assessing the impact of potential therapeutics on oxidative stress–induced cellular damage and investigating the role of carbonylation in cellular signaling pathways. In conclusion, the presented protocol offers a valuable tool for researchers in the field of oxidative stress, enabling live-cell imaging of carbonylation with potential applications in various scientific domains. The stepwise synthesis and quantitative analysis contribute to the protocol's robustness, though ongoing research will refine its specificity and broaden its applicability.
Materials and reagents
Cell lines
A-498, primary human kidney epithelial carcinoma, adherent (ATCC, catalog number: Htb-44)
ACHN, metastatic renal cell adenocarcinoma, adherent (ATCC, catalog number: Crl-1611)
HDF, human dermal fibroblast, adherent (ATCC, catalog number: PCS 201-012)
Chemicals
Chemical synthesis
Silica gel (silica gel 60-200 mesh) (2.5 kg) (Merck, catalog number: 107734)
Hexane (2.5 L) (Merck, catalog number: 104368)
Ethanol (EtOH) (100%) (Merck, catalog number: M.100986.2500)
Chloroform (Merck, catalog number: SC.CL.0200.2500)
Dichloromethane (2.5 L) (Merck, catalog number: M.106050.2500)
Acetone (2.5 L) (Merck, catalog number: M.100013.2500)
Hydrochloric acid (HCl) (2.5 L) (Merck, catalog number: 100317)
Sodium hydroxide pellets (Merck, catalog number: 106498)
Sodium chloride (VWR, catalog number: SC.SO.0227.1000)
Methanol (spectral grade, anhydrous) (2.5 L) (Merck, catalog number: 106009)
Dioxane (1 L) (Spectrophotometric, catalog number: 154822)
Methanol (normal, 2.5 L) (Sigma-Aldrich, catalog number: 34885)
Sodium nitrite (1 kg) (Merck, catalog number: 106544)
Tin (II) chloride (Stannous chloride) (100 g) (Fluka, catalog number: 31669)
2-Amino-5-Nitrophenol (100 g) (Sigma, catalog number: 303585)
Diethyl ether (1 L) (Merck, catalog number: 100921)
TLC aluminum sheets (Merck, catalog number: 105554)
Cell culture media
Dulbecco’s modified Eagle’s medium (DMEM), high glucose (Gibco, catalog number: 41966)
Fetal bovine serum (FBS), cell culture tested (Gibco, catalog number: 10082)
Other reagents for cell culture
Bovine serum albumin (BSA), protein standard (Sigma, catalog number: P0834)
Dimethyl sulfoxide (DMSO) (Santa Cruz, catalog number: Sc-202581)
Dulbecco’s phosphate buffered saline (PBS) (Pan Biotech, catalog number: P04-53500)
H2O2 (50 wt % in H2O) (Sigma, catalog number: 519813)
L-Glutamine (Invitrogen, catalog number: 25030)
Methanol 99% (Sigma, catalog number: 34885)
Phenylmethanesulfonylfluoride (PMSF) (Sigma, catalog number: 78830)
Protease inhibitor (Pi) (Sigma, catalog number: P8340)
Penicillin-streptomycin (Thermo Scientific, catalog number: Sv30010 or Biochrom, catalog number: A2213)
Trypsin-EDTA (Biochrom, catalog number: L2153)
Kits
Cell Proliferation Reagent WST-1 (Roche, catalog number: 05015944001)
Protein Assay Reagent A (Bio-Rad, catalog number: 5000113)
Protein Assay Reagent B (Bio-Rad, catalog number: 5000114)
DCFDA, Cellular Reactive Oxygen Species Detection Assay Kit (Abcam, catalog number: Ab113851)
Small equipment and other supplies
Bright-LineTM hemocytometer (Sigma-Aldrich, catalog number: Z359629)
Coverslip (Sigma-Aldrich, catalog number: Z375357)
Electronic pipette (CAPP Aid)
Filter 0.22 mm (TPP), 0.45 mm (Santorium Stedim Biotech)
Graduated cylinder 50, 250, 500, 1,000 mL (Isolab)
Micropipettes 10, 20, 100, 200, 1,000 μL (Eppendorf Research)
Pipette tips 10, 100, 200, 1,000 µL (Capp Expell Plus)
Polypropylene centrifuge tubes 0.5, 1.5, 2, 15, 50 mL (Isolab)
Serological pipettes 2, 5, 10, 25 mL (Grenier Bio or Axygen)
Tissue culture flasks, T-25, T-75, T-150 (TPP or Grenier-Bio)
Multiple-well cell culture plates and cryovials (TPP or Grenier-Bio)
Whatman paper (Isolab)
Equipment
-80 °C freezer (Thermo, model: Forma -86 C ULT Freezer)
Bruker Avance III 500 MHz Spectrometry
Centrifuge (Hettich, model: Mikro 22r and Sigma, model: 2-5 Centrifuge)
CO2 Incubator (Nuaire, model: Nu5510/E/G)
Confocal microscope (Zeiss, model: Lsm 800)
Fluorescence microscope (Nikon, model: 80i Eclipse Fluorescence Microscope)
Fume Hood (Greenlab)
Heater (Bioer, model: Mb102)
Laminar flow cabinet (ESCO Lab culture Class II Biohazard Safety Cabinet 2A)
Light microscope (Nikon, model: Eclipse #Ts100)
Magnetic stirrer (Heidolph, model: Mr 3004)
pH meter (Hanna Instruments, model: Ph211)
Rotary evaporator (Heidolph, model: Hei-VAP Silver Packages)
UV lamb, cabinet (CAMAG)
Varioskan Lux multimode microplate reader (Thermo Fisher)
Vortex (Stuart Sa8p)
Water bath (Stuart, model: Sb540)
Software and datasets
GraphPad Prism 6
Free versions of SigmaPlot Software and ChemDraw
Procedure
Organic syntheses
Synthesis of 2-hydrazinyl-5-nitrophenol HCl salt
In a fume hood, prepare a cold solution of sodium nitrite (106 mg, 1.5 mmol) by dissolving it in 385 μL of water. Keep this solution cold by storing it in an ice container. In a separate container, prepare a cold solution of 2-Amino-5-nitrophenol (200 mg, 1.3 mmol) in 648 μL of HCl and keep it on ice as well. Slowly add the cold sodium nitrite solution dropwise to the cold 2-Amino-5-nitrophenol solution while maintaining the mixture on ice. Make sure to perform this step under a chemical fume hood to safely handle the hazardous chemicals.
Stir the mixture for 1 h at -5 °C, keeping it in a container filled with ice. Add sodium chloride to the ice to maintain the temperature at -5 °C or lower. Meanwhile, prepare a solution of stannous chloride (931 mg, 4.1 mmol) by dissolving it in 927 μL of cold HCl. Slowly add this stannous chloride solution to the reaction mixture while maintaining the low temperature.
Mix the reaction for 1 h at -5 °C.
Vacuum filter the mixture and wash the precipitate with cold methanol and diethyl ether. The reaction should yield dried 2-Hydrazine-5-nitrophenol HCl salt (177 mg) with a yield of 66%.
Synthesis of 2-salicylaldehyde-5-nitrophenol
Stir 2Hzin5NP (400 mg, 2.36 mmol) and salicylaldehyde (2.5 mL, 23.6 mmol) in 19 mL of MeOH for 1 h at room temperature.
After stirring, cool the reaction mixture in an ice bath and then filter the mixture.
Wash the solid product with cold MeOH and dry it under vacuum. This should yield 155 mg of fluorescent product with a yield of 24%. The product exhibits an Rf value of 0.39 on TLC plates using a mixture of hexane and ethyl acetate (7:4) as eluents.
Synthesis of 2-acetaldehyde-5-nitrophenol
Stir 2Hzin5NP (90 mg, 0.53 mmol) and acetaldehyde (29 μL, 0.44 mmol) in 2.4 mL of EtOH for 1 h at room temperature.
After stirring, cool the reaction mixture in an ice bath, then filter the mixture.
Wash the solid product with cold EtOH and dry it under vacuum. This should yield 90 mg of fluorescent product with a yield of 87%.
Protocol for Rf calculation using thin layer chromatography (TLC)
Materials needed:
TLC plate (silica gel or another appropriate stationary phase)
Solvent (mobile phase): Prepare 7:4 hexane/ethyl acetate solvent
Sample(s) to analyze
Developing chamber (e.g., beaker with lid)
Pencil
Ruler
UV lamp or staining reagent (if needed)
Tweezers
Procedure
Prepare the TLC plate
Use a pencil to draw a baseline approximately 1 cm from the bottom edge of the TLC plate. Do not use a pen, as the ink may dissolve in the solvent.
Mark the spots where you will apply the sample along the baseline.
Spot the samples
Using a capillary tube, carefully apply a small drop of each sample dissolved in methanol onto the marked spots on the baseline. Allow the spots to dry before proceeding.
Prepare the developing chamber
Fill the developing chamber with a small amount of solvent (mobile phase) to a depth of approximately 0.5 cm. Make sure the solvent level is below the baseline on the TLC plate.
Place a lid or cover on the chamber to saturate the atmosphere with solvent vapor.
Develop the TLC plate
Carefully place the TLC plate in the developing chamber using tweezers, ensuring that the baseline remains above the solvent level.
Allow the solvent to rise up the plate by capillary action until it reaches approximately 1–2 cm below the top edge of the plate.
Remove the plate from the chamber and immediately mark the solvent front (the highest point reached by the solvent) with a pencil.
Visualize the spots
Examine the plate under UV light or use a suitable staining reagent to visualize the spots. Mark the center of each visible spot with a pencil.
Measure and calculate the Rf values
Measure the distance from the baseline to the center of each spot (distance traveled by the compound).
Measure the distance from the baseline to the solvent front (distance traveled by the solvent).
Calculate the Rf value for each compound using the formula:
Rf = Distance traveled by the compound/Distance traveled by the solvent
Document the results
Record the Rf values along with any observations (e.g., color, intensity) in your lab notebook.
Interpret the results
Compare the calculated Rf values with reference values to identify the compounds.
Notes:
i. Ensure that the TLC plate is handled carefully to avoid contamination.
ii. The Rf value is unitless and typically ranges from 0 to 1.
iii. Perform the TLC in a well-ventilated area or fume hood if using volatile or hazardous solvents.
Percentage yield calculations: Calculate the percentage yield of each reaction based on the following equation:
% Yield = experimental yield/theoretical yield × 100
Cell culture
Preparation of cells
Culture HDF, A-498, and ACHN cells in DMEM containing 4.5 g/L glucose, 1 mmol/L sodium pyruvate, and 200 mM L-glutamine supplemented with 10% (v/v) FBS and 1% (v/v) penicillin-streptomycin.
Maintain them in a humidified incubator with 5% CO2 at 37 °C.
Once the cells reach 80% confluency (occupying 80% of the available surface area of the culture vessel), subculture them using a trypsin-EDTA solution (0.05%).
Passage of cell lines
Subculture A498, ACHN, and HDF cells when they reach approximately 80% cell confluency. After discarding the media, rinse the cell monolayer with PBS. Incubate the cells with 25% Trypsin/5 mM EDTA solution in PBS (pH 7.4) at 37 °C for 5 min.
Mix the detached cell suspension with 10% FBS (v/v) complete DMEM in twice the volume of trypsin to inhibit trypsin activity. Centrifuge the cell suspension at 300× g for 5 min.
Discard the supernatant and resuspend the cell pellet in complete DMEM growth medium. Seed the cells in a new tissue culture flask.
Determination of cell number
Use a hemocytometer for cell counting. Load a 10 µL aliquot of the cell suspension into the square of the hemocytometer.
Count the cells three times, focusing on the center of the square under the inverted light microscope using a 20× objective lens. Calculate the cell concentration using the following equation:
Cell number/mL = number of counted cells × dilution factor/mm2 × chamber depth
Cryopreservation of cell lines
After trypsinizing the cells and counting them, suspend the cells with a freezing mixture consisting of 10% (v/v) DMSO in heat-inactivated FBS.
Suspend approximately 1 × 106 cells in 1 mL of the freezing mixture and transfer the suspension into a cryovial.
Place the cryovials in a -80 °C freezer for at least 16 h. Then, transfer the cryovials into a liquid nitrogen tank for long-term storage.
Thawing of cell lines
Retrieve the cryopreserved cells from either the liquid nitrogen tank or the -80 °C freezer and quickly warm them up to 37 °C in a water bath.
Add the cell suspension dropwise into complete DMEM to prevent cell disruption due to the osmotic pressure difference between the freezing mixture and the medium.
Centrifuge the cell suspension at 300× g for 5 min.
Discard the supernatant and resuspend the cell pellet in 5 mL of complete DMEM.
Seed the cells into a T-25 tissue culture flask.
After the cells attach to the flask, which usually takes approximately 12–24 h, exchange the medium with fresh complete DMEM to remove excess DMSO.
WST-1 cell proliferation assay
Seed HDF (5,000 cells/well), A-498 (10,000 cells/well), and ACHN (10,000 cells/well) cells in 100 μL into 96-well plates and allow them to adhere overnight (~16 h).
Treat the cells with H2O2 (0.5, 1, 1.5, 2, and 2.5 mM) in FBS-free DMEM for 2 h and wash the cells with serum-free media to remove H2O2.
Incubate cells with 2Hzin5NP (5, 10, 15, 20, 25, and 50 μM) in PBS (pH 7.4) for 30 min at 37 °C.
After removing the media, wash the cells thrice with PBS (5 min washes) and then incubate them with DMEM at 37 °C for 24 h.
Assess the effects of H2O2 and subsequent 2Hzin5NP treatment on cell proliferation using the WST1 assay, following the manufacturer’s instructions.
Measure absorbance values at 450 and 650 nm using a Varioskan Lux multimode microplate reader. Calculate the percentage of cell viability by normalizing the values to non-treated control cells, which should be adjusted to 100% using the formula: Asample × 100/Acontrol.
Labeling of carbonylation
Pre-treat HDF, A-498, and ACHN cells with increasing concentrations of sodium pyruvate (0, 1, and 2 mM) in complete DMEM for 1 h at 37 °C. After incubation, treat the cells with 2 mM H2O2 in FBS-free medium at 37 °C for 2 h.
Following H2O2 treatment, discard the medium and wash the cells once with PBS.
For hydrazine labeling, incubate A-498 cells with 20 μM 2Hzin5NP, while subjecting ACHN and HDF cells to 15 μM 2Hzin5Np in PBS for 30 min.
To induce endogenous carbonylation, serum-starve the cells in DMEM for up to 24 h and then label them with 2Hzin5NP.
Image the cells using a Zeiss LSM 800 confocal microscope at room temperature after washing them once with PBS.
Excite the samples using diode 405 and 488 lasers and collect the emission using long pass (LP) 435 and 518 filters.
Quantification of carbonylation in cell lysate
Collect cell pellets from HDF, A-498, and ACHN cells and suspend them in 500 μL of lysis buffer containing 0.05 mM PMSF and a protease inhibitor cocktail dissolved in dH2O.
Lyse the cells by subjecting them to six consecutive freeze-thaw cycles using liquid nitrogen and a water bath at 37 °C.
Determine the protein content using the DC protein assay following the manufacturer’s instructions. Generate a standard curve using BSA standards in the concentration range of 0.05–1 mg/mL.
Measure the fluorescence intensity of H2O2-treated and labeled cell lysates using a Varioskan Lux multimode microplate reader with excitation wavelength set at 396 nm and emission wavelength set at 502 nm.
Biochemical analysis
Cytotoxicity of H2O2 treatment
Seed A498, ACHN, and HDF cells at a density of 10,000 cells/well, and HDF cells at a density of 5,000 cells/well into a 96-well plate.
Incubate the cells for 9 h.
Treat the cells with 0.5, 1, 1.5, 2, and 2.5 mM H2O2 in FBS-free standard DMEM for 120 min.
Following H2O2 treatment, change the medium with standard complete DMEM and incubate the cells at 37 °C for 24 h.
Dissolve 10% WST-1 reagent in standard complete DMEM, then add 50 µL of the WST-1 mix to the cells and incubate for 1 h at 37 °C.
Measure absorbance values at 450 and 650 nm using a Varioskan Lux multimode microplate reader.
Subtract the background absorbance at 650 nm from the formazan absorbance at 450 nm.
Cytotoxicity of 2-Hydrazine 5-Nitrophenol labeling
Plate A498, ACHN, and HDF cells at a density of 10,000 cells/well, and HDF cells at a density of 5,000 cells/well into a 96-well plate.
Incubate the cells for 9 h and treat the cells with 5, 10, 15, 20, 25, and 50 µM 2Hzin5np in PBS (pH 7.4) for 30 min at 37 °C.
After treatment, discard the labeling reagent and wash the cells with PBS (pH 7.4).
Incubate the cells with standard complete DMEM at 37 °C for 24 h.
To measure cell viability, add 50 µL of standard complete DMEM containing 10% WST-1 reagent to each well and incubate for 1 h at 37 °C.
Reactive oxygen species detection assay
To detect the level of ROS after hydrogen peroxide treatment, label cells with DCFDA (2′,7′-dichlorofluorescin diacetate), a cellular reactive oxygen species detection assay kit. Count A498 and ACHN cells, seed them into a 96-well plate at a density of 10,000 cells/well, and incubate for 24 h.
While A498 cells are incubated with the maximum non-toxic dose of H2O2 at 2.5 mM, ACHN cells are incubated with the maximum non-toxic dose of H2O2 at 2 mM in FBS-free standard DMEM for 120 min. After treatment, discard the medium and wash the cells with PBS (pH 7.4).
Then, load 50 µL of 20 µM DCFDA in PBS (pH 7.4) into each well and incubate cells for 3 min in the dark at room temperature. Subsequently, wash cells with PBS (pH 7.4) and immediately image with a fluorescence microscope using a green fluorescence filter.
Spectroscopic and chemical analysis
General calibration and maintenance protocol for fluorescence and UV-Vis experiments
Fluorescence experiments (FluoroMax-4)
Warm-up period: Turn on the FluoroMax-4 and allow it to warm up for at least 30 min prior to use. This ensures optimal performance and accurate measurements.
Automatic calibration: The FluoroMax-4 automatically calibrates itself upon startup. Confirm that the instrument has completed its calibration and is ready for new experiments or stored routines.
Baseline check with water sample: Before beginning any analyses, run a water sample to verify that the fluorescence spectrum is correctly set and that there are no unexpected peaks or noise. This step helps ensure that the instrument is functioning properly.
Sample consistency: Prepare fresh samples for analysis and ensure that all emission spectra measurements are conducted on the same day to avoid any variations due to sample degradation or changes in environmental conditions.
Routine maintenance:
Clean the sample compartment: Regularly clean the sample compartment and cuvette holders to prevent contamination and ensure accurate readings.
Inspect and clean optical components: Periodically check the mirrors, lenses, and filters for dust or smudges. Clean with appropriate optical-grade materials as needed.
Software updates and calibration verification: Check for any software updates and perform manual calibration verification if required, especially after any major software or hardware changes.
UV-Vis experiments (UV-Vis spectrophotometer)
Warm-up period: Turn on the UV-Vis spectrophotometer and allow it to warm up for at least 30 min before use. This is essential for obtaining stable and reliable measurements.
Blank measurement: Before analyzing the actual sample, run a blank measurement using the same solvent as the sample. This step zeroes the instrument and accounts for any absorbance from the solvent, ensuring that only the sample's absorbance is measured.
Sample analysis: After recording the blank, proceed with the analysis of your sample. Make sure to use matched cuvettes and handle them carefully to avoid fingerprints or scratches that could interfere with the measurement.
Routine maintenance:
Clean cuvettes and sample holders: Clean all cuvettes and sample holders before and after each use to prevent cross-contamination.
Inspect optical pathways: Regularly check and clean the light source, detectors, and optical pathways. Ensure there are no obstructions or contaminants that could affect the results.
Wavelength calibration check: Perform a wavelength calibration check periodically to ensure the spectrophotometer is accurately measuring across its range. Use calibration standards or reference materials as needed.
Environmental conditions: Maintain a stable environment around both instruments, free from vibrations, temperature fluctuations, and excessive light, to ensure the accuracy and reliability of measurements.
UV-Vis absorbance and emission analysis
Acquire absorption spectra using an Agilent/HP 8453 UV-Visible spectrophotometer, employing a Starnacell Hellma quartz back wall cuvette with a 1 cm path length.
Record emission spectra at room temperature using a Jobin Yvon Horiba FluoroMax-4 spectrofluorometer, utilizing a Starnacell Hellma 2 × 10 mm fluorescence cuvette, oriented such that the light passes through the shorter path.
Fluorescence quantum yield
Determine fluorescence quantum yields (ΦF) in dilute solutions with an absorbance below 0.1 at the excitation wavelength.
Quinine sulfate in 0.1 M H2SO4 (λex = 347 nm, ΦF = 0.57) serves as the standard.
All spectra were recorded with a Fluoromax-4 spectrophotometer at 23 °C. Quantum yields are calculated using the following equation:
ΦF=ΦR·(AR/A) ΦS·(AS/A)
NMR analysis
Run the 1H-NMR and 13C-NMR spectra for the following listed synthesized molecules using an AVANCE III 500 MHz spectrometer (Bruker), with TMS as the internal standard. Chemical shift multiplicities are denoted using the following abbreviations: s = singlet, d = doublet, t = triplet, q = quartet, m = multiplet (denotes a complex pattern), dd = doublet of doublets, dt = doublet of triplets.
2-hydrazinyl-5-nitrophenol HCl salt
2-salicylaldehyde-5-nitrophenol
2-acetaldehyde-5-nitrophenol
Statistical analysis
Obtain all data from three independent experiments and present them as the mean ± SD (with error bars) for clarity.
Analyze the quantification of carbonylation in cell lysates using a two-tailed student t-test. Consider a p-value less than 0.05 as statistically significant.
Validation of protocol
Erkan et al. [18]. Design of Fluorescent Probes for Bioorthogonal Labeling of Carbonylation in Live Cells. Sci Rep. 10 (1): 7668.
Acknowledgments
This protocol was adapted from our previous work, published in Sci Rep. 2020 [18].
The presented protocols and work have been partially supported by TUBITAK (113S812) and COST Action CM1004.
Competing interests
The authors declare no conflicts of interest.
Ethical considerations
No human or animal subjects were included in this study.
References
Dilek, O. and Bane, S. L. (2011). Synthesis and Spectroscopic Characterization of Fluorescent Boron Dipyrromethene-Derived Hydrazones. J Fluoresc. 21(1): 347–354.
Chen, X. and Wu, Y. W. (2016). Selective chemical labeling of proteins. Org Biomol Chem. 14(24): 5417–5439.
Dalle-Donne, I., Rossi, R., Giustarini, D., Milzani, A. and Colombo, R. (2003). Protein carbonyl groups as biomarkers of oxidative stress. Clin Chim Acta. 329: 23–38.
Dilek, O. (2022). Current Probes for Imaging Carbonylation in Cellular Systems and Their Relevance to Progression of Diseases. Technol Cancer Res Treat. 21: 153303382211373.
Beatty, K. E., Liu, J. C., Xie, F., Dieterich, D. C., Schuman, E. M., Wang, Q. and Tirrell, D. A. (2006). Fluorescence Visualization of Newly Synthesized Proteins in Mammalian Cells. Angew Chem Int Ed. 45(44): 7364–7367.
Brand, M. D. (2016). Mitochondrial generation of superoxide and hydrogen peroxide as the source of mitochondrial redox signaling. Free Radical Biol Med. 100: 14–31.
Suzuki, Y. J., Carini, M. and Butterfield, D. A. (2010). Protein Carbonylation. Antioxid Redox Signal. 12(3): 323–325.
Schieber, M. and Chandel, N. S. (2014). ROS Function in Redox Signaling and Oxidative Stress. Curr Biol. 24(10): R453–R462.
Plass, T. and Schultz, C. (2011). Covalent Labeling of Biomolecules in Living Cells. In: Advanced fluorescence reporters in chemistry and biology III: applications in sensing and imaging. 225–261.
Cairns, R. A., Harris, I. S. and Mak, T. W. (2011). Regulation of cancer cell metabolism. Nat Rev Cancer. 11(2): 85–95.
Fedorova, M., Bollineni, R. C. and Hoffmann, R. (2013). Protein carbonylation as a major hallmark of oxidative damage: Update of analytical strategies. Mass Spectrom Rev. 33(2): 79–97.
Mukherjee, K., Chio, T. I., Gu, H., Banerjee, A., Sorrentino, A. M., Sackett, D. L. and Bane, S. L. (2017). Benzocoumarin Hydrazine: A Large Stokes Shift Fluorogenic Sensor for Detecting Carbonyls in Isolated Biomolecules and in Live Cells. ACS Sens. 2(1): 128–134.
Mukherjee, K., Chio, T. I., Sackett, D. L. and Bane, S. L. (2015). Detection of oxidative stress-induced carbonylation in live mammalian cells. Free Radical Biol Med. 84: 11–21.
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Article Information
Publication history
Received: Apr 5, 2024
Accepted: Sep 25, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biophysics > Microscopy > Two-photon laser scanning microscopy
Cell Biology > Cell imaging > Fluorescence
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Measurement of the Activity of Wildtype and Disease-Causing ALPK1 Mutants in Transfected Cells With a 96-Well Format NF-κB/AP-1 Reporter Assay
TS Tom Snelling
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5113 Views: 273
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Original Research Article:
The authors used this protocol in Proceedings of the National Academy of Sciences of the United States of America Dec 2023
Abstract
Alpha-protein kinase 1 (ALPK1) is normally activated by bacterial ADP-heptose as part of the innate immune response, leading to the initiation of downstream signalling events that culminate in the activation of transcription factors such as NF-κB and AP-1. In contrast, disease-causing mutations in ALPK1 that cause ROSAH syndrome or spiradenoma allow ALPK1 to be activated in cells in the absence of bacterial infection (i.e., without ADP-heptose). This protocol describes a semi-quantitative reporter assay based on ALPK1 knockout HEK-Blue cells that measures the activity of transfected wildtype and disease-causing forms of ALPK1 by virtue of their ability to activate the transcription factors NF-κB and AP-1. These cells express a synthetic gene encoding alkaline phosphatase under the control of an NF-κB/AP-1-dependent promoter, and consequently, the activation of ALPK1 leads to the production of alkaline phosphatase, which is secreted into the culture media and can be measured colorimetrically at 645 nm after the addition of a detection reagent.
Key features
• Highly sensitive reporter assay allowing detection of low-level activity arising from ALPK1 mutants
• Optimised in 96-well plate format, requiring only 60,000 transfected ALPK1 KO HEK-Blue cells per well
• Rapid experimental design, taking only four days from start to finish
• Suitable for screening ALPK1 variants of unknown significance in an arrayed 96-well format
Keywords: ALPK1 ADP-heptose ROSAH Spiradenoma Transcriptional reporter Transfection NF-κB AP-1
Background
Alpha-protein kinase 1 (ALPK1) is an atypical protein kinase that is activated allosterically by the binding of a bacterial nucleotide sugar, ADP-heptose, to its N-terminal ADP-heptose binding domain [1]. This enables ALPK1 to phosphorylate TIFA (TRAF-interacting protein with forkhead-associated domain) at Thr9, triggering its polymerization and the consequent activation of downstream signalling events that lead to activation of transcription factors such as NF-κB and AP-1 [1,2].
ROSAH syndrome (retinal dystrophy, optic nerve edema, splenomegaly, anhidrosis, and migraine headache) is an autosomal dominant genetic disorder caused by specific mutations in ALPK1 [3]. Most of the cases of ROSAH syndrome reported so far involve the mutation of Thr237 to Met, but cases have also been identified to be caused by the mutation of Tyr254 to Cys or Ser277 to Phe [4,5]. Thr237 directly interacts with ADP-heptose, whereas Tyr254 and Ser277 are outside of the ADP-heptose binding site itself but interact with each other through a hydrogen bond [5]. In contrast, ALPK1[Val1092Ala] is a driver mutation of a rare type of hair follicle tumour called spiradenoma, which can transform into an invariably fatal form known as spiradenocarcinoma [6].
The overexpression of these disease-causing ALPK1 mutants in ALPK1 knockout (KO) cells leads to the activation of NF-κB/AP-1-dependent gene transcription in the absence of ADP-heptose by a mechanism that is dependent on an intact ADP-heptose binding site [5,7]. This conundrum was resolved when it was found that these disease-causing ALPK1 mutants are activated by mammalian nucleotide sugars such as UDP-mannose and ADP-ribose, in addition to ADP-heptose [5,7]. In contrast, the normal form of ALPK1 is specifically activated by ADP-heptose.
This protocol outlines a cell-based assay designed to measure the activation of NF-κB and AP-1-dependent gene transcription by transfected ALPK1 mutants as utilised in previous studies on this topic [5,7]. The assay uses ALPK1 KO HEK-Blue cells, which express a synthetic gene encoding a secreted form of alkaline phosphatase under the control of an NF-κB/AP-1-dependent promoter. When these cells are transfected with plasmids encoding either wildtype or mutant forms of ALPK1, their ability to activate gene transcription in the presence or absence of ADP-heptose can be quantified by measuring the levels of alkaline phosphatase in the culture medium. This is achieved through a simple colorimetric assay where the absorbance is measured at 645 nm after the addition of a detection reagent, providing an efficient method for assessing ALPK1 activity. These reporter cells have been used previously to screen for activators or inhibitors of immune pathways [8], but here we expand their utility by showing their successful use to study variants of unknown significance.
The overexpression of ALPK1 mutants in this assay enables the detection of low-level activity, which can be difficult to observe when mutants are instead expressed under their endogenous promoter in immune cells and the resulting production of cytokines or chemokines is subsequently measured. Moreover, generating complex cell models such as these would be impractical for the screening of multiple ALPK1 variants of unknown significance at scale, but can be completed with minimal effort in four days using the protocol described here.
Materials and reagents
Biological materials
Cells should be cultured by incubation at 37 °C with 5% CO2 and tested regularly for mycoplasma using a MycoAlert Mycoplasma Detection Kit (Lonza, catalog number: LT07-318). The cells should be passaged once confluent at a ratio of 1:10 (v/v), and not used beyond 30 passages.
ALPK1 KO HEK-Blue cells (InvivoGen, #hkb-koalpk)
Reagents
The storage conditions for reagents are given in parentheses when not at room temperature. Commercial stock solutions are listed where possible, but many of these can also be prepared using standard methods.
ADP-heptose triethylammonium salt (InvivoGen, catalog number: tlrl-adph-l) (-80 °C)
Note: Resuspend 250 μg in 694 μL of PBS to generate a 0.5 mM stock. Aliquot and store at -80 °C.
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: 11960-085) (4 °C)
Heat-inactivated foetal bovine serum (FBS) (Labtech, catalog number: FB-1001/500) (-80 °C)
Note: FBS must be heat-inactivated to avoid high background alkaline phosphatase activity.
200 mM L-glutamine (Gibco, catalog number: 25030024) (-20 °C)
Penicillin-streptomycin 100× stock (Gibco, catalog number: 15140122) (-20 °C)
Opti-MEM I reduced serum medium (OptiMEM) (Gibco, catalog number: 31985062) (4 °C)
Lipofectamine 2000 (Thermo Fisher, catalog number: 11668019) (4 °C)
Trypan Blue solution (Gibco, catalog number: 11538886)
Quanti-BLUE solution 100× stock (InvivoGen, catalog number: rep-qbs) (-20 °C)
Note: Dilute the provided solutions together to 1× in water, prepare 20 mL aliquots, and store at -20 °C.
1 M magnesium chloride (Sigma-Aldrich, catalog number: M8266)
Phosphate-buffered saline (PBS) (Gibco, catalog number: 10010023) (4 °C)
Trypsin-EDTA solution (Gibco, catalog number: 25200056) (4 °C)
SDS sample buffer 4× stock (Millipore, catalog number: 70607) (4 °C)
Benzonase endonuclease (Millipore, catalog number: E1014) (-20 °C)
Complete EDTA-free protease inhibitor cocktail (Roche, catalog number: 11873580001) (4 °C)
Note: Dissolve a tablet in 1 ml of water to generate a 50× stock. Aliquot and store at -20 °C.
Antibodies
FLAG antibody (Sigma-Aldrich, catalog number: F3165), store at -20 °C
GAPDH (Cell Signalling Technology, catalog number: 2118), store at -20 °C
Plasmids
These plasmids encode WT and mutant forms of FLAG-ALPK1 under the control of a CMV promoter and are available to request via the MRC PPU Reagents and Services website (https://mrcppureagents.dundee.ac.uk). They were purified using NucleoBond Xtra Midi Endotoxin-Free kits (Macherey-Nagel, catalog number: 740420), resuspended to 0.5 mg/mL in endotoxin-free water and stored at -20 °C.
pcDNA5-FRT-TO-FLAG (empty vector, denoted “EV”) (catalog number: DU41457)
pcDNA5-FRT/TO-FLAG-ALPK1 (catalog number: DU65668)
pcDNA5-FRT-TO-FLAG-ALPK1[T237M] (catalog number: DU65723)
pcDNA5-FRT-TO-FLAG-ALPK1[Y254C] (catalog number: DU71685)
pcDNA5-FRT-TO-FLAG-ALPK1[S277F] (catalog number: DU71952)
pcDNA5-FRT-TO-FLAG-ALPK1[V1092A] (catalog number: DU65703)
pcDNA5-FRT-TO-FLAG-ALPK1[R150A] (catalog number: DU71740)
pcDNA5-FRT-TO-FLAG-ALPK1[R150A/T237M] (catalog number: DU71743)
pcDNA5-FRT-TO-FLAG-ALPK1[R150A/Y254C] (catalog number: DU71741)
pcDNA5-FRT-TO-FLAG-ALPK1[R150A/S277F] (catalog number: DU71954)
pcDNA5-FRT-TO-FLAG-ALPK1[R150A/V1092A] (catalog number: DU71742)
Solutions
Note: Storage conditions are given in parentheses.
Antibiotic-free culture media (4 °C) (see Recipes)
Culture media (4 °C) (see Recipes)
SDS lysis buffer (use immediately) (see Recipes)
Recipes
Antibiotic-free culture media (1 bottle)
Reagent Final concentration Quantity to add
DMEM Not applicable 500 mL
FBS 10% (v/v) 50 mL
200 mM L-Glutamine 2 mM 5.6 mL
Culture media (1 bottle)
Reagent Final concentration Quantity to add
DMEM Not applicable 500 mL
FBS 10% (v/v) 50 mL
200 mM L-Glutamine 2 mM 5.6 mL
Penicillin-streptomycin 100× 1× 5.6 mL
SDS lysis buffer (5 mL)
Reagent Final concentration Quantity to add
SDS sample buffer 4× 1× 1.25 mL
Benzonase endonuclease 0.2% (v/v) 10 μL
1 M magnesium chloride 1 mM 5 μL
Protease inhibitor cocktail 50× 1× 100 μL
Water Not applicable 3.635 mL
Laboratory supplies
Similar products can also be used, but those marked by an asterisk are highly recommended.
10 cm Nunc cell culture dishes (Thermo Fisher, catalog number: 150318)*
96-well Nunc cell culture plates (Thermo Fisher, catalog number: 167008)*
25 mL sterile reagent reservoir (Thermo Fisher, catalog number: 11405758)
Clear sealing tape for 96-well plates (Thermo Fisher, catalog number: 10105383)
50 mL conical centrifuge tubes (Greiner, catalog number: 227261)
15 mL conical centrifuge tubes (Greiner, catalog number: 188271)
Serological pipettes (Thermo Fisher, catalog number: 10710810)
Safe-Lock 1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 30123611)
Cellometer counting chambers (Nexcelom, catalog number: 11522186)
Combitips advanced 0.5 mL (Eppendorf, catalog number: 30089421)
Equipment
Similar equipment from other suppliers can also be used instead of those listed below. Those that have been discontinued do not have a catalog number indicated.
Cellometer Auto 2000 (Nexcelom Bioscience)
Multipette M4 (Eppendorf, catalog number: 4982000012)
8-channel adaptor (Integra Biosciences, catalog number: 10023451)
8-channel pipettor 20–200 μL (VWR, catalog number: 89079-948)
Pipetman 4-Pipette Kit (Gilson, catalog number: F167360)
Stripettor Ultra pipet controller (Corning, catalog number: 4099)
CB150 cell culture incubator (Binder)
BioMAT 2-SF cell culture hood (Conditioned Air Solutions)
Allegra X-12 benchtop centrifuge (Beckman Coulter, catalog number: 392474)
QBT2 dry block heater (Grant)
Epoch plate reader (BioTek)
Procedure
All steps are performed in a sterile culture hood, and the cells are cultured at 37 °C with 5% CO2. Before starting the procedure, a confluent 10 cm dish of ALPK1 KO HEK-Blue cells in culture media is required, sufficient for 2 × 96-well plates.
Prepare complexes of lipofectamine 2000 and plasmid in a 96-well culture plate (Day 1)
Dilute lipofectamine 2000 and add to the required wells:
Calculate the number of wells to be transfected plus 10% extra (for 88 wells, prepare 96 wells).
Each well requires 0.5 μL of lipofectamine 2000 in 24.5 μL of OptiMEM. Prepare a mastermix for 96 wells by diluting 48 μL of lipofectamine 2000 in 2352 μL of OptiMEM. Invert five times to mix thoroughly.
Transfer diluted lipofectamine 2000 to a reagent reservoir and use a multi-channel pipette to add 25 μL per required well within a 96-well culture plate.
Prepare a mastermix for each plasmid and add to the relevant wells:
Calculate the number of wells to be transfected with each plasmid plus 20% extra. Each plasmid should be assayed in quadruplicate, with and without ADP-heptose. Leave an empty column for a media-only control in the 96-well plate.
Each well requires 0.2 μg of plasmid in 25 μL of OptiMEM. For a 10-well mastermix of plasmid encoding WT ALPK1 (i.e. sufficient for the required 8 wells, plus 20% extra), add 2 μg of this plasmid to 250 μL of OptiMEM. Invert five times to mix thoroughly.
Add 25 μL of each diluted plasmid to relevant wells of the 96-well culture plate using a repeat pipettor, remembering to change tips for different plasmids.
Leave the culture plate undisturbed for at least 20 min, but no more than 6 h, to allow the formation of lipid-DNA complexes.
Note: Prepare the cell suspension below in the meantime, which should take approximately 20 min.
Add cell suspension directly into the wells of the 96-well plate containing lipid-DNA complexes (Day 1)
Aspirate the culture medium from a confluent 10 cm dish of ALPK1 KO cells and replace with 10 mL of PBS using a serological pipette.
Aspirate the PBS and add 2 mL of trypsin-EDTA solution. Return the dish to the incubator until the cells have detached, which takes typically 2–3 min.
Note: If trypsinization is taking significantly longer than this, ensure that all residual culture medium and PBS are removed during the aspiration steps since FBS present in the culture media can inhibit trypsin activity.
Add 10 mL of antibiotic-free culture media and pipette up and down with a serological pipette until a single-cell suspension has been produced. Transfer this cell suspension to a 15 mL canonical tube.
Mix 20 μL of cell suspension with 20 μL of trypan blue solution and count the number of cells using standard methods, ensuring that cell viability is at least 90%.
Dilute the cell suspension to 600,000 cells/mL in antibiotic-free culture media, ensuring that the total volume of the diluted cell suspension is at least 10% more than the total volume needed for the experiment.
Add 100 μL per required well (i.e., 60,000 cells) into the wells of the 96-well plate that contain lipid-DNA complexes using a reagent reservoir and multi-channel pipette (see Figure 1A for expected density).
Figure 1. Appearance of ALPK1 KO HEK-Blue cells. Density of ALPK1 KO HEK-Blue cells 1 h after plating into 96-well plates (A) and 24 h post-transfection, prior to stimulation (B).
Return the cells to the 37 °C incubator (5% CO2) for 24 h.
Replacement of culture media and stimulation of relevant wells with ADP-heptose (Day 2)
Prepare a mastermix of culture media containing either PBS or 5 μM ADP-heptose:
At this stage, cells should be confluent (see Figure 1B). Each well requires 75 μL of culture media containing either 5 μM ADP-heptose or an equivalent volume of PBS.
For PBS control wells, dilute 36 μL of PBS in 3600 μL of culture media.
For ADP-heptose wells, dilute 36 μL of 0.5 mM ADP-heptose in 3600 μL of culture media.
Aspirate wells and add the culture media:
After 24 h of transfection, carefully aspirate the existing culture media using an 8-channel adaptor, being careful not to disturb the cells.
Replace with 75 μL of culture media (with antibiotics) containing either PBS or ADP-heptose according to the experimental plan and return cells to the incubator for 24 h.
As a control for the absorbance of the culture media itself, transfer 75 μL of the fresh culture media to an empty column.
Quantification of alkaline phosphatase activity in the culture media (Day 3)
Collection of culture media and lysis of cells:
After 24 h of stimulation, transfer 50 μL of culture media from each well into a new 96-well plate using a multi-channel pipette, changing tips each time.
Carefully aspirate the remaining culture media using an 8-channel adaptor, being careful not to disturb the cells. Remove residual media by centrifuging the culture plate upside down on a stack of tissues for 30 s at 500× g.
To each well, add 30 μL of SDS lysis buffer, seal with plastic, and discard the plate lid.
Incubate the plate at 75 °C for 10 min by placing it on a heat block.
Analyse the unstimulated samples in duplicate (i.e., 2 of the 4 technical replicates) by SDS-PAGE followed by immunoblotting with anti-GAPDH and anti-FLAG antibodies using standard methods.
Note: This step is essential as it will indicate any variability in the levels of expression of the different ALPK1 variants as well as the similarity in the expression levels between technical replicates.
Detection of absorbance at 645 nm:
Add 150 μL of Quanti-Blue 1× solution to each well containing 50 μL of culture media (including the control column with fresh culture media), changing tips each time.
Incubate at room temperature and take absorbance readings at 645 nm every 15 min until an optimal signal-to-noise ratio is achieved, typically 1 h (Figure 2A).
Figure 2. Disease-causing ALPK1 mutants have activity in the absence of ADP-heptose, which is dependent on an intact ADP-heptose binding site. (A) Appearance of the reporter plate after incubation of the culture media for 1 h with Quanti-Blue solution to detect alkaline phosphatase activity. (B) Absorbance values at 645 nm were plotted, with background absorbance from culture media alone subtracted from all other values. The bar heights represent the mean values and the error bars indicate plus and minus one standard deviation.
Data analysis
Two alternative approaches can be used for data analysis. In both cases, the bar heights should represent the mean values, with error bars indicating plus and minus one standard deviation.
In the first approach, the average absorbance values from the culture medium-only control column are subtracted from all other values, and the resulting data are plotted (see Figure 2B). The resulting y-axis is the relative activation of NF-κB/AP-1, with the culture medium-only control set to zero since no cells were plated. In the second approach, the culture medium-only control can be excluded, and the average absorbance values from the empty vector-only controls can instead be subtracted from the values obtained from conditions where ALPK1 was expressed. This approach removes basal NF-κB/AP-1-dependent gene transcription that is independent of ALPK1, and the y-axis is, therefore, the relative ALPK1 activity.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Snelling, T et al (2024). Discovery and Functional analysis of a novel ALPK1 variant in ROSAH syndrome. bioRxiv (Figure 2, panels A–C; Figure 4, panels B–C)
Snelling, T et al (2023). ALPK1 mutants causing ROSAH syndrome or Spiradenoma are activated by human nucleotide sugars. Proc Natl Acad Sci USA (Figure 2, panel C; Figure 3, panels A–C)
General notes and troubleshooting
Troubleshooting
Low or undetectable levels of NF-κB/AP-1 dependent gene transcription
This is most likely caused by poor transfection efficiency or low-quality plasmid DNA. To confirm this, check the percentage of GFP-positive cells by flow cytometry/microscopy 24 h after transfection with a GFP plasmid such as pcDNA5-FRT/TO-GFP-ALPK1 (MRC PPU Reagents and Services, catalog number: DU78380).
Below are suggestions for troubleshooting:
Ensure that 10 cm dishes of cells are not overconfluent prior to plating into 96-well plates, as this can reduce the transfection efficiency and expression of the ALPK1 constructs.
Ensure that cells are evenly distributed after plating into 96-well plates and are not clumped together.
Confirm that cells are free from mycoplasma and other contaminants by routine testing.
Thaw a new vial of cells and ensure that they are passaged at least twice prior to transfection experiments.
Check DNA purity by measuring the ratio of absorbance at 260 and 280 nm using a spectrophotometer, which should be between 1.8 and 2.0. In addition, confirm DNA integrity by agarose gel electrophoresis.
Confirm that cells still express the gene encoding alkaline phosphatase by stimulation with 10 ng/mL human IL-1β (InvivoGen, catalog number: rcyec-hil1b) and measuring alkaline phosphatase activity in the culture media after 24 h.
Cell death during the transfection procedure
Antibiotics should be excluded from the culture media during the process of transfection, which is observed to minimise cell toxicity in this assay.
Ensure that cells are plated at the required density to be confluent 24 h later, as it is observed that cell toxicity from transfection is greater at lower cell densities.
High background NF-κB/AP-1 dependent gene transcription
To determine the source of a high background signal, perform control experiments comparing alkaline phosphatase activity in the culture media of cells transfected with empty vector compared to lipofectamine alone (i.e., without DNA) and in fresh culture media with and without FBS (i.e., no cells).
Ensure that FBS is heat-inactivated to denature any alkaline phosphatases that are present. It may be necessary to test multiple batches of FBS from different suppliers to minimise the background signal.
Confirm that cells are free from mycoplasma and other contaminants by routine testing.
Thaw a new vial of cells and passage at least twice prior to performing experiments.
Ensure that plasmids are not contaminated with bacterial components by using endotoxin-free kits.
Cell detachment during aspiration and wash steps
HEK293 cells are weakly adherent and therefore easily detached during aspiration and addition steps.
Do not touch the bottom of the wells when aspirating or adding culture media.
Use gel-loading tips intended for SDS-PAGE on the end of an 8-channel adaptor to minimise cell detachment during aspiration steps.
When adding media, pipette slowly against the side of each well to minimise cell detachment.
Acknowledgments
This work was supported by a PhD studentship (#2087974) from the MRC.
Competing interests
The author declares no competing interests associated with this manuscript.
References
Zhou, P., She, Y., Dong, N., Li, P., He, H., Borio, A., Wu, Q., Lu, S., Ding, X., Cao, Y., et al. (2018). Alpha-kinase 1 is a cytosolic innate immune receptor for bacterial ADP-heptose. Nature. 561(7721): 122–126. https://doi.org/10.1038/s41586-018-0433-3
Snelling, T., Shpiro, N., Gourlay, R., Lamoliatte, F. and Cohen, P. (2022). Co-ordinated control of the ADP-heptose/ALPK1 signalling network by the E3 ligases TRAF6, TRAF2/c-IAP1 and LUBAC. Biochem J. 479(20): 2195–2216. https://doi.org/10.1042/bcj20220401
Williams, L. B., Javed, A., Sabri, A., Morgan, D. J., Huff, C. D., Grigg, J. R., Heng, X. T., Khng, A. J., Hollink, I. H., Morrison, M. A., et al. (2019). ALPK1 missense pathogenic variant in five families leads to ROSAH syndrome, an ocular multisystem autosomal dominant disorder. Genet Med. 21(9): 2103–2115. https://doi.org/10.1038/s41436-019-0476-3
Kozycki, C. T., Kodati, S., Huryn, L., Wang, H., Warner, B. M., Jani, P., Hammoud, D., Abu-Asab, M. S., Jittayasothorn, Y., Mattapallil, M. J., et al. (2022). Gain-of-function mutations in ALPK1 cause an NF-κB-mediated autoinflammatory disease: functional assessment, clinical phenotyping and disease course of patients with ROSAH syndrome. Ann Rheum Dis. 81(10): 1453–1464. https://doi.org/10.1136/annrheumdis-2022-222629
Snelling, T., Garnotel, L. O., Jeru, I., Tusseau, M., Cuisset, L., Perlat, A., Minard, G., Benquey, T., Maucourant, Y., Wood, N. T., et al. (2024). Discovery and Functional analysis of a novel ALPK1 variant in ROSAH syndrome. bioRxiv: e612837. https://doi.org/10.1101/2024.09.13.612837
Rashid, M., van der Horst, M., Mentzel, T., Butera, F., Ferreira, I., Pance, A., Rütten, A., Luzar, B., Marusic, Z., de Saint Aubain, N., et al. (2019). ALPK1 hotspot mutation as a driver of human spiradenoma and spiradenocarcinoma. Nat Commun. 10(1): e1038/s41467–019–09979–0. https://doi.org/10.1038/s41467-019-09979-0
Snelling, T., Saalfrank, A., Wood, N. T. and Cohen, P. (2023). ALPK1 mutants causing ROSAH syndrome or Spiradenoma are activated by human nucleotide sugars. Proc Natl Acad Sci USA. 120(50): e2313148120. https://doi.org/10.1073/pnas.2313148120
Hu, Z., Zhang, T., Jiang, S. and Yin, H. (2022). Protocol for evaluation and validation of TLR8 antagonists in HEK-Blue cells via secreted embryonic alkaline phosphatase assay. STAR Protoc. 3(1): 101061. https://doi.org/10.1016/j.xpro.2021.101061
Article Information
Publication history
Received: Jul 25, 2024
Accepted: Sep 24, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Cell Biology > Cell-based analysis > Protein synthesis
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Preparation of Protein Lysates Using Biorthogonal Chemical Reporters for Click Reaction and in-Gel Fluorescence Analysis
YX Yaxin Xu
TP Tao Peng
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5114 Views: 364
Reviewed by: Neha NandwaniDhananjay D ShindeFan Zheng
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Original Research Article:
The authors used this protocol in Molecular & Cellular Proteomics Mar 2024
Abstract
Bioorthogonal chemical reporters are non-native chemical handles introduced into biomolecules of living systems, typically through metabolic or protein engineering. These functionalities can undergo bioorthogonal reactions, such as copper-catalyzed alkyne-azide cycloaddition (CuAAC), with small-molecule probes, enabling the tagging and visualization of biomolecules. This approach has greatly enhanced our understanding of cellular dynamics, enzyme targeting, and protein post-translational modifications. Herein, we report a protocol for preparing protein lysates for click reaction and in-gel fluorescence analysis, exemplified using alk-16, a terminal alkyne-functionalized stearic acid analog, to investigate proteins with fatty acylation. This protocol provides methods for the fluorescent visualization of chemical reporter–labeled proteomes or individual proteins of interest (POIs).
Key features
• Metabolic incorporation of bioorthogonal chemical reporters into proteins in living cells
• Visualization of proteomes or specific proteins labeled with chemical reporters via in-gel fluorescence analysis
• Reliable, non-radioactive methods for investigating protein fatty acylation and other post-translational modifications
Keywords: Click chemistry Bioorthogonal chemical reporter Metabolic labeling In-gel fluorescence Post-translational modifications
Background
Bioorthogonal chemical reporters refer to non-native, non-perturbing chemical handles that can be specifically modified in living systems through highly selective reactions with exogenously delivered probes [1]. The bioorthogonal chemical reporter strategy involves incorporating unique functionalities, such as azide, alkyne, or alkene groups, into target biomolecules using the cellular biosynthetic machinery. Chemical labeling with small-molecule probes is then achieved via bioorthogonal click reactions, which are characterized by exceptional biorthogonality, biocompatibility, rapid kinetics, and high specificity in biological environments [1]. A prominent example of such click reactions is the copper-catalyzed alkyne-azide cycloaddition (CuAAC). In this reaction, an alkyne-tagged molecule reacts with an azide-tagged molecule in the presence of copper (I) to form a stable 1,4-disubstituted 1,2,3-triazole product via a [3+2] cycloaddition [2,3]. Over the past two decades, these strategies have enabled precise tracking and analysis of proteins and other biomolecules in complex biological environments, revolutionizing our understanding of biological systems. They have been applied to monitoring dynamic changes in cellular activity, profiling various cell types, states, or mutations, identifying enzyme targets, and exploring a wide range of post-translational modifications (PTMs) [4]. Herein, we present a detailed experimental protocol for the preparation of protein lysates for click reaction and in-gel fluorescence analysis, using alk-16 as a representative example. Alk-16 is a terminal alkyne-functionalized stearic acid analog that mimics endogenous long-chain fatty acids and can undergo a click reaction, allowing for the investigation of proteins with fatty acylation [5–7]. This includes the metabolic incorporation of chemical reporters into proteins in living cells, global visualization of reporter-labeled proteins in gels by selectively reacting alkynyl chemical reporter–labeled proteins in cell lysates with azido-rhodamine, and the analysis of reporter-labeled candidate proteins using immunoprecipitation, click chemistry, and fluorescence scanning (Figure 1). We aim to provide a general protocol for in-gel fluorescence analysis of proteins that are labeled by bioorthogonal chemical reporters. While we use alk-16 as an example to describe the protocol, it is also adaptable to other bioorthogonal chemical reporters, such as alkynyl-functionalized YnLac [8] and HMGAMyne [9] to detect and identify the cellular lactylated and HMGylated proteins.
Figure 1. Workflow for in-gel fluorescence analysis of alkynyl chemical reporter–labeled proteins. (A) Schematic for in-gel fluorescence analysis of chemical reporter-labeled proteomes. (B) Schematic for in-gel fluorescence analysis of chemical reporter-labeled protein of interest (POI).
Materials and Reagents
Reagents
DMSO (Sigma-Aldrich, catalog number: D2650)
DMEM (Thermo Fisher Scientific, catalog number: 11995065) or other culture media for growing the cell type of interest
Fetal bovine serum (FBS) (Thermo Fisher Scientific, catalog number: 26140079)
Charcoal/dextran-treated fetal bovine serum (Cytiva, catalog number: SH30068)
Opti-MEM (Thermo Fisher Scientific, catalog number: 31985070)
ViaFect transfection reagent (Promega, catalog number: E4981)
Benzonase (Sigma-Aldrich, catalog number: E1014)
EDTA-free protease inhibitor cocktail (Roche, catalog number: 11873580001)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S3014)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9541)
Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S3264)
Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P9791)
Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: L3771)
Trizma hydrochloride (Tris-HCl) (Sigma-Aldrich, catalog number: T3253)
Hydrochloric acid (HCl) (Sigma-Aldrich, catalog number: 258148)
Triton X-100 (Sigma-Aldrich, catalog number: T8787)
Sodium deoxycholate (Sigma-Aldrich, catalog number: D6750)
Triethanolamine (TEA) (Sigma-Aldrich, catalog number: 90279)
4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Sigma-Aldrich, catalog number: 54457)
BCA assay reagents (Thermo Fisher Scientific, catalog number: A55864)
Red anti-HA affinity gel (Sigma-Aldrich, catalog number: E6779)
Red anti-FLAG M2 affinity gel (Sigma-Aldrich, catalog number: F2426)
Anti-GFP nanobody agarose beads (AlpaLifeBio, catalog number: KTSM1301)
Tert-butanol (t-BuOH) (Sigma-Aldrich, catalog number: 471712)
Methanol (Sigma-Aldrich, catalog number: 34860)
Glycerol (Sigma-Aldrich, catalog number: G5516)
Bromophenol blue (Sigma-Aldrich, catalog number: B8026)
Bond-breaker TCEP solution, neutral pH (Thermo Fisher Scientific, catalog number: 77720)
Hydroxylamine hydrochloride (NH2OH·HCl) (Sigma-Aldrich, catalog number: 55460)
Tween-20 (Sigma-Aldrich, catalog number: 655205)
Acetic acid (Sigma-Aldrich, catalog number: A6283)
Clarity western ECL substrate (Bio-Rad, catalog number: 1705060)
Alk-16 (Sigma-Aldrich, catalog number: O8382; alternatively, it can be synthesized as described in Charron et al. [5])
Azido-rhodamine (the synthetic method has been described in Charron et al. [5])
Tris(2-carboxyethyl) phosphine hydrochloride (TCEP) (Sigma-Aldrich, catalog number: C4706)
Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA) (Sigma-Aldrich, catalog number: 678937)
Copper sulfate pentahydrate (CuSO4·5H2O) (Sigma-Aldrich, catalog number: C8027)
Tris-MOPS-SDS running buffer (GenScript, catalog number: M00138)
Coomassie Brilliant Blue staining solution (Beyotime, catalog number: P0003S)
Solutions
50 mM alk-16 (see Recipes)
PBS (1 L) (see Recipes)
RIPA buffer (see Recipes)
1% SDS lysis buffer (see Recipes)
4% SDS buffer (see Recipes)
HEPES buffer (see Recipes)
2 M NH2OH solution (see Recipes)
5 mM azido-rhodamine stock solution (see Recipes)
50 mM TCEP solution (see Recipes)
2 mM TBTA stock solution (see Recipes)
50 mM CuSO4 solution (see Recipes)
4× SDS-PAGE loading buffer (see Recipes)
Destaining buffer (see Recipes)
Recipes
50 mM alk-16
Dissolve 14.0 mg of alk-16 in 1 mL of DMSO.
PBS (1 L)
Measure and mix the following components:
8 g of NaCl (137 mM)
0.2 g of KCl (2.7 mM)
1.44 g of NaHPO (10 mM)
0.24 g of KHPO (1.8 mM)
Add ultrapure water to a final volume of 1 L.
Adjust the pH to 7.3 using HCl.
RIPA buffer (1 L)
Measure and mix the following components:
10 mL of Triton X-100 (1% v/v)
10 g of sodium deoxycholate (1% w/v)
1 g of SDS (0.1% w/v)
8.78 g of NaCl (150 mM)
50 mL of 1 M Tris-HCl pH 7.4 (50 mM)
Add ultrapure water to a final volume of 1 L.
1% SDS lysis buffer (1 L)
Measure and mix the following components:
10 g of SDS (1% w/v)
8.78 g of NaCl (150 mM)
50 mL of 1 M HEPES pH 7.4 (50 mM)
Add ultrapure water to a final volume of 1 L.
4% SDS buffer (1 L)
Measure and mix the following components:
40 g of SDS (4% w/v)
8.78 g of NaCl (150 mM)
50 mL of 1 M TEA pH 7.4 (50 mM)
Add ultrapure water to a final volume of 1 L.
HEPES buffer (1 L)
Measure and mix the following components:
8.78 g of NaCl (150 mM)
50 mL of 1 M HEPES pH 7.4 (50 mM)
Add ultrapure water to a final volume of 1 L.
2 M NH2OH solution (10 mL)
Dissolve 1.39 g of NH2OH·HCl in 8 mL of ultrapure water.
Adjust the pH to 7.4 using NaOH.
Add ultrapure water to a final volume of 10 mL.
5 mM azido-rhodamine stock solution
Dissolve 34.3 mg of azido-rhodamine in 10 mL of DMSO.
50 mM TCEP solution
Dissolve 14.3 mg of TCEP in 1 mL of ultrapure water.
2 mM TBTA stock solution
Dissolve 10.6 mg of TBTA in 10 mL of a 1:4 (v/v) mixture of DMSO and t-BuOH.
50 mM CuSO4·5H2O solution
Dissolve 12.5 mg of CuSO4·5H2O in 1 mL of ultrapure water.
4× SDS-PAGE loading buffer (100 mL)
Measure and mix the following components:
8 g of SDS (8% w/v)
20 mL of 1 M Tris-HCl pH 6.8 (200 mM)
40 mL of glycerol (40% v/v)
0.4 g of bromophenol blue (0.4% w/v)
Add ultrapure water to a final volume of 100 mL.
Destaining buffer (1 L)
Measure and mix the following components:
500 mL of methanol (50% v/v)
100 mL of acetic acid (10% v/v)
400 mL of ultrapure water (40% v/v)
Mix thoroughly to ensure the components are well combined.
Laboratory supplies
60 mm TC-treated culture dish (Corning, catalog number: 430166)
6-well clear TC-treated multiple-well plate (Corning, catalog number: 3516)
Pipette tips (Rainin, catalog numbers: 30180889, 30374583, 30296781)
10 mL serological pipette (Thermo Fisher Scientific, catalog number: 170367N)
Cell scraper (Corning, catalog number: 3010)
High-binding ELISA plate (JET-Bio, catalog number: FEP100096)
1.5 mL microfuge tube (Axygen, catalog number: MCT-150-C)
2.0 mL dolphin microcentrifuge tube (Sigma-Aldrich, catalog number: Z717533)
15-well 4%–20% Bis-Tris protein gel (GenScript, catalog number: M42015C)
Protein molecular weight standards (Yeasen, catalog number: 20351ES72)
Equipment
-80 °C freezer (Thermo Fisher Scientific, model: TDE50086FV-ULTS)
-20 °C freezer (Thermo Fisher Scientific, model: ES Series)
Biological safety cabinet (Thermo Fisher Scientific, model: 1300 Series A2)
Pipette (Rainin, model: LTS Pipette)
CO2 incubator (Thermo Fisher Scientific, model: Forma Series 3 WJ)
Inverted microscope (Nikon, model: Eclipse TS2)
Barnstead GenPure Pro ultrapure water system (Thermo Fisher Scientific, model: GenPure UV/UF)
Refrigerated centrifuge (Eppendorf, model: 5424R)
ThermoMixer (Eppendorf, model: ThermoMixer F1.5)
SpeedVac concentrator (Eppendorf, model: Concentrator Plus)
pH meter (Mettler Toledo, model: FE20K)
Multi-mode microplate reader (BioTek, model: EPOCH)
Analytical balance (Mettler Toledo, model: XS64)
Vacuum aspirator system (Dragon LAB, model: SAFEVAC)
Genie 2 vortex mixer (Scientific Industries, model: G560E)
Orbital shaker (Kylin-Bell, model: TS-200)
Dry bath incubator (Blue Pard, model: TU-10)
Protein electrophoresis system (Bio-Rad, model: PowerPac-Basic)
Trans-Blot SD semi-dry electrophoretic transfer cell (Bio-Rad, model: Trans-Blot Turbo)
ChemiDoc MP imaging system (Bio-Rad, model: ChemiDoc MP)
Procedure
Part I: Metabolic labeling of cellular proteins with alkynyl chemical reporters in living cells
Cell culture
Seed cells in 12-well plates or 60 mm dishes with normal growth medium (e.g., DMEM supplemented with 10% FBS). Incubate the cells overnight at 37 °C with 5% CO to allow them to adhere and grow.
Notes:
Seed the cells to reach 70%–80% confluency by the next day.
Both HeLa and HEK293T cells were used in this study, but other adherent cell types can also be used.
Cells in one well of a 12-well plate (approximately 100 μg of protein in cell lysates) are enough for in-gel fluorescence analysis of chemical reporter-labeled proteomes, while cells in a 60 mm dish (approximately 600 μg of protein in cell lysates) are needed for in-gel fluorescence analysis of individual POIs.
Transfection (optional)
Prepare transfection mixture. For cells cultured in a 12-well plate:
In a microcentrifuge tube, dilute 1 μg of plasmid encoding the wild-type (WT) or mutant POI in 100 μL of Opti-MEM.
Add 2.5 μL of ViaFect transfection reagent to the diluted DNA.
Mix the solution gently by pipetting and incubate the mixture at room temperature for 15 min.
Add the transfection mixture dropwise into the cell media.
Incubate the cells for 6 h at 37 °C in a CO incubator.
Optional: Transfection is optional; it depends on the experimental requirement. To analyze endogenous proteins labeled by chemical reporters, transfection is not needed. However, transfection to express an exogenous enzyme may be performed to analyze the substrates of that enzyme.
Notes:
The recommended concentration of plasmid stock for transfection is approximately 200–500 ng/μL.
Gently mix the transfection reagent and plasmid to avoid shear forces that can damage the plasmid DNA.
Select an appropriate transfection reagent and method based on the cell type.
Metabolic labeling with the chemical reporter
Carefully replace the existing cell culture media with fresh media, supplemented either with the chemical reporter or the solvent (e.g., DMSO) used to dissolve the chemical reporter as the control.
Notes:
In this study, the bioorthogonal chemical reporter alk-16, an alkynyl-functionalized fatty acid analog, was utilized to metabolically label fatty-acylated proteins in living cells.
Add fresh media gently to avoid dislodging the cells.
Optimize the concentration of the chemical reporter as needed. For alk-16, a final concentration of 50 µM has been found effective for efficient labeling.
Choose the appropriate type of cell culture media based on the experiment requirement. In this study, DMEM supplemented with 2% charcoal/dextran-treated fetal bovine serum was used to dissolve alk-16 and incubate the cells. The charcoal/dextran treatment removes lipids from the serum, facilitating the uptake of the fatty acid chemical reporter (e.g., alk-16) without competition from serum lipids.
Ensure to process a vehicle well without the chemical reporter as the control to evaluate the background fluorescence signal.
Incubate cells with the chemical reporter for 16 h at 37 °C.
Notes:
The incubation time can be optimized and adjusted based on specific experimental needs.
For different samples, ensure that the labeling period and incubation temperature are consistent.
After the incubation, discard the medium and resuspend the cells in 1 mL of ice-cold PBS by gently pipetting up and down. Centrifuge at 4,000× g for 2 min at 4 °C and discard the supernatant. Add fresh ice-cold PBS, resuspend the cells again by pipetting, and centrifuge once more. Remove the supernatant and collect the cell pellet.
Notes:
Use a cell scraper to detach adherent cells, avoiding trypsin as it may degrade cell surface proteins.
Ensure thorough washing to remove any residual media and chemical reporters (e.g., alk-16).
Handle the cells gently to avoid cell damage.
If not proceeding immediately to the following experiments, freeze the cell pellets in liquid nitrogen and store them at -80°C until further use.
Part II: In-gel fluorescence analysis of chemical reporter-labeled proteomes
Cell lysis and protein quantification
Lyse the cells labeled with chemical reporters by adding 50 μL of 1% SDS lysis buffer (see Recipes), supplemented with EDTA-free protease inhibitor cocktail and 0.2 μL of benzonase (50 U).
Notes:
Benzonase degrades nucleic acids to reduce viscosity.
Ensure that the lysis buffer is freshly prepared and supplemented with the protease inhibitor cocktail immediately before use. Avoid using protease inhibitors containing EDTA, as EDTA is incompatible with the click reaction.
It is preferable to use the lysis buffer in a volume to achieve a protein concentration above 2 mg/mL in the cell lysate.
Vortex vigorously to ensure complete lysis.
Centrifuge the lysates at 12,000× g for 20 min at room temperature. Collect the supernatant and discard the pellet (cell debris).
Determine the protein concentration using a standard BCA assay.
Note: Quantification of protein concentration is crucial for subsequent steps.
Aliquot equal amounts of protein (e.g., 100 μg) into 1.5 mL tubes and adjust the volume to 50 μL with the 1% SDS lysis buffer.
Add 39 μL of HEPES buffer (see Recipes) to bring the total volume to 89 μL.
Click reaction
Prepare the click reaction cocktail by adding the following components sequentially into an Eppendorf tube: 10 μL of 5 mM azido-rhodamine in DMSO, 25 μL of 2 mM TBTA in DMSO/t-BuOH, 10 μL of 50 mM TCEP in H2O, and 10 μL of 50 mM CuSO4·5H2O in H2O. Vortex the mixture thoroughly.
Notes:
Prepare TCEP and CuSO4·5H2O solutions freshly.
Azido-rhodamine should be protected from light.
Add 11 μL of the click reaction cocktail to each lysate sample prepared in step A of part II. Vortex to mix thoroughly.
Incubate the reaction mixture in the dark at room temperature for 2 h.
Note: A 2-h incubation time ensures the click reaction is as thorough as possible; however, the duration can be adjusted based on specific experimental needs, such as reducing it to 1 h.
Protein precipitation and washing
Terminate the click reaction by adding 500 μL of ice-cold methanol. Store the mixture at -20 °C overnight.
Note: Methanol should be ice-cold to ensure efficient protein precipitation.
Centrifuge the mixture at 20,000× g for 20 min at 4 °C.
Discard the supernatant carefully and retain the protein pellet.
Note: Remove the supernatant cautiously to avoid disturbing the protein pellet.
Wash the protein pellets twice by adding 1 mL of ice-cold methanol, gently inverting the tube, and centrifuging at 20,000× g for 20 min at 4 °C. The protein pellet should be at the bottom of the tube.
Air-dry the protein pellets on the bench for 60 min at room temperature.
Note: Ensure that pellets are completely dry before proceeding to subsequent steps.
Sample preparation for SDS-PAGE
Resuspend the protein pellet in 57.5 μL of 4% SDS buffer (see Recipes). Shake the samples for 15 min at 1,500 rpm to ensure complete dissolving.
Notes:
Ensure complete dissolution of the protein pellet for subsequent analysis.
Use a bath sonicator to aid solubilization if necessary. Sonication helps to dissolve the protein pellets.
Add 12.5 μL of 2 M NH2OH to each sample.
Optional: NH2OH selectively cleaves thioester bonds but not amide bonds, so treatment with NH2OH can minimize interference from S-palmitoylated proteins. If NH2OH treatment is not needed, add 70 μL of 4% SDS buffer (see Recipes) directly to dissolve the protein pellet in the previous step.
Add 25 μL of 4× SDS-PAGE loading buffer (see Recipes) and 5 μL of bond-breaker TCEP solution.
Heat the samples at 95 °C for 5 min to denature the proteins. Briefly vortex and then centrifuge the samples for 1 min at 10,000× g at room temperature.
SDS-PAGE and in-gel fluorescence visualization
Load 20 μg of protein per lane onto a 4%–20% Bis-Tris gel. Run the gel for 1 h at 140 V.
Note: Use a fluorescent protein ladder as the molecular weight standard to facilitate accurate size determination during fluorescence scanning.
Destain the gel by rocking it for 1 h in destaining buffer (see Recipes) at room temperature.
Note: The destaining step removes the remaining traces of the loading buffer and unreacted rhodamine.
Scan the gel on a ChemiDoc MP imager using “rhodamine” mode.
Following fluorescence scanning, stain the gel with Coomassie Brilliant Blue staining to confirm equal protein loading of samples.
Notes:
Coomassie Brilliant Blue staining provides a visual check for total protein amount and loading consistency.
As an example, the result for in-gel fluorescence analysis of fatty-acylated proteins using metabolic labeling with the bioorthogonal chemical reporter alk-16 and click reaction is shown in Figure 2A.
Figure 2. Results for in-gel fluorescence analysis of alk-16-labeled fatty-acylated proteins. (A) In-gel fluorescence analysis of alk-16-labeled fatty-acylated proteins in HeLa cells. Coomassie Brilliant Blue (CBB) staining is shown to confirm equal protein loading. (B) Validation of fatty-acylation of Ras-related protein R-Ras (RRAS) by in-gel fluorescence analysis. HEK293T cells were transfected with HA-tagged RRAS. The cells were metabolically labeled with alk-16 and subjected to immunoprecipitation and in-gel fluorescence assay. Anti-HA immunoblotting is shown to confirm sample loading.
Part III: Metabolic labeling of the protein of interest with alkynyl chemical reporters in living cells
Cell culture
Seed cells in 60 mm dishes with normal growth medium as described in step A of part I.
Transfection
Prepare transfection mixture. For cells cultured in a 60 mm dish:
In a microcentrifuge tube, dilute a total of 4 μg of plasmid encoding the WT or mutant POI in 400 μL of Opti-MEM.
Add 10 μL of ViaFect transfection reagent to the diluted DNA.
Mix the solution gently by pipetting and incubate the mixture at room temperature for 15 min.
Note: In this study, HEK293T cells were transfected with HA-tagged Ras-related protein R-Ras (RRAS) to validate the fatty-acylation of RRAS by in-gel fluorescence assay.
Add the transfection mixture dropwise into the cell media.
Incubate the cells for 6 h at 37 °C in a CO incubator.
Notes:
The transfection procedure is essential for overexpressing the POI with an affinity tag. In this study, HEK293T cells were transfected with HA-tagged RRAS to validate the fatty-acylation of RRAS by in-gel fluorescence assay.
In initial experiments, it is crucial to check transfection efficiency and ensure the effective expression of the POI. If the POI is fused with a fluorescent protein tag, such as GFP or RFP, use fluorescence microscopy to assess the transfection efficiency and expression level of the POI. If the POI does not have a fluorescent tag, perform western blotting to evaluate the protein expression level.
Metabolic labeling with the chemical reporter
Metabolic labeling of proteins with chemical reporters in living cells and harvest cells as described in Step C of part I.
Part IV: In-gel fluorescence analysis of the protein of interest
Cell lysis and protein quantification
Lyse the cells labeled with chemical reporters by adding 200 μL of RIPA buffer (see Recipes) supplemented with EDTA-free protease inhibitor cocktail and 0.4 μL of benzonase (100 U). Vortex vigorously and then rotate at 4 °C for 30 min.
Note: Other lysis buffers compatible with immunoprecipitation may be used in place of RIPA buffer.
Centrifuge the lysates at 12,000× g for 20 min at 4 °C. Collect the supernatant and discard the pellet (cell debris).
Quantify protein concentration using a standard BCA assay.
Aliquot equal amounts of protein (e.g., 600 μg) into 2.0 mL dolphin microcentrifuge tubes and adjust the volume with the RIPA buffer to 1.5 mg/mL.
Immunoprecipitation
For the immunoprecipitation of proteins tagged with HA/GFP/FLAG, use the corresponding anti-HA/GFP/FLAG agarose beads. Equilibrate the beads by washing them with PBS before use.
Add the agarose beads (e.g., 20 μL of bead slurry per 600 μg of cell lysate) to the lysate and incubate the mixture on a rotator at 4 °C for 2 h. Gentle rotation is essential for efficient binding of the tagged protein to the beads.
After incubation, centrifuge the beads–lysate mixture at 8,000× g for 1 min at 4 °C to pellet the beads. Carefully remove the supernatant without disturbing the bead pellet.
Wash the beads three times with 1 mL of chilled RIPA buffer to remove non-specifically bound proteins. For each wash, gently rotate the beads on a rotator at 4 °C for 5 min, then centrifuge at 8,000× g for 1 min at 4 °C.
After washing with RIPA buffer, wash the beads three additional times with 1 mL of chilled PBS.
Resuspend the beads in 44.5 μL of PBS.
Click reaction
Add 5.5 μL of freshly prepared click reaction cocktail (as described in step B1 of part II) to the resuspended beads.
Incubate the reaction mixture in the dark for 2 h at room temperature.
Wash the agarose beads three times with 1 mL of RIPA buffer.
Sample preparation for SDS-PAGE
Resuspend the beads with 17.25 μL of ultrapure water, 7.5 μL of 4× SDS-PAGE loading buffer, and 1.5 μL of bond-breaker TCEP solution.
Add 3.75 μL of 2 M NH2OH to each sample.
Optional: If NH2OH treatment is not required, add 21 μL of ultrapure water directly in the previous step.
Heat the samples at 95 °C for 5 min to denature the proteins. Briefly vortex and then centrifuge the samples for 1 min at 8,000× g at room temperature to pellet agarose beads.
SDS-PAGE and in-gel fluorescence visualization
Load 20 μL of the supernatant per lane onto a 4%–20% Bis-Tris gel for SDS-PAGE and run at 140 V for 1 h. This gel is used for fluorescent gel scanning.
Add 20 μL of 1× SDS-PAGE loading buffer to the remaining sample, vortex, and centrifuge the samples at 8,000× g for 1 min at room temperature. Load 20 μL of the supernatant per lane onto another 4%–20% Bis-Tris gel for SDS-PAGE and run at 140 V for 1 h. This gel serves as a control to confirm protein loading through western blotting analysis.
After running, destain the first gel and perform fluorescence scanning and Coomassie Brilliant Blue staining as described in steps E2–4 of part II.
Transfer the proteins from the second gel onto a nitrocellulose membrane and detect the POI using standard western blotting procedures.
Note: As an example, the result for in-gel fluorescence analysis of fatty-acylation of HA-tagged RRAS is shown in Figure 2B.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
• Xu et al. [7]. Chemical Proteomics Reveals Nε-Fatty-Acylation of Septins by Rho Inactivation Domain (RID) of the Vibrio MARTX Toxin to Alter Septin Localization and Organization. Mol Cell Proteomic.
Rho inactivation domain (RID) from Vibrio cholerae and Vibrio vulnificus efficiently catalyzes the covalent attachment of long-chain fatty acyl groups (e.g., palmitoyl, stearyl) to the ϵ-amino group of lysine residues on substrate proteins in host cells [7, 10] (Figure 3A). To globally profile RID-mediated Nϵ-fatty acylation, HeLa cells were transfected with either wild-type RID (RID-WT) or the inactive mutant RID-C2835A (RID-CA), incubated with alk-16 overnight, and then lysed. Proteins in the lysates were subjected to click chemistry with azido-rhodamine for in-gel fluorescence analysis, as described in parts I and II (Figure 3B). To reduce interference from S-palmitoylated proteins, lysates were treated with NH2OH to selectively cleave thioester bonds. The in-gel fluorescence revealed specific alk-16 labeling of many proteins in RID-WT lysates, but not in RID-CA lysates (Figure 3C), indicating that RID mediates Nϵ-fatty-acylation of these host proteins in living cells.
Figure 3. Workflow and result for in-gel fluorescence analysis of alk-16-labeled fatty-acylated proteins. (A) RID of the Vibrio MARTX toxin catalyzes Nϵ-fatty-acylation modifications on lysines of its substrate proteins. (B) Schematic for in-gel fluorescence analysis of RID-mediated Nϵ-fatty-acylation of proteins using metabolic labeling with the bioorthogonal chemical reporter alk-16 and click reaction. (C) In-gel fluorescence analysis of alk-16-labeled fatty-acylated proteins in mock- and RID (WT or CA)-transfected HeLa cells. Coomassie Brilliant Blue (CBB) staining is shown to confirm equal protein loading.
To further validate that RID mediates the Nϵ-fatty-acylation of Ras-related protein Rac1 (RAC1), an in-gel fluorescence assay was performed. Specifically, HEK293T cells were co-transfected to express RID and HA-tagged RAC1, followed by labeling with alk-16. Cell lysates were subjected to immunoprecipitation, click-reacted with azido-rhodamine, and analyzed by in-gel fluorescence after NH2OH treatment, as described in parts III and IV (Figure 4A). The results showed that RAC1 was labeled by Alk-16 only in the presence of RID-WT, but not upon expression of RID-CA (Figure 4B), confirming that RID catalyzes the Nϵ-fatty acylation of RAC1.
Figure 4. Workflow and result for in-gel fluorescence analysis of RID-mediated Nϵ-fatty-acylation of HA-tagged RAC1. (A) Workflow for analysis of RID-mediated Nϵ-fatty-acylation of candidate proteins of interest (POI). HEK293T cells were co-transfected with RID and individual tagged POI, metabolically labeled with alk-16, and lysed. The cell lysates were subjected to immunoprecipitation, click reaction with azido-rhodamine, NH2OH treatment to cleave thioester bonds, and in-gel fluorescence analysis. (B) Validation of RID-mediated Nϵ-fatty-acylation of RAC1 by in-gel fluorescence analysis. Samples were prepared as in (A). Anti-HA immunoblotting is shown to confirm sample loading.
General notes and troubleshooting
Troubleshooting
Cells unhealthy after transfection (step B of parts I and III)
Ensure cell confluency is around 70% before transfection.
Adjust the transfection reagent-to-plasmid ratio and consider shortening the transfection time.
Replace the media 6 h after transfection.
Check if overexpression of the POI is toxic to the cells.
Cells detaching during media change (step C1 of parts I and III)
Gently aspirate and replace the media, avoiding direct pipetting onto the cells.
Slowly add media near the edge of the dish to minimize disruption to the cell monolayer.
Consider using an electronic pipette aid with a gravity drain feature.
Using poly-D-lysine-coated culture dishes may improve cell attachment.
Click chemistry reaction not proceeding efficiently (step B of part II and step C of part IV)
Always use freshly prepared TCEP and CuSO4 solutions, as TCEP may oxidize over time and Cu(I) is critical for the reaction. Prepare these solutions just before use to ensure optimal reaction efficiency.
Poor protein precipitation after the click reaction (step C of part II)
Ensure that methanol is ice-cold and has been stored at -20 °C overnight before centrifugation.
Increase methanol volume if needed to improve precipitation.
Incomplete resolubilization of the protein pellet (step D1 of part II)
Ensure the protein pellet is completely dry before dissolving.
Use 4% SDS buffer combined with sonication to fully resolubilize the protein.
High background fluorescence in fluorescence gel scans (step E3 of parts II and IV)
Wash the protein pellet twice with ice-cold methanol during the precipitation step to remove unreacted azido-rhodamine.
Thoroughly destain the gel after SDS-PAGE to eliminate residual fluorescence.
Handle the gel by its edges with clean gloves to avoid smudges or fingerprints.
Failure to detect the protein of interest in immunoprecipitation (step B of part IV)
If low protein expression levels are suspected or epitope loss occurs during lysis, verify protein expression on a small sample before immunoprecipitation.
If the protein is localized in the nucleus or detergent-resistant membranes (DRMs), using 1% SDS lysis buffer may be necessary to lyse the cells. Dilute SDS to 0.1% before incubating with beads.
For inefficient binding, extend the incubation time and increase the amount of beads used.
If protein loss occurs during washing, use gentler washing conditions and avoid vigorous vortexing of the beads.
Acknowledgments
We thank Peng lab members for the helpful discussion. T.P. acknowledges grant support from National Natural Science Foundation of China (22277008).
Author Contributions
Conceptualization, T. P.; Investigation, X. Y. Writing – Original Draft, X. Y.; Writing – Review & Editing, T. P.; Funding Acquisition, T. P.
Competing interests
The author declares no competing interests.
References
Prescher, J. A. and Bertozzi, C. R. (2005). Chemistry in living systems. Nat Chem Biol. 1(1): 13–21.
Demko, Z. P. and Sharpless, K. B. (2002). A click chemistry approach to tetrazoles by Huisgen 1,3-dipolar cycloaddition: synthesis of 5-acyltetrazoles from azides and acyl cyanides. Angew Chem, Int Ed Engl. 41(12): 2113–2116.
Wang, Q., Chan, T. R., Hilgraf, R., Fokin, V. V., Sharpless, K. B. and Finn, M. G. (2003). Bioconjugation by copper(I)-catalyzed azide-alkyne[3 + 2] cycloaddition. J Am Chem Soc. 125(11): 3192–3193.
Grammel, M. and Hang, H. C. (2013). Chemical reporters for biological discovery. Nat Chem Biol. 9(8): 475–484.
Charron, G., Zhang, M. M., Yount, J. S., Wilson, J., Raghavan, A. S., Shamir, E. and Hang, H. C. (2009). Robust fluorescent detection of protein fatty-acylation with chemical reporters. J Am Chem Soc. 131(13): 4967–4975.
Liu, W., Zhou, Y., Peng, T., Zhou, P., Ding, X., Li, Z., Zhong, H., Xu, Y., Chen, S., Hang, H. C. and Shao, F. (2018). N ϵ-fatty acylation of multiple membrane-associated proteins by Shigella IcsB effector to modulate host function. Nat Microbiol. 3(9): 996–1009.
Xu, Y., Ding, K. and Peng, T. (2024). Chemical proteomics reveals Nϵ-fatty-acylation of septins by Rho inactivation domain(RID) of the Vibrio MARTX toxin to alter septin localization and organization. Mol Cell Proteomics. 23(3): 100730–100753.
Sun, Y., Chen, Y. and Peng, T. (2022). A bioorthogonal chemical reporter for the detection and identification of protein lactylation. Chem Sci. 13(20): 6019–6027.
Bao, X., Xiong, Y., Li, X. and Li, X. D. (2018). A chemical reporter facilitates the detection and identification of lysine HMGylation on histones. Chem Sci. 9(40): 7797–7801.
Zhou, Y., Huang, C., Yin, L., Wan, M., Wang, X., Li, L., Liu, Y., Wang, Z., Fu, P., Zhang, N., Chen, S., Liu, X., Shao, F. and Zhu, Y. (2017). N ϵ-fatty acylation of Rho GTPases by a MARTX toxin effector. Science. 358(6362): 528–531.
Article Information
Publication history
Received: Jul 28, 2024
Accepted: Sep 18, 2024
Available online: Oct 17, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Category
Biochemistry > Protein > Fluorescence
Biochemistry > Protein > Posttranslational modification
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A Simple Staining Method Using Pyronin Y for Laser Scanning Confocal Microscopy to Evaluate Gelatin Cryogels
BR Brianna Reece
EB Elizabeth V. Bahar
AP Angel Cabrera Pereira
LW Lukasz Witek
KK Katsuhiro Kita
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5115 Views: 452
Reviewed by: Hélène LégerHsih-Yin Tan Anonymous reviewer(s)
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Abstract
This study explores the novel application of pyronin Y for fluorescently labeling extracellular matrices (ECMs) and gelatin cryogels, providing a simple and reliable method for laser scanning confocal microscopy. Pyronin Y exhibited remarkable staining ability of the porous structures of gelatin cryogels, indicating its potential as a reliable tool for evaluating such biomaterials. Confocal imaging of pyronin Y–stained cryogels produced high signal-to-noise ratio images suitable for quantifying pores using Fiji/Image J. Importantly, pyronin Y enabled effective dual-color imaging of cryogel-labeled mesenchymal stem cells, expanding its utility beyond traditional RNA assays. Traditional staining methods like Mason’s trichrome and Sirius Red have limitations in cryogel applications. Pyronin Y emerges as a powerful alternative due to its water solubility, minimal toxicity, and stability. Our results demonstrate pyronin Y’s ability to specifically stain gelatin cryogel's porous structures, surpassing its weak staining of ECMs in 2D. Confocal imaging revealed enduring staining even under rigorous scanning, with no notable photobleaching observed. Furthermore, pyronin Y's combination with Alexa Fluor 647 for dual-color imaging showed promising results, validating its versatility in fluorescence microscopy. In conclusion, this study establishes pyronin Y as a cost-effective and rapid option for fluorescent staining of gelatin cryogels. Its simplicity, efficacy, and compatibility with confocal microscopy make it a valuable tool for characterizing and evaluating gelatin-based biomaterials, contributing significantly to the field of cryogel imaging. The study opens new avenues for dual-color imaging in biomaterial research and tissue engineering, advancing our understanding of cellular interactions within scaffolds.
Key features
• Fluorescent staining of gelatin-based cryogels with an inexpensive yet less time-consuming protocol.
• Pyronin Y staining is suitable for dual-color confocal imaging by combining with far-red fluorophores (such as Alexa Fluor 647).
• The method is conducted routinely.
Keywords: Pyronin Y Cryogels Collagen Gelatin Glutaraldehyde Fluorescence labeling
Graphical overview
Gelatin cryogel staining using pyronin Y
Background
In this study, we report the novel application of pyronin Y dye in cryogel imaging combined with confocal microscopy, focusing on the detailed visualization of the cryogel’s porous structure. To our knowledge, this is the first protocol demonstrating a very fast yet reliable fluorescent staining of gelatin-based cryogels. We also show its compatibility with dual-color imaging.
The lack of a straightforward, rapid, and reliable fluorescent labeling method for cryogels is one obstacle in the field. Traditional methods like Mason’s trichrome for collagen [1] and the anionic dye, Sirius Red, for polarized microscopy [2] have limitations in cryogel applications. Mason’s trichrome staining is optimized to stain collagen fibers of tissue sections, not biomaterials [3]. While immunofluorescent staining of cryogels is possible using anti-gelatin antibodies, it is cost-prohibitive and not used for cryogels. Covalent conjugation of fluorescent dyes to gelatin is effective in visualizing the architecture of gelatin-based cryogels [4]; however, it is time-consuming.
Initially, we evaluated the dye’s potential for staining extracellular matrix (ECM)-coated glass surfaces in 2D cultures and discovered that pyronin Y can specifically stain 3D porous structures. Traditionally, pyronin Y has been a staple in RNA analysis [5–7]. Cryogels are unique porous scaffolds formed at sub-zero temperatures (around -20 °C) through a freeze-thaw process resulting in the formation of a porous network of polymer strands, post-thawing, due to the sublimation of ice crystals [8,9]. The interconnected porosity of cryogels allows efficient diffusion of nutrients and oxygen, rendering them advantageous for a variety of applications in tissue engineering, including superior cell growth [10].
Pyronin Y (C17H19CIN2O) features a dimethylamino group [N(CH3)2] and a xanthene-3-iminium group, which render the dye its cationic nature and distinctive red fluorescence. Additionally, pyronin Y is water-soluble and therefore easy to apply to various biological applications. Furthermore, it is considered to be safe for use in biological research, due to its relatively low toxicity. Lastly, pyronin Y is a stable dye that can be stored for extended periods without significant degradation or loss of fluorescence intensity. Past studies [5,7,11] have demonstrated pyronin Y’s efficacy in RNA analysis, owing to its ability to emit fluorescent light upon binding to RNA through ionic interactions. Thus, pyronin Y is established as a crucial component in a variety of RNA-based assays, such as in situ hybridization, [11], flow cytometry [5], and real-time RNA imaging [7]. However, no studies explored pyronin Y's potential for the imaging of ECMs, as shown here.
The properties of pyronin Y, such as its vivid red fluorescence, positive charge, water solubility, minimal toxicity, and chemical stability, render it a highly adaptable dye for the analysis of various extracellular matrices. Gelatin, which primarily constitutes heterogeneous peptides, is a predominantly negatively charged protein in alkaline pH [10]. This protocol explores the efficacy of pyronin Y in staining gelatin-based cryogels.
Materials and reagents
Biological materials
StemProTM BM mesenchymal stem cell (Gibco, Thermo Fisher Scientific, catalog number: A15652)
Note: Other mesenchymal stem cells (MSCs) may be used, such as those isolated from the umbilical cord [12] or from other sources.
Reagents
Pyronin Y (Alfa Aesar, Brand, catalog number: J61068, lot F18Z037), stable at room temperature
Glutaraldehyde 25% solution (Sigma-Aldrich, catalog number: G6257); once opened, you may store it in a refrigerator, tightly sealing the container
Sodium borohydride (Sigma-Aldrich, catalog number: 452882)
Sodium carbonate (Fisher Scientific, catalog number: S64-500) (any equivalent or higher quality of the same chemicals should work fine)
Hydrochloric acid (for pH adjustment) (Fisher Scientific, catalog number: A1445-500) (any equivalent or higher quality of the same chemicals should work fine)
16% paraformaldehyde (Electron Microscopic Sciences, catalog number: 15700); once opened, you may store it in a refrigerator, tightly sealing the container
Triton X-100 (Sigma-Aldrich, catalog number: T8787)
Fish skin gelatin (Sigma-Aldrich, catalog number: G7041)
Alexa Fluor 647 phalloidin (Invitrogen, Thermo Fisher Scientific, catalog number: A22287)
Dulbecco’s modified Eagle medium (Gibco, Thermo Fisher Scientific, catalog number: 11995), store at 4 °C
Fetal bovine serum (premium) (Gibco, Thermo Fisher Scientific, catalog number: A5670701)
Dulbecco’s phosphate-buffered saline (PBS) without calcium and magnesium (Gibco, Thermo Fisher Scientific, catalog number: 14190144)
Solutions
Gelatin cryogel solution (see Recipes)
0.1 M sodium bicarbonate buffer (see Recipes)
Pyronin Y solution (see Recipes)
4% Paraformaldehyde solution (see Recipes)
0.5% Triton X-100 solution (see Recipes)
Recipes
Gelatin cryogel solution
Reagent Final concentration Amount (per 1.5 mm thick gel)
Gelatin (8.6%) 4.3% 3,000 μL
Glutaraldehyde (25%) * 0.3% 72 μL
H2O n/a 2,928 μL
Total n/a 6,000 μL
*Add after mixing other parts.
0.1 M sodium carbonate buffer (pH 9.2)
Reagent Final concentration Amount
Sodium carbonate 0.1 M 10.6 g (when anhydrous is used)
HCl n/a (add dropwise to adjust the pH)
Total with H2O n/a 1,000 mL
Pyronin Y solution
Reagent Final concentration Amount
Pyronin Y 1 mg/mL 1 mg
ddH2O n/a 1 mL
Total n/a 1 mL
It is highly recommended to spin briefly (~30 s) at 10,000× g to remove any insoluble pyronin Y particles after dissolving the pyronin Y powder.
4% paraformaldehyde solution
Reagent Final concentration Amount
16% paraformaldehyde solution 4% 2.5 mL
PBS n/a 7.5 mL
Total n/a 10 mL
You can downscale the volume.
0.1% Triton X-100 solution
Reagent Final concentration Amount
Triton X-100 0.1% 10 μL
PBS n/a 9,990 μL
Total n/a 10 mL
You can downscale the volume. It will be fast to dissolve Triton X-100 by using a magnetic stirrer bar in a small beaker. Alternatively, making a 10% Triton X-100 stock solution helps this process. If a 10% stock solution is used, the volume of Triton X-100 should be 100 μL.
Laboratory supplies
1.5 mL or 2.0 mL microcentrifuge tubes
Micropipette (P200 and P1000)
1.5 mL tube rack
15 mL tubes
100 mm plastic Petri dishes or square dishes (e.g., Carolina Biological Supply, catalog number: 741470)
Gloves
Flat-tip forceps
35 mm glass bottom dishes (Mattek, catalog number: P35G-1.5-7-C)
Note: As long as the thickness is #1.5, any glass-bottom dishes for cellular imaging would be suitable.
1.5 mm spacer glass plates (Bio-Rad, catalog number: 1653312)
Short glass plates (Bio-Rad, catalog number: 1653308)
Mini-PROTEAN tetra cell casting stand & clamps (Bio-Rad, catalog number: 1658050)
Mini-PROTEAN casting stand gasket (Bio-Rad, catalog number: 1653305)
(alternative for 10/11) glass tube (the bottom has to be tightly sealed with a removable plug) or plastic disposable syringes (no needle required; the top has to be cut off)
Equipment
LSM 880 (Carl Zeiss)
20× objective lens; a high numerical aperture, longer working distance objective lens is strongly recommended, such as Plan-Apochromat 20×/N.A. = 0.8 M27 (Carl Zeiss, catalog number: 420650-9901-000)
-20 °C freezer (Thermo Fisher Scientific)
UV crosslinker (such as Spectronics Corporation, catalog number: XL-1000) or a cell culture hood equipped with a UV lamp
Software and datasets
Fiji/Image J (2.15.1), available without any cost
Microsoft Excel (no specific version as long as Office 2003 or newer versions are used)
Fluorescence Spectraviewer (Bio-Rad; https://www.bio-rad-antibodies.com/spectraviewer.html), available without any costs. Only required if you want to confirm the compatibility of your microscopic illumination settings.
Procedure
Preparation of gelatin cryogels
Set up the gel casting apparatus
Use either a protein gel apparatus (to prepare a sheet-type cryogel), glass tubes, or small syringes (to prepare a tube-type cryogel).
Make sure to measure the volume of liquid that your casting can hold (depending on this volume, you may scale up/down the total volume of the cryogel) (Recipe 1). Mix water, gelatin, and glutaraldehyde in an appropriate tube, such as a 15 mL plastic tube.
Critical: Add glutaraldehyde solution after gently mixing water and gelatin, because chemical crosslinking will start soon after adding glutaraldehyde. After the addition of glutaraldehyde, gently mix the solution again.
Pour the mixed solution into an apparatus. Leave on the bench (room temperature) for 30 min.
Carefully place an apparatus in a -20 °C freezer. Leave in a freezer overnight (~16 h).
Critical: Freeze-thawing is the critical step for the formation of porous structure. Make sure that other people do not open the freezer door (especially during the initial freezing process).
Taking out cryogels
Take out a gel apparatus from the -20 °C freezer and leave it on a bench until completely thawed.
Critical: If a cryogel is not completely thawed, gels or pore structures may be damaged during the process of retrieving a cryogel from an apparatus. See General note 1.
Set up a dish (such as a 100 mm Petri dish or a rectangular plastic dish) filled with 0.1 M sodium carbonate buffer (Recipe 2).
Carefully separate glass plates or slowly push the gel (if a glass tube or a syringe is used).
Rinse cryogels with deionized water. Then, transfer cryogels to a dish filled with 0.1 M sodium carbonate buffer.
Add a small pinch of sodium borohydride powder to the solution. One previous study indicates 0.1 M freshly prepared sodium borohydride solution [4]. This is equivalent to the concentration of 3.78 mg/mL. You will start seeing bubbles (sodium borohydride reacts with remaining glutaraldehyde and producing hydrogen). To quench enough, you may leave cryogels in the sodium borohydride solution overnight on a rocking shaker (Pause point). Cryogels can be kept in sodium carbonate buffer with or without sodium borohydride. However, we recommend starting the quenching with sodium borohydride soon after taking out gels from the casting apparatus, as it will take at least overnight to quench residual sodium borohydride well. Remaining sodium borohydride can cause cell death if you grow cells.
You can keep cryogels in 0.1 M sodium carbonate buffer. Keep it in a refrigerator (Pause point). Rinse well with deionized water before use.
Staining of cryogels
Cutting cryogels
Use a razor blade to slice a piece of cryogel. Make a small rectangular piece (approximately 5–10 mm is sufficient)
Transfer a piece of cryogel onto the glass surface of a glass-bottom dish.
(Not necessary: We conducted this to compare pyronin Y staining in 2D and 3D) To stain ECM coating, the surface of a glass bottom dish should be coated with 1 mg/mL of ECM solution first. Then, the same staining process (step B2) needs to be done.
Pyronin Y staining
Critical: If MSCs are included, go to section E, then come back here after Alexa Fluor 647-phalloidin staining of MSCs.
Place a piece of cryogel from the previous step onto a glass bottom dish (#1.5 thickness).
Critical: The thickness of coverslips is critical to acquiring microscopic images with the optimum conditions.
Add the pyronin Y solution (Recipe 3) to cover the entire gel. Although the volume will depend on the size of a piece of cryogel, typically 50–200 μL of the pyronin Y solution is sufficient.
Leave at room temperature for 5 min.
Rinse the stained cryogel with deionized water three times (5 min each). If over-staining happens, try again with a shorter staining time (see step C1h and General note 5).
Imaging of cryogels with a laser-scanning confocal microscope*
*Many people may use their core imaging facility. It may be easy for you to consult with your facility manager/director.
Critical: Please let them know that the settings described in steps C1c–d are critical, as pyronin Y’s imaging probably requires less laser power and relatively low digital gain; see General note 2.
Determine the acquisition settings
Place a 35 mm glass-bottom dish with a piece of the stained cryogel onto the stage of a confocal microscope.
Use an appropriate objective lens (see General note 2).
Set the lowest laser power that is necessary to obtain sufficient signals. With a Zeiss 880 equipped with a highly sensitive Gallium-Arsenide-Phosphide (GaAsP) detector, ~0.1% was sufficient for the excitation of pyronin Y (with a 561 nm laser). When MSCs (stained with Alexa Fluor 647-phalloidin) are also included for dual-color imaging, a slightly higher laser power for a 633 nm laser may be needed (see General note 3).
Select the laser for pyronin Y excitation (such as 561 nm).
Set the scanning area, digital zoom, and scanning speed (important; see General note 4). We used the highest scanning speed (8; resulting in a resolution of 0.415 μm). The scanning area is up to your choice. Because the magnification is low in this case (20×), we set 1,024 × 1,024. 512 × 512 may be too small.
Set the master gain. Probably ~500 is appropriate. (Critical: Too high master gain can damage the detector.) Because pyronin Y staining of gelatin cryogels gives fairly high signals, we strongly suggest keeping the digital gain at 1.0 and digital offset at 0.
Set the pinhole diameter. In Zeiss confocal operated with Zen, you simply need to hit 1 AU, which will set the optimum pinhole diameter (with a 561 nm excitation wavelength, it is 36 μm).
Hit Live to check that the setting is suitable for imaging. Keep your eye on the histogram of the signal to make sure that the signal is not saturated (see General note 5).
If you conduct dual-color imaging with MSCs stained with Alexa Fluor 647-phalloidin, repeat steps C1f–h with the laser for the excitation of Alexa Fluor 647-phalloidin. [In our case, we used a 633 nm laser; an excitation wavelength closer to 647 nm is fine (better) if available.] Also, carefully check what MSCs look like (see Troubleshooting).
Acquire images
In the control panel, first set live to determine the start and the end plane.
When you set the z-axis step, a 1.0 μm interval may be sufficient, because a relatively low magnification of an objective lens is used.
Then, hit the start experiment button. The following YouTube video may be helpful: https://www.youtube.com/watch?v=QhFb6vU1Iyg.
Save z-stack images. Keep the original data files without modification (important).
Quantification of pores using Image J/Fiji [13]
Export files from the original z-stack data files
Open the original data files with the software to control the microscope and export them to create TIFF files (see General note 6). If Zeiss confocal is used, the original files can be opened with Fiji.
Adjust the contrast if necessary (see General note 7).
Quantification of pores (see General note 8)
Copy the selected z-plane image from the stack (Image → Duplicate). When a small window pops up, type the desired single-plane range (for example, 8-8).
Set the threshold (Image → Adjust → Threshold).
The following YouTube video may be very useful: https://www.youtube.com/watch?v=4gm9Q_TdRvY.
MSC culture
Rinsing cryogels
Rinse a piece of cryogel with deionized water extensively.
After rinsing, dip a piece of gel in 70% ethanol, then place it on a 35 mm dish (or a 12-well plate for more convenient handling). Then, place a dish/plate in a UV crosslinker for approximately 2 min. If a UV crosslinker is not available, placing it under a cell culture hood for approximately 5–10 min (then turning on the UV lamp) can be an alternative.
Caution: Do not overdo this process, especially with a UV crosslinker. It can dry the gel too much and cause shrinking. Also, cryogels should be handled aseptically after this process. Contamination of microorganisms has to be avoided. We used a UV crosslinker only to sterilize cryogels. Chemical crosslinking is not the purpose of this procedure; therefore, sterilization under cell culture hoods with UV lamps will also work fine.
MSC culture and staining
Slowly seed 10 μL of trypsinized MSCs suspension (~400,000 cells/mL) onto a piece of cryogel in a 12-well tissue culture plate to absorb the cell suspension.
Put the plate back in a CO2 incubator and leave it for 30 min to allow MSCs to adhere to a cryogel piece.
Caution: If your gel is too dry, you may want to add a little drop of cell culture medium after the previous step to avoid cells dying due to drying.
Add 1 mL of culture medium.
Aspirate the medium a few hours later and fix cells with 4% paraformaldehyde in PBS for 20 min.
Permeabilize the cell membrane with 0.1% Triton X-100 in PBS (5 min).
Rinse with PBS three times (5 min each).
Stain F-actin with Alexa Fluor 647-phalloidin (1/100) in PBS for 15–30 min (see General note 9).
Rinse with PBS three times (5 min each).
Conduct the Pyronin Y staining (as described in B2; also see General note 10).
Data analysis
Although no special computational skills are needed, some try-and-error type patience is required if you try a semi-automated quantification of pores. The video mentioned in step C2c is useful.
Validation of protocol
In order to test the specificity of pyronin Y dye to different ECMs, pyronin Y was first applied to stain the various ECMs in 2D. Glass-bottom dishes were coated with each ECM protein and then incubated with pyronin Y solution. The comparison of the average maximum fluorescence intensity showed minimal staining of the ECMs coating of the glass surface (Figure S1). The fluorescence intensity, however, yielded a slightly higher signal level in gelatin- and collagen-coated dishes. This result implied that pyronin Y may potentially recognize a specific structure present in those proteins, such as glycine-proline-hydroxyproline triplets. Remarkably, gelatin cryogels showed approximately 8-fold higher average signals than gelatin or collagen coating (Figure S1). Indeed, pyronin Y staining of a gelatin cryogel resulted in exceptionally clear visualization of porous structures (Figure 1).
Figure 1. Examination of pyronin Y staining in three channels. No staining (top) and pyronin Y staining (bottom) of cryogels imaged with three different laser lines (488, 561, and 633 nm). Scale bar = 50 μm.
Through threshold adjustment, followed by conversion to a black-and-white format and inversion of the binary image, the production of a clear image of the mesh-like pores (Figure 2A) was possible. Using the object counting function in Fiji software allowed the semi-automatic outlining and quantification of pore sizes (section C).
Employing a 561 nm laser resulted in more than 50% of the peak emission of pyronin Y, which has an optimum excitation peak of around 544 nm (Figure S2). In contrast, a 488 nm laser, though less efficient, still presented approximately 15% of the maximum emission. Consequently, some degree of bleed-through was expected by combining pyronin Y with green fluorescent dyes for dual-color imaging. However, at 633 nm, pyronin Y is not expected to emit any fluorescence. This theoretical bleed-through was experimentally validated by imaging a pyronin Y-stained gelatin cryogel with three laser lines (488, 561, and 633 nm). As presented in Figure 1, excitation of a pyronin Y–stained cryogel with a 488 nm laser yielded a low, yet recognizable level of bleed-through, whereas excitation with a 633 nm laser showed no detectable emission.
Therefore, we hypothesized that combining pyronin Y with a far-red fluorescence dye, such as Alexa Fluor 647, may be more effective for dual-color imaging. Based on these insights, we conducted dual-color imaging of MSCs and a piece of cryogel. Figure 2B demonstrates that the combination of pyronin Y and Alexa Fluor 647 phalloidin staining enabled successful dual-color confocal imaging of cryogels and MSCs. Although there was a subtle staining inside of MSCs observed in the red channel (Figure 2B), it was believed to be due to the binding of pyronin Y to RNA inside of MSCs. The far-red channel clearly shows distinct actin staining (center, Figure 2B). This compelling evidence reinforces the versatility of pyronin Y as a powerful tool in fluorescence microscopy, particularly for dual-color imaging applications. The successful combination of pyronin Y with Alexa Fluor 647 phalloidin in our experiments not only validates our hypothesis but also sets a new precedent for advanced imaging techniques in cellular and molecular biology.
Figure 2. Examples of semi-automated pore analysis and dual-color imaging of a pyronin Y–stained cryogel with MSCs. (A) An example of a single-plane staining image of a gelatin cryogel processed for quantification. (B) Dual-color imaging of a cryogel and MSCs. red: pyronin Y-stained cryogel; magenta: MSCs stained with Alexa Fluor 647. Scale bar = 50 μm.
General notes and troubleshooting
General notes
Pyronin Y solution is stable at room temperature for at least a few weeks. The solution shows a slightly dark yet vivid color. However, if this color does not appear to be vivid, preparing a fresh pyronin Y solution is necessary. If you have not used the stock solution for a while, we strongly recommend conducting a quick staining test with a piece of gelatin cryogel. You can quickly see if fluorescence is present by checking with a conventional fluorescent microscope equipped with a Texas Red/X-Rhodamine filer. However, a conventional epi-fluorescence microscope gives significantly higher background signals and images will not be suitable to semi-automatically quantify with Fiji/Image J. Therefore, a combination of a confocal microscope and the indicated laser(s) wavelength would give you the best result. If you just want a routine quality check of cryogels, the use of a conventional epi-fluorescence microscope is sufficient. We have not tried imaging pyronin Y–stained cryogels with a spinning disk confocal microscope, although it should not be a problem. Note that the detector(s) used for a spinning disk confocal microscope could be a charge-coupled device (CCD), a complementary metal oxide semiconductor (cMOS), or an electron-multiplying charge-coupled device (EM-CCD) camera. We do not think EM-CCD is necessary for the imaging of pyronin Y staining; however, please note that the sensitivity of CCD and cMOS will be lower than that of the GaAsP detector. You need to adjust (increase) the laser power. To minimize photobleaching, increased exposure time is efficient for acquiring high-contrast images when those cameras with a spinning disk confocal are used. Because magnification is relatively low, binning (such as 2 × 2) is also effective, depending on the pixel size of CCD and cMOS cameras. However, those acquisition settings are beyond the scope of this protocol, and we will omit the discussion.
Please note that the protocol using Fiji/Image J shown here does not allow 3D measurements―such tasks may be done with Imaris or other software (not free).
Despite some possible limitations, we would like to emphasize that pyronin Y staining is a quite reasonably performed, very simple, easy, and yet reproducible fluorescent labeling method for gelatin cryogels.
When cryogels are successfully made, it appears as a uniform, very slightly yellow/tan-colored material. If something is wrong during the freeze-thawing, you may see a half-transparent appearance (Maybe ice crystals, but we do not know exactly.) In such cases, it would be better to start all over again to prepare a new cryogel. After the successful quenching with sodium borohydride, a subtle yellow/tan color of cryogels will disappear and cryogels’ color will change to white (Figure 3).
Figure 3. Cryogels before (left arrow) and after quenching (right arrow) with sodium borohydride
If higher magnifications, such as 60× (63× in Zeiss) and 100×, are used, please keep in mind that the working distance of microscopic objective lenses becomes shorter and thus may not be suitable for pore imaging of cryogels. Because the purpose of this protocol is to stain and evaluate the overall architectures of cryogels, we think that lower magnifications are more appropriate. 10–20× objective lenses with relatively high numerical aperture (N.A.) values work best and, therefore, we used 20× (N.A. = 0.8) objective in this example (N.A. higher than 0.8 is great but such lenses are optimized for multi-photon imaging and thus very expensive). If conventional 10–20× objective lenses are used, Plan Apochromat or Plan Fluor quality lenses are highly preferred. Please note that the use of lower N.A. lenses will result in less signal; thus, acquisition settings would need to be adjusted accordingly (a little more laser power and slightly higher master gain of a GaAsP detector). Also, if you use a laser scanning confocal (either earlier models from Zeiss or other manufacturers’ laser scanning confocal microscopes) without high-sensitive detectors such as a GaAsP detector, you need to increase the laser power (and probably digital gain as well). Traditionally, photomultipliers (PMT) have been used as detectors in laser scanning confocal microscopy. However, its sensitivity is approximately half for pyronin Y’s emission wavelength and ~30% for Alexa Fluor 647’s emission wavelength.
We confirmed that the combination of 488 and 561 nm cannot avoid breed-through because of the excitation spectrum of pyronin Y (see Figure S2). A 633 nm laser was used to excite Alexa Fluor 647-phalloidin because a 633 nm laser is the standard configuration for Zeiss 880. If your microscopic settings allow the use of a laser closer to 647 nm, even better.
Please note that the scan line speed was set at the highest in our case. Although higher scan line speed can be disadvantageous, such as reduced signal-to-noise ratios and spatial resolution, please note that too low scan line speed (= more laser scanning) is not necessary with 20× objective lenses. If you use the formula d = 0.61 λ/N.A., the spatial resolution in our case (using a 20× objective lens with a 561 nm laser) should be 428 nm (0.428 μm). Thus, the maximum scan line speed (again, our case is Zeiss 880 with Zen) gives a sufficient spatial resolution.
Overstaining of cryogels must be avoided because it does not allow quantitative imaging. Too much pyronin Y signals might also bother the dual-color imaging. Overstaining of cryogels can easily occur when pyronin Y staining takes longer than 5 min. It may be necessary to optimize the staining time (may be a little shorter). Saturation of digital signals also occurs by the detector’s settings (the image would be 16-bit, so the maximum signal value in grayscale should be 65,535).
When you export files from the original data, make sure to split the channels if you conduct dual-color imaging (i.e., pyronin Y–stained cryogels and Alexa Fluor 647-phalloidin-labeled MSCs). This can be done with microscopic software (Zen) when you export; alternatively, you can also split the channels after opening the original stack files with Fiji.
Changing the Gamma value is not necessary and not recommended. Although the later procedure using Fiji/Image J will create binary images for semi-automated quantification, note that modifying the Gamma value will introduce the nonlinear fluorescence intensity adjustment in an image [14]. With pyronin Y, gelatin cryogels should be able to give relatively high signal-to-noise ratio images. If not, either staining or acquisition conditions may need to be checked.
Please note that successful quantification relies on image quality. If the signal-to-noise ratio of the original images is not great, semi-automated quantification will become quite difficult.
We did not attempt blocking (of non-specific binding of Alexa Fluor 647-phalloidin to cryogels) with bovine serum albumin, serum, or skim milk. Prolonged incubation of this step may cause significant Alexa Fluor 647-phalloidin binding to cryogels, but we have not tested it. One fear of doing enough blocking is that it might also bother the staining of MSCs. At least, a relatively short incubation time was sufficient for dual-color imaging of pyronin Y–stained cryogels and Alexa Fluor 647-phalloidin-labeled MSCs.
We conducted Alexa Fluor 647-phalloiding staining (MSCs) first, then moved on to the pyronin Y staining of a cryogel. The order could be reversed. However, it is important to keep in mind that 1) the cell membrane needs to be permeabilized with a detergent such as Triton X-100, as shown in this protocol, to stain F-actin, and 2) pyronin Y is not covalently bound to gelatin. So, if pyronin Y staining is done first, you may lose significant pyronin Y staining during Triton X-100 treatment.
Troubleshooting
To culture MSCs, you should rinse cryogels very well before setting the cell culture. Residual buffer or glutaraldehyde from the manufacturing process of cryogels can cause unhappy MSCs (or, in the worst case, cell death). After the quenching of glutaraldehyde with sodium borohydride, cryogels are submerged in 0.1 M sodium carbonate buffer, whose pH is basic. It may be necessary to rinse cryogels with deionized water for more than a day.
Acknowledgments
We would like to thank Dr. Karen Kasza and Christian Cupo (Columbia University) for allowing us to use the Zeiss 880 confocal microscope. The authors also appreciate Dr. Björn Kafsack (Weill Cornell Medicine) for sharing pyronin Y. This study was partially supported by a St. Francis College faculty research grant (K.K.). St. Francis College STEM Resource Center (funded by the Department of Education, STEM Success & Articulation grant P031C210140; for B.R. and E.B.) and Collegiate Science and Technology Entry Program (CSTEP) from the New York State Education Department (CSTEP grant #0537-22-2016; for E.B.) provided support for faculty-mentored research of students. Lastly, we thank Dr. Allen Burdowski (St. Francis College) and Noemi Rivera (St. Francis College STEM Resource Center) for their tireless effort to support student engagement in research. This is an original work and has not been published in any other papers.
Competing interests
There are no conflicts of interest or competing interests.
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Supplementary information
The following supporting information can be downloaded here:
Figure S1. A graph summarizing the average of maximum fluorescence intensity in a field (n = 3–5 pictures)
Figure S2. The comparison of pyronin Y (A) excitation (dotted line) and emission spectra (solid line) data with three laser lines (488, 561, and 633 nm; B–D)
Article Information
Publication history
Received: Jun 17, 2024
Accepted: Sep 23, 2024
Available online: Oct 22, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,116 | https://bio-protocol.org/en/bpdetail?id=5116&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Preparing Chamber Slides With Pressed Collagen for Live Imaging Monolayers of Primary Human Intestinal Stem Cells
JB Joseph Burclaff
SM Scott T. Magness
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5116 Views: 281
Reviewed by: Laxmi Narayan MishraSarah ShortAnu Thomas
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Original Research Article:
The authors used this protocol in Scientific Reports Jul 2024
Abstract
Primary human intestinal stem cells (ISCs) can be cultured and passaged indefinitely as two-dimensional monolayers grown on soft collagen. Culturing ISCs as monolayers enables easy access to the luminal side for chemical treatments and provides a simpler topology for high-resolution imaging compared to cells cultured as three-dimensional organoids. However, the soft collagen required to support primary ISC growth can pose a challenge for live imaging with an inverted microscope, as the collagen creates a steep meniscus when poured into wells. This may lead to uneven growth toward the center of the well, with cells at the edges often extending beyond the working distance of confocal microscopes. We have engineered a 3D-printed collagen mold that enables the preparation of chamber slides with flat, smooth, and reproducible thin collagen layers. These layers are adequate to support ISC growth while being thin enough to optimize live imaging with an inverted microscope. We present methods for constructing the collagen press, preparing chamber slides with pressed collagen, and plating primary human ISCs for growth and analysis.
Key features
• This protocol describes how to construct and use collagen presses for chamber slides, as demonstrated in Cotton et al. [1].
• The soft collagen and culture media presented are optimized for primary human intestinal stem cells.
• The full protocol, including 3D printing, preparing collagen-coated chamber slides, and plating cells can be completed in under one week.
• This protocol requires access to a 3D printer.
Keywords: Collagen press Chamber slide Primary human intestinal stem cells Live imaging Microphysiological platform
Graphical overview
Preparing the collagen press and chamber slide for culturing human intestinal stem cells
Background
The intestinal epithelium is a dynamic tissue with differentiated cell types arising from a common crypt base columnar intestinal stem cell (ISC) [2]. Cells of the intestinal epithelium serve many roles, including absorbing nutrients, secreting mucus and hormones, providing a selectively permeable barrier between the body and luminal contents, and managing the microbiota. While much knowledge has been gleaned about the mammalian intestine using model organisms such as rodents, differences between the human intestine and those of model organisms are becoming increasingly known [3,4]. This makes it important to design systems for studying primary human intestinal cells. A common strategy for studying the human intestinal epithelium is to culture ISCs in a soft matrix and allow growth into 3-dimensional organoids (also referred to as enteroids) [5]. Organoids can be derived from primary crypts from donor epithelia [6] or via induced differentiation of pluripotent stem cells [7]. While organoids have greatly advanced our ability to study the human intestinal epithelium, they have limitations, such as an enclosed luminal compartment that is difficult to access and imaging challenges, as the cells grow in a spherical shape rather than in a flat plane. In 2017, a novel method for cultivating human small intestinal [8] or colon ISCs [9] as a self-replicating monolayer was introduced. We showed that primary cells grown on a soft collagen substrate (~9 kPa) in a growth medium rich in stem cell factors could be continuously propagated and passaged indefinitely [8]. Our recent work analyzes single-cell transcriptomic data to demonstrate that these growing monolayers largely consist of proliferating stem and progenitor cells, with minimal differentiated cells present [1]. Our optimized method for genetically modifying these primary human ISCs [10] to incorporate fluorescent reporter genes has enabled high-resolution analysis of various cell functions, including stemness [10], tight junctions [11], and the cell cycle [1]. However, we found that the thick collagen needed for maintaining primary ISC proliferation and stemness can also negatively affect high-resolution live imaging by adding distance between the growing cells and the microscope objective. When we used less collagen in a chamber slide to address this issue, we observed that the formation of a collagen meniscus at the edges of the chamber slides (Figure 1A) made consistent and clear imaging unfeasible. Cells in the thin central region of the well mostly exited the cell cycle and exhibited phenotypes indicative of auto-differentiation, while those in the thicker areas near the well walls proliferated but often risked growing beyond the working distance of our microscope (Figure 1B) [1]. To resolve this issue, we designed a collagen press that creates a flat, smooth collagen layer with a controlled and uniform thickness (Figure 1C). This innovation led to significantly more consistent cell growth and maintenance of stemness (Figure 1D). In this protocol, we describe how to design and construct collagen presses, how to pour reproducibly coated chamber slides, and how to culture and image primary human ISCs.
Figure 1. Effect of the collagen press on cell growth in chamber slide. A) Model of meniscus formation following freely poured collagen (orange) setting in chamber slides. B) Immunofluorescence of nuclei (DAPI, blue) and proliferation (EdU uptake, white) in two separate wells of intestinal stem cells (ISCs) in chamber slides poured with no press. Insets show differences in proliferative capacity and cellular spacing across areas of the same chamber well after three days of culture. C) Model for 3D-printed collagen press designed to sit across chamber slide walls and reach into wells to prevent meniscus formation. Coverslip glass glued to the leg bottoms forms a smooth surface. Photograph of a collagen press on a chamber slide. D) Immunofluorescence of nuclei (DAPI, blue) and proliferation (EdU uptake, white) in ISCs grown in two separate wells of a chamber slide using the 3D-printed collagen press. Insets indicate uniform proliferation across colonies. Adapted from Cotton et al. Scientific Reports [1]. © The Authors, some rights reserved; exclusive licensee, Springer Nature. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC) http://creativecommons.org/licenses/by-nc/4.0/.
Materials and reagents
Biological materials
LWRN Cells (ATCC, catalog number: CRL-3276)
Reagents
N-acetylcholine (NAC) (Sigma-Aldrich, catalog number: A9165)
Dulbecco's phosphate-buffered saline (dPBS) (Gibco, catalog number: 14190-144)
Na2HPO4 (Sigma, catalog number: S7907)
KH2PO4 (Sigma, catalog number: P5655)
NaCl (Sigma, catalog number: S5886)
KCl (Sigma, catalog number: P5405)
Sucrose (Fisher BP, catalog number: 220-1)
D-sorbitol (Fisher BP, catalog number: 439-500)
Y27632 (Selleck Chemical, catalog number: S6390)
DPBS (10×) (Gibco, catalog number: 14200-075)
Sodium bicarbonate (7.5%) (Gibco, catalog number: 25080-094)
Ethylenediaminetetraacetic acid (EDTA) (Corning, catalog number: 46-034-Cl)
Dithiothreitol (DTT) (Fisher Scientific BP, catalog number: 172-5)
Protease VIII (Sigma, catalog number: P5380)
Advanced DMEM/F12 (Gibco, catalog number: 12634-010)
Bovine serum albumin (Fisher Scientific, catalog number: BP1600-1)
Fetal bovine serum tetracycline negative (Gemini, catalog number: 100-800)
Primocin (InvivoGen, catalog number: ant-pm-05)
Gentamycin (Sigma-Aldrich, catalog number: G1914)
Amphotericin B (Sigma-Aldrich, catalog number: A2942)
Collagenase IV (Thermo Fisher, catalog number: LS004189)
TrypLE Express (Gibco, catalog number: 12605-010)
GlutaMAX (Thermo Fisher, catalog number: 35050061)
HEPES (Corning, catalog number: 25-060-CI)
Primocin (InvivoGen, VWR, catalog number: mspp-ant-pm2)
Pen/Strep (Thermo Fisher, catalog number: 15070063)
N-acetylcysteine (Sigma-Aldrich, catalog number: A9165)
EGF, murine (PeproTech, catalog number: 315-09)
Nicotinamide (Sigma-Aldrich, catalog number: N0636)
B27 supplement (Thermo Fisher, catalog number: 12587001)
Gastrin (Sigma-Aldrich, catalog number: G9145)
Prostaglandin E2 (PeproTech, catalog number: 3632464)
A 83-01 (Sigma-Aldrich, catalog number: SML0788)
SB202190 (PeproTech, catalog number: 1523072)
5-ethynyl-2'-deoxyuridine (EdU) (Molecular Probes, catalog number: C10634)
CuSO4 (Fisher Scientific, catalog number: S25286)
Sulfo-CY5-Azide (Lumiprobe, catalog number: A3330)
Ascorbic acid (Fisher Scientific, catalog number: AC352681000)
Cultrex rat collagen I (R&D Systems, catalog number: 3443-100-01)
Paraformaldehyde (Acros Organics, catalog number: 41678-0010)
DAPI (Invitrogen, catalog number: D3571)
NaOH (Thermo Fisher, catalog number: 12426)
NaHCO3 (Sigma-Aldrich, catalog number: S6014)
DPBS, 10× (Gibco, catalog number: 14200075)
Triton X-100 (MP Biomedicals, catalog number 02194854-CF)
Solutions
Collagen solution (see Recipes)
Neutralization buffer (see Recipes)
Small intestine (SI) maintenance media (see Recipes)
4% paraformaldehyde fixative (See Recipes)
EdU reaction buffer (see Recipes)
Sulfo-Cyanine5-azide (see Recipes)
Permeabilization solution (see Recipes)
EdU staining solution (see Recipes)
DAPI solution (see Recipes)
Recipes
Collagen solution
Note: Neutralization buffer can be made in advance and stored at 4 °C for up to one month.
Reagent Final concentration Volume for 12 mL total
Rat collagen I (3 mg/mL) 1 mg/mL 4 mL
Neutralization buffer (Recipe 2) - 8 mL
Neutralization buffer
Note: NaOH is added to neutralize the acetic acid in the Collagen I solution. The values here are for using Cultrex rat collagen supplied at 3 mg/mL in 20 mM acetic acid. The amount of NaOH will vary if collagen stocks with other concentrations are used. NaOH should be used at a 1.15 molar equivalent to the acetic acid added with the collagen.
Reagent Final concentration Volume for 8 mL total
10× dPBS 1× 1.2 mL
HEPES (1 M) 0.02 M 0.24 mL
NaHCO3 (7.5%) 0.45% w/v 0.72 mL
NaOH (1 N) see note 0.092 mL
DI water 50% total volume 6 mL
Small intestine (SI) maintenance media
Note: All reagents, except the conditioned media and mEGF, can be mixed together separately to make a “Basal Media” that can be kept at 4 °C for up to one month. Then, to make SI maintenance media, use basal media and conditioned media at a 1:1 ratio, with mEGF added at 1:20,000.
*Note: The protocol for preparing L-WRN conditioned media has been described previously [12,13]
Reagent Final concentration Volume for 50 mL total
L-WRN conditioned media* 50% 25 mL
Adv DMEM/F12 - 22.5 mL
B-27 (without Vit A) 1× 1 mL
Nicotinamide (1 M) 10 mM 0.5 mL
HEPES (1 M) 10 mM 0.5 mL
GlutaMAX (100×) 0.5× 0.25 mL
Pen/Strep (100×) 0.5× 0.25 mL
N-acetyl cysteine (NAC, 0.5 M) 1.25 mM 0.125 mL
Primocin (50 mg/mL) 50 µg/mL 0.05 mL
SB202190 (30 mM) 3 µM 5 µL
Gastrin I, human (100 μM) 10 nM 5 µL
Prostaglandin E2 (PGE2, 1 mM) 10 nM 0.05 µL
Murine EGF (1 mg/mL) 50 ng/mL 2.5 µL
4% paraformaldehyde fixative
Note: Add paraformaldehyde powder to 800 mL of PBS, start a stir bar spinning, then warm to just below 70 °C. Once all solids have dissolved, turn off the heat, allow to cool, filter the solution, and then bring to 1 L with PBS.
The final product can be stored at -20 °C indefinitely; the thawed solution should be used within two weeks.
Reagent Final concentration Volume for 1 L total
Paraformaldehyde 4% 40 g
PBS - To 1 L
EdU reaction buffer
Note: Can be stored at 4 °C indefinitely.
Reagent Final concentration Volume for 38 mL total
10× dPBS 2.6× 10 mL
DI water - 28 mL
Sulfo-Cyanine5-azide (0.8 mM)
Note: Can be stored at -20°C indefinitely.
Reagent Final concentration Volume
Sulfo-Cyanine5-azide 0.8 mM 1 mg
DI water - 1.318 mL
Permeabilization solution
Note: Can be stored at 4°C indefinitely.
Reagent Final concentration Volume
Triton-X100 0.5% 1 mL
DI water - 200 mL
EdU staining solution
Reagent Final concentration Volume for 1 mL total
EdU reaction buffer (2.63× PBS) 2× 760 µL
CuSO4 (100 mM) 0.02 M 40 µL
Sulfo-Cyanine5-azide (0.8 mM) 0.45% w/v 2.5 µL
L-ascorbic acid (1 M) 0.2 M 200 µL
DAPI solution
Note: 5 mg/mL DAPI can be stored at -20 °C indefinitely. Avoid repeated freeze/thaws. Make the final product fresh on the day of use.
Reagent Final concentration Volume for 38 mL total
DAPI (5 mg/mL in DI water) 5 µM 1 µL
PBS - 10 mL
Laboratory supplies
Chlorinated polyethylene
Cover glass thickness 1, 18 × 18 mm (Corning, catalog number: 2845-18)
Glass glue (Loctite)
15 mL conical tube (Corning, catalog number: CLS430790)
Costar Stripette, 5 mL (Corning, catalog number: 4487)
µ-Slide 4 Well ibiTreat chamber slides (Ibidi, catalog number: 80426)
Rain-X glass water repellent (Rain-X, catalog number: 800002250)
Equipment
Ultimaker S3 3D printer (Ultimaker)
Engraving pen (Millipore Sigma, catalog number: Z225568)
Incubator set to 37 °C and 5% CO2
Water bath set to 37 °C
Clinical centrifuge capable of spinning 15 mL conical tubes at least 800× g
Confocal microscope with environmental chamber
For fluorescence live imaging of PIP-H2A cell cycle reporter lines [1], we used an Andor Dragonfly spinning disk confocal microscope mounted on a Leica DMi8 microscope stand, using a Leica HC PL APO 20×/0.75 LWD air objective with pinhole size set to 40 μm. Light was collected with a 593/43 Semrock emission filter using a HC Fluotar L 25×/0.95 W 0.17 VISIR objective or a 514 nm laser was used for excitation using a 538/20 Semrock emission filter.
Procedure
Construct the collagen presses
3D-print the bodies of the collagen press using chlorinated polyethylene, following the specifications shown in Figure 2 or the design file included as File S1 (see General note 1).
Only µ-Slide 4-well ibiTreat chamber slides have been used for this protocol. If slides are used with different dimensions, the dimensions of the collagen press may need to be altered accordingly.
Figure 2. Collagen press specifications. Side lengths (in mm) shown for 3D-printing a collagen press designed to press collagen to a thickness of 0.3 mm in an Ibidi µ-slide 4-well ibiTreat chamber slide.
Prepare coverslip glass covers for each leg of the press.
Use a diamond-tipped engraving pen to cut 18 × 18 mm square coverslips to fit the downward face of each leg of the collagen press (see Video 1).
i. Working on a clean surface, overlay a leg of the collagen press onto the coverslip, aligning three of the sides.
ii. Use a fine-tipped marker to trace immediately outside the leg onto the glass. Remove the collagen press.
iii. Lay a flat, straight edge onto the coverslip and use a diamond-tipped engraving pen to cut just inside (on the side where the leg was) of the marked line. Pass the engraving pen along the full line with very light pressure and then again with slightly heavier pressure until the glass snaps off. No marker should remain on the final piece and the resulting edge should be straight.
iv. Discard any pieces that break in any way other than the intended straight line.
v. Check that the cut piece fits entirely within the base of the collagen press leg, with no glass protruding beyond the edges of the press leg.
Video 1. Cutting coverslips to cover the legs of the collagen press. Demonstration of step A2a showing how to use the collagen press to mark where to cut on the coverslip and then how to use a diamond-tipped engraving pen with a straight edge to cut a flat edge.
Glue the trimmed coverslips onto the legs of the collagen press using Glass Glue (Locktite).
i. Carefully align the edges of coverslips exactly with the sides of the press. Glass protruding beyond the wall of the press leg often disrupts the collagen when removing the press in the next step.
ii. Allow to air dry for 1 h at room temperature.
Make collagen presses hydrophobic.
Coat the collagen press with Rain-X glass treatment, following the manufacturer’s instructions. Only coat the coverslip glass adhered to the bottoms of the legs and the sides of the legs; the top of the press that rests on the chamber walls does not need to be coated.
Wipe the press (with attached glass coverslips) clean before starting. Apply Rain-X onto a small dry cloth. Wipe onto the press using small circular motions, taking care to contact all sides of the legs. Allow to dry. Repeat. Allow to dry. Clean the final product with a gentle spray of water, then dry with a paper towel.
Prepare collagen-coated chamber slides
Sterilize collagen presses.
Do not autoclave collagen presses as this can cause them to warp.
On the day of use, spray collagen presses generously with 70% ethanol. Then, set in a laminar flow hood under ultraviolet light for 1 h, turning 90° every 15 min.
Keep collagen presses in the laminar flow hood until use. No pre-warming is necessary.
Prepare collagen solutions on ice.
Prepared neutralization buffer can be stored in a sealed container at 4 °C for over a month without an obvious effect on collagen quality.
Add the desired amount of neutralization buffer to a 15 mL conical tube. Store neutralization buffer and the bottle of rat collagen I on ice for 10 min to allow everything to become ice cold. Per well, 250 µL of final neutralized collagen solution is needed, resulting in 1 mL of collagen per full chamber slide. As the collagen solution is viscous, we regularly prepare ~20% more solution than needed to avoid running out before the last chamber.
Add collagen to the neutralization buffer.
i. For 3 mg/mL collagen stock, use neutralization buffer and collagen at a 2:1 ratio to reach a final concentration of 1 mg/mL collagen.
For example, if pouring four chamber slides, prepare 5 mL of total solution (1.66 mL of collagen + 3.33 mL of neutralization buffer) to have 1 mL per slide with 20% excess.
ii. The collagen solution is viscous; use a 5 mL stripette and pipette up/down multiple times in the tube of neutralization buffer to wash out all collagen from the stripette.
iii. Cap the tube tightly and gently invert 10 times. Return the tube to ice and use as quickly as possible.
We recommend using the neutralized collagen solution within 10 min post-mixing.
Add collagen and press to the chamber slide.
Open µ-slide 4-well ibiTreat chamber slides inside of a clean laminar flow hood. Place the chamber slide on the floor of the hood and remove the cover.
Add collagen to each well.
i. Mix tube of final collagen solution (pre-chilled solutions) and then store on ice. Add 250 µL of collagen solution to each well. Use a pipette tip and/or tilting and tapping the slide to cover the whole bottom of all four wells with collagen solution with no dry spots (Video 2).
ii. Once no dry spots remain, gently lower the collagen press onto the inner walls of the chamber slide (as shown in Figure 1C). Adjust so the legs are freely floating into the middle of the wells and not pressing against any of the walls of the wells.
iii. Transfer the chamber slide and collagen press to a 37 °C incubator for 90 min.
iv. Work quickly; prepare only 1–2 chamber slides side-by-side before transferring to the incubator.
We recommend moving plates to the incubator within 2 min after adding liquid collagen.
Video 2. Pouring collagen with the press. Demonstration of step B3b showing how to add collagen to each chamber of the slide, tap to remove any bubbles and cover the full base of each chamber, then place the collagen press on top so it is not pressing against any wall of the chamber.
Remove the collagen press and store the finished collagen-coated chamber slide.
Remove the collagen press.
i. After 90 min incubation at 37 °C, remove the chamber slide and collagen press from the incubator and transfer back into the laminar flow hood.
ii. Carefully remove the press following the technique shown in Video 3.
a). With the slide turned perpendicular to your field of vision, carefully tilt the collagen press toward you, then lift up and out of the well.
b). Watch for any wells that show any movement of collagen. If any part of the final patty is pulled up by the collagen press, such as being stuck on a glass overhang (see note in step A2b), mark the outside of the well with a marker and refrain from using that well.
Video 3. Removing collagen press. Demonstration of step B4a showing how to carefully remove the press, check for any large bubbles or gaps in the coverage, and then add PBS for storage of the coated slide. The small bubbles seen following the removal of the press are normal and not expected to have any downstream impact.
iii. Once the press is removed, visually inspect all wells for full collagen coating.
a). Tiny bubbles trapped around the collagen base are normal and have not been seen to affect cell growth.
b). Watch for wells with large bubbles, dry spots, or any areas where the collagen is visibly pulled off or folding over. Mark the outside of these wells with a marker and refrain from using them.
Add 700 µL of room-temperature, sterile dPBS to each chamber. Pipette the dPBS gently to avoid disrupting the collagen coating. Cover each chamber slide tightly with the supplied covers.
Store collagen-coated chamber slides in a sealed zip-top bag at room temperature and protected from light until use.
Note: Collagen-coated plates have been used up to 6 months post-coating with no apparent effect on cell growth as long as they are protected from evaporation.
Seeding cells
Prepare primary human ISCs for seeding.
Primary human ISC monolayers should be expanded on thick collagen patties, as previously described [8,9]. Cells are grown in SI maintenance media as described in a recent protocol paper [14]. One day prior to passaging, add Y-27632 to the media at a final concentration of 10 µM.
Grow cells at least two days prior to passaging; 3–6 days is generally desired. Weaker growth has been noted for cells passaged after only one day or beyond 7 days.
Passage cells onto chamber slides.
Use a pipette to transfer the collagen patty and 1 mL of culture medium to a conical tube. Add 150 µL of 5,000 U/mL collagenase IV. Cap and incubate in a 37 °C water bath for ~25 min, swirling occasionally, until all collagen is dissolved and cells fall to the bottom of the tube.
i. During this incubation, wash the collagen-coated chamber slide by incubating three times for 5 min each with 700 µL of dPBS at room temperature. No shaking is necessary.
ii. Be careful not to touch the collagen with the pipette/aspirator tip when removing the dPBS as it can easily be pulled off of the chamber slide.
Centrifuge at 800 RCF for 2 min.
Aspirate out the media and wash pellet with 3 mL of sterile dPBS.
Centrifuge at 800 RCF for 2 min.
Aspirate out the dPBS and resuspend the pellet in 1 mL of pre-warmed TrypLE. Incubate in a 37 °C water bath for 5 min. Break up the pellet by pipetting, then quench by adding an equal volume of pre-warmed advanced DMEM/F12 + 10% fetal bovine serum.
Centrifuge at 800 RCF for 2 min, aspirate out the liquid, then resuspend the pellet in SI maintenance media.
Add the desired number of cells to each chamber in 700 µL of total media (see General note 2).
Incubate at 37 °C, changing media every 1–2 days until live imaging.
Immunofluorescence staining for EdU
The easiest way to validate that the pressed collagen is supporting ISC growth is to stain for EdU to mark active proliferation following 3+ days of culture in SI maintenance media.
Fix cells following growth on the pressed chamber slides.
i. Aspirate maintenance media, add 500 µL of room-temperature 4% paraformaldehyde fixative to each chamber, incubate at room temperature for 20 min, aspirate off fixative, and wash twice in dPBS.
ii. Store fixed slides with PBS in each chamber in a zip-top bag at 4 °C until staining.
On the day of staining, wash fixed chambers three times with PBS.
Add enough permeabilization solution to cover the full base of the well. Incubate for 20 min at room temperature.
Wash two times with PBS, 5 min each.
Add EdU staining solution. Incubate for 1 h at room temperature, protected from light.
Wash with PBS for 5 min, protected from light.
Add DAPI staining solution for 3 min, protected from light.
Wash three times with PBS, 5 min each, protected from light.
Store in PBS in a sealed zip-top bag at 4 °C until imaging. Image within two weeks to avoid fluorescent signal fading.
Validation of protocol
The major readout of a properly functioning collagen patty in this format is that primary human ISCs maintain growth and proliferation for multiple days when cultured in the wells. Since primary human ISC monolayers require a soft surface (~9 kPa) to preserve stemness, ISCs cease to proliferate if the collagen hydrogel is too thin. Figure 1B illustrates the phenotypic and growth differences observed when ISCs are cultured on a collagen layer that is too thin and uneven. In contrast, Figure 1D shows optimal ISC growth, with thick bands of proliferating ISCs evenly distributed along the edges of all monolayers after three days of growth, regardless of their position within the chamber well.
This protocol, or components of it, has been employed and validated in the following research article(s): Cotton et al. [1]. An in vitro platform for quantifying cell cycle phase lengths in primary human intestinal epithelial cells. Scientific Reports (Figures 3, 4, 6). In this publication, collagen-pressed chamber slides from various preparation batches were utilized with multiple cell cycle reporter lines of primary human ISCs. All assays confirmed sustained ISC proliferation, with the cell cycles of hundreds of control cells closely matching the 12–21 h cell cycle lengths observed in mouse models (Figure 3) [15,16]. This demonstrates that the collagen press protocol reliably produces collagen-coated chamber slides optimized for maintaining ISC growth and enabling live imaging of fluorescence reporters over multiple days. Specifics regarding cell cycle reporter ISC lines and their corresponding live imaging and analysis parameters can be found in this previous publication.
Figure 3. Cell cycle phases directly quantified from human intestinal stem cells (ISCs) grown on chamber slides with pressed collagen. Violin plots denoting cell cycle phase lengths (in hours) across 284 primary human ISCs from n = 2 wells using our novel PIP-H2A reporter. Reprinted from Cotton et al. Scientific Reports [1]. © The Authors, some rights reserved; exclusive licensee, Springer Nature. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC) http://creativecommons.org/licenses/by-nc/4.0/.
General notes and troubleshooting
General notes
Collagen presses can be 3D printed from a variety of materials with no expected effect on function. Our group has used presses printed using chlorinated polyethylene and acrylonitrile butadiene styrene and both worked equivalently. A potential issue may arise with sterilizing presses before use, so it is suggested to avoid materials that are easily degraded by moderate exposure to ethanol.
The number of cells passaged to the chamber slides will depend on the assay being run. Primary human intestinal monolayers are generally passed at ratios between 1:3 and 1:5 from a confluent well for normal passaging. Ibidi µ-slide 4-well chamber slides have 2.5 cm2 of growth area per well, ~25% higher than a typical well of a 24-well culture plate, which has 1.9 cm surface per well. For applications such as monitoring cell cycles, we passage cells intentionally thin to avoid contact inhibition. However, higher coverage and larger colonies may be desired for other assays, such as those looking at tight junctions or cell differentiation. We recommend trialing passaging density on collagen-coated 24-well plates prior to using the chamber slides if cost is an issue.
While we maintain our ISCs as monolayers for routine expansion prior to imaging assays, previous studies demonstrate that primary human ISCs cultured as 3-dimensional organoids can also be plated onto collagen patties to form monolayers [9]. Therefore, it is not necessary for a lab that routinely cultures primary human ISCs as organoids to establish separate lines as monolayers; the cells grown as organoids can be plated directly onto pressed collagen chamber slides.
We recommend testing different lengths of legs for the collagen press to ascertain that the thickness of the resulting collagen coating maintains the cellular properties to be analyzed over several days. For example, we made a collagen press with staggered leg lengths to find whether thin collagen layers support primary human ISC growth (Figure 4A). We saw sustained EdU uptake in all four wells after three days of growth (Figure 4B), indicating that thicknesses from 0.3 to 0.9 mm were all sufficient to support growth and stemness, so we chose the thinnest collagen tested to optimize imaging. We suggest that each group starts with a similar analysis to determine that their properties of interest (stemness markers, growth rate, -omics analyses, etc.) are similar between the pressed chamber slide and their original platform. Also, biological differences can arise between platforms, especially if the original platform is notably different such as 3D organoids. For this reason, controls and experimental groups should be run side-by-side on the pressed chamber slides to avoid platform-derived differences in the analysis.
Figure 4. Designing a collagen press with staggered leg lengths to test different thicknesses of collagen coating. A) Leg lengths shown are measured from the bottom of the horizontal base, not including the portion of the press that sits atop the chamber slide walls. Values in blue are the calculated thickness of collagen expected after accounting for the lengths of the legs and the attached cover slip-thickness glass. B) Representative images of cells cultured on collagen patties with thicknesses ranging from 0.3 to 0.9 mm stained for nuclei (DAPI, blue) and proliferation (EdU uptake, white).
While we have only used these chamber slides with pressed collagen for primary human intestinal stem cells, they can likely be used for other cell and tissue types as well, especially those already known to grow on a collagen matrix. If using this collagen press for a different cell type, we recommend testing different thicknesses of collagen, as in General note 4 and Figure 4, to check that the platform is optimized for supporting your cell type before beginning analysis.
It should be noted that a comprehensive survey of the capabilities and limitations of this platform has not been done. As discussed in General note 4, the major intended readout of this platform for the original study was tracking cell cycle, so all analyses by our group pertained to proliferation over time. We have not tested sustained cell culture past 5 days on the pressed slides, passaged cells from pressed slides for further culturing, or analyzed our cells for properties other than cell cycle. As cells appear to proliferate normally in days 3–5, we expect that they would continue to act equivalently to cells grown on typical collagen-coated wells, but researchers will need to test further biological properties individually.
Troubleshooting
Problem 1: Collagen coatings on each well appear uneven.
Possible cause: The collagen press is warped due to autoclaving or exposure to high heat.
Solution: Re-make the collagen presses and sterilize with a gentle method such as ultraviolet light and ethanol.
Problem 2: Glass panes coating the legs of the collagen press detach into the wells.
Possible cause: Many common super-glue brands are not made for glass.
Solution: Use Locktite glue glass instead of a general super glue.
Problem 3: The collagen press pulls up the collagen patty upon removal after incubation.
Possible cause: Any amount of overhang from the glass pane will easily get caught in the solidifying collagen.
Solution: Re-cut the coverslips to better fit the legs. Slightly less glass is better than too much. Test each leg of the collagen press for overhangs before first use. After adhering the panes to the collagen press legs and allowing time to dry, take a small tool such as a metal spatula and rub it along the sides of the collagen press, moving from the base toward the glass. If the tool hits the glass pane, pry it off and reattach the glass with no overhang.
Problem 4: The full layer of cells moves downward (in the Z direction) outside of the frame of imaging during extended live imaging.
Possible cause: We often see a small amount of settling (10–20 µm) during a 48-h live image, but exaggerated sinking could be due to evaporation.
Solution: All live imaging should image across a z-stack to account for small changes in collagen thickness and cell movement. We use up to 60 µm when possible. To avoid evaporation, make sure that the cover of the chamber slide is firmly attached during imaging, fill a humidity chamber in the air input line if available, and add open water to the environmental chamber by filling any moats or adding conical tube caps filled with water to buffer the amount of evaporation that occurs.
Acknowledgments
The authors thank the human donor for the gift of tissue. We thank Steven Emanual and Michael Cotton for 3D printing the collagen presses and Henry Taylor for helping film the videos. The Microscopy Services Laboratory, Department of Pathology and Laboratory Medicine, is supported in part by the P30 CA016086 Cancer Center Core Support Grant to the UNC Lineberger Comprehensive Cancer Center. The Andor Dragonfly microscope was funded with support from National Institutes of Health grant S10OD030223. This research was supported by the AGA-Bristol Myers Squibb Research Scholar Award in IBD (J.B.), R01DK115806 and R01DK109559 (S.T.M.), by CGIBD support through funding from the National Institutes of Health, P30 DK034987, and the Katherine E. Bullard Charitable Trust for Gastrointestinal Stem Cell and Regenerative Research.
Competing interests
S.T.M has a financial interest in Altis Biosystems Inc., which licenses technology used in this study.
References
Cotton, M. J., Ariel, P., Chen, K., Walcott, V. A., Dixit, M., Breau, K. A., Hinesley, C. M., Kedziora, K. M., Tang, C. Y. and Zheng, A. (2024). An in vitro platform for quantifying cell cycle phase lengths in primary human intestinal epithelial cells. Sci Rep. 14(1): 15195.
Barker, N., (2013). Adult intestinal stem cells: critical drivers of epithelial homeostasis and regeneration. Nat Rev Mol Cell Biol. 15(1): 19–33.
Burclaff, J., Bliton, R. J., Breau, K. A., Ok, M. T., Gomez-Martinez, I., Ranek, J. S., Bhatt, A. P., Purvis, J. E., Woosley, J. T. and Magness, S. T. (2022). A Proximal-to-Distal Survey of Healthy Adult Human Small Intestine and Colon Epithelium by Single-Cell Transcriptomics. CMGH Cell Mol Gastroenterol Hepatol. 13(5): 1554–1589.
Malonga, T., Vialaneix, N. and Beaumont, M. (2024). BEST4+ cells in the intestinal epithelium. Am J Physiol, Cell Physiol. 326(5): C1345–C1352.
Stelzner, M., Helmrath, M., Dunn, J. C. Y., Henning, S. J., Houchen, C. W., Kuo, C., Lynch, J., Li, L., Magness, S. T. and Martin, M. G. (2012). A nomenclature for intestinal in vitro cultures. Am J Physiol Gastrointest Liver Physiol. 302(12): G1359–G1363.
Zachos, N. C., Kovbasnjuk, O., Foulke-Abel, J., In, J., Blutt, S. E., de Jonge, H. R., Estes, M. K. and Donowitz, M. (2016). Human Enteroids/Colonoids and Intestinal Organoids Functionally Recapitulate Normal Intestinal Physiology and Pathophysiology. J Biol Chem. 291(8): 3759–3766.
Spence, J. R., Mayhew, C. N., Rankin, S. A., Kuhar, M. F., Vallance, J. E., Tolle, K., Hoskins, E. E., Kalinichenko, V. V., Wells, S. I. and Zorn, A. M. (2010). Directed differentiation of human pluripotent stem cells into intestinal tissue in vitro. Nature. 470(7332): 105–109.
Wang, Y., Gunasekara, D. B., Reed, M. I., DiSalvo, M., Bultman, S. J., Sims, C. E., Magness, S. T. and Allbritton, N. L. (2017). A microengineered collagen scaffold for generating a polarized crypt-villus architecture of human small intestinal epithelium. Biomaterials. 12844–55.
Wang, Y., DiSalvo, M., Gunasekara, D. B., Dutton, J., Proctor, A., Lebhar, M. S., Williamson, I. A., Speer, J., Howard, R. L. and Smiddy, N. M. (2017). Self-renewing Monolayer of Primary Colonic or Rectal Epithelial Cells. CMGH Cell Mol Gastroenterol Hepatol. 4(1): 165–182.e7..
Breau, K. A., Ok, M. T., Gomez-Martinez, I., Burclaff, J., Kohn, N. P. and Magness, S. T. (2022). Efficient transgenesis and homology-directed gene targeting in monolayers of primary human small intestinal and colonic epithelial stem cells. Stem Cell Rep. 17(6): 1493–1506.
Ok, M. T., Liu, J., Bliton, R. J., Hinesley, C. M., San Pedro, E. E. T., Breau, K. A., Gomez-Martinez, I., Burclaff, J. and Magness, S. T. (2023). A leaky human colon model reveals uncoupled apical/basal cytotoxicity in early Clostridioides difficile toxin exposure. Am J Physiol Gastrointest Liver Physiol. 324(4): G262–G280.
Miyoshi, H. and Stappenbeck, T. S. (2013). In vitro expansion and genetic modification of gastrointestinal stem cells in spheroid culture. Nat Protoc. 8(12): 2471–2482.
VanDussen, K. L., Sonnek, N. M. and Stappenbeck, T. S. (2019). L-WRN conditioned medium for gastrointestinal epithelial stem cell culture shows replicable batch-to-batch activity levels across multiple research teams. Stem Cell Res. 37101430.
Hinman, S. S., Wang, Y., Kim, R. and Allbritton, N. L. (2020). In vitro generation of self-renewing human intestinal epithelia over planar and shaped collagen hydrogels. Nat Protoc. 16(1): 352–382.
Schepers, A. G., Vries, R., van den Born, M., van de Wetering, M. and Clevers, H. (2011). Lgr5 intestinal stem cells have high telomerase activity and randomly segregate their chromosomes. EMBO J. 30(6): 1104–1109.
Matsu-ura, T., Dovzhenok, A., Aihara, E., Rood, J., Le, H., Ren, Y., Rosselot, A. E., Zhang, T., Lee, C. and Obrietan, K. (2016). Intercellular Coupling of the Cell Cycle and Circadian Clock in Adult Stem Cell Culture. Mol Cell. 64(5): 900–912.
Supplementary information
The following supporting information can be downloaded here:
File S1: Collagen Press Design File.
Article Information
Publication history
Received: Aug 14, 2024
Accepted: Sep 27, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Stem Cell > Adult stem cell > Intestinal stem cell
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Cell Biology > Cell imaging > Live-cell imaging
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Peer-reviewed
A Protocol for co-Injecting Cells with Pulverized Fibers for Improved Cell Survival and Engraftment
AS Ana I. Salazar-Puerta *
NO Neil Ott *
LD Ludmila Diaz-Starokozheva
DD Devleena Das
WL William R. Lawrence
JJ Jed Johnson
RH Robert Houser
NH Natalia Higuita-Castro
KS Kristin I. Stanford
DG Daniel Gallego-Perez
(*contributed equally to this work)
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5117 Views: 264
Reviewed by: Pilar Villacampa AlcubierreShivaram Selvam Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Biomedical Materials Research Part A Nov 2023
Abstract
Adipose tissue is crucial for medical applications such as tissue reconstruction, cosmetic procedures, and correcting soft tissue deformities. Significant advances in the use of adipose tissue have been achieved through Coleman’s studies in fat grafting, which gained widespread acceptance due to its effectiveness and safety. Despite its benefits, adipose tissue grafting faces several limitations, including high absorption rates due to insufficient support or anchorage, replacement by fibrous tissue, migration from the intended site, and loss of the initial desired morphology post-administration. To counteract these constraints, there is a need for improved grafting techniques that enhance the predictability and consistency of outcomes. Biomaterials are extensively used in tissue engineering to support cell adhesion, proliferation, and growth. Both natural and synthetic materials have shown promise in creating suitable microenvironments for adipose tissue regeneration. PLGA, a synthetic copolymer, is particularly notable for its biocompatibility, biodegradability, and tunable mechanical properties. Here, we describe a protocol using milled electrospun poly(lactic-co-glycolic acid) (PLGA) fibers combined with lipoaspirated tissue to create a fibrous slurry for injection. By pulverizing PLGA fiber mats to create fiber fragments with increased pore size and porosity, we can influence key cellular responses and enhance the success of adipose tissue–grafting procedures. This approach improves anchorage and support for adipocytes, thereby increasing cell viability. This method aims to enhance vascularity, perfusion, and volume retention in adipose tissue grafts, which addresses many of the limitations of current approaches to adipose tissue grafting and holds promise for more consistent and successful outcomes.
Key features
• Adipose tissue for tissue reconstruction.
• Need for improved engraftment and volume retention.
• Pulverized PLGA fiber mats to create a fibrous "slurry" that allows injection.
• PLGA fibers co-injected with lipoaspirated tissue.
• Improved adipose engraftment outcomes (e.g., perfusion, vascularity, and retention of graft volume).
Keywords: Pulverized PLGA fibers Adipose tissue engraftment Volume retention Increased vascularity Tissue reconstruction Soft-tissue augmentation
Graphical overview
Overview of the subcutaneous implantation of human adipose tissue and pulverized electrospun poly(lactic-co-glycolic acid) (PLGA) fibers in a mouse model
Background
Adipose tissue plays a crucial role in numerous medical applications, such as tissue reconstruction, cosmetic procedures, and the correction of soft tissue deformities. The first use of autologous adipose tissue in humans was described in 1889 by Van der Meulen [1]. While many subsequent studies have explored this technique to correct various defects, a significant advancement in fat transplantation occurred with the publication of Coleman’s studies in fat grafting [2]. The Coleman technique, which utilizes autologous adipose tissue, has gained widespread acceptance due to its effectiveness and safety [3]. This method involves harvesting adipose tissue via liposuction, processing it, and reinjecting it into various tissue depths. Studies have demonstrated its efficacy in yielding viable adipocytes and sustaining optimal cellular function within fat grafts, making it a valuable method for soft-tissue augmentation and tissue repair [4].
The use of adipose tissue has led to numerous advancements in the use of autologous fat grafts not only in aesthetic treatments but also in various medical specialties such as therapies for breast cancer [2,5]. The regenerative potential of adipose tissue is attributed to the presence of stem cells, which makes it an ideal filler due to its availability, low donor-site morbidity, cost-effectiveness, and biocompatibility [6]. Adipose-derived stromal cells can secrete various growth factors, such as VEGF, HGF, and TGF-β, which play a crucial role in tissue remodeling. These growth factors influence the differentiation of stem cells and promote angiogenesis, thereby enhancing the efficacy of adipose tissue in tissue engineering approaches [7]. Moreover, after transplantation, ischemic adipocytes attract macrophages, initiating revascularization through neoangiogenesis, suggesting thus that the foreign body response is associated with increased vascularization [6,8].
However, existing autologous adipose engraftment methods have several limitations that impact the success and consistency of clinical outcomes. In clinical practice, a major issue with adipose tissue autotransplantation is the absorption rate over time, which ranges from 25% to 70% of the total implanted volume [2,6,9]. In 1987, the American Society of Plastic and Reconstructive Surgeons reported that only 30% of the injected autologous fat was expected to survive for one year [9]. Additionally, it was observed that the transplanted tissue often became filled with connective tissue, indicating the death of the adipose tissue followed by its replacement with fibrous tissue or newly formed metaplastic fat [9]. In contrast, Peer proposed the cellular survival theory, which suggests that the final volume after an adipose tissue transplant depends on the number of living adipocytes at the time of transplantation [9,10]. These challenges underscore the need for better grafting techniques to improve the predictability and consistency of adipose grafting results.
Biomaterials are extensively utilized in tissue engineering for their capability to support cell adhesion, proliferation, and growth during the development of new tissues. For instance, porous scaffolds and hydrogels, both natural and synthetic, have shown promise in improving adipose tissue grafting procedures. These materials have been found to enhance cell viability, promote vascularization, and support the growth of adipocytes derived from progenitor cells. Collagen-based scaffolds, hyaluronan-heparin-collagen hydrogels, fibrinogen hydrogels, and decellularized extracellular-matrix scaffolds are among the natural materials that have demonstrated potential in creating a suitable microenvironment for adipose tissue regeneration [11–13]. On the other hand, synthetic materials such as poly(N-isopropylacrylamide) (PNIPAm), polyglycolic acid (PGA), polymeric nanocomposites, and poly(lactic-co-glycolic acid) (PLGA) have shown promise in tissue regeneration, contributing to adipose tissue grafting [14,15].
In this context, we propose a protocol to improve grafting outcomes using milled/pulverized electrospun PLGA fibers combined with lipoaspirated tissue to create a fibrous slurry for injection into recipient tissue. PLGA is a copolymer widely used in tissue engineering due to its excellent biocompatibility, biodegradability, and mechanical properties. PLGA degrades into lactic acid and glycolic acid, which are naturally metabolized by the body, minimizing its toxicity [16]. Additionally, the mechanical properties (e.g., degradation rate) can be adjustable by altering the ratio of lactic acid to glycolic acid monomers. Higher lactic acid content results in slower degradation, while higher glycolic acid content leads to faster degradation. The ability to control the degradation rate by adjusting the ratio is a unique feature of PLGA that sets it apart from other commonly used biodegradable polymers, such as PGA or polylactic acid (PLA) [17,18]. While PLGA can be easily electrospun into microscale fibers for scaffolding in cell culture, these fibers must be processed into small pieces to make them suitable for injection. Therefore, we pulverize the electrospun fiber mats using a mini mill to create small fibrous clusters with increased pore size and porosity, which are critical for influencing key cellular responses.
We present a detailed protocol for using milled electrospun PLGA fibers co-injected with adipose tissue, aimed at enhancing vascularity, perfusion, and graft volume retention. This approach offers the potential to improve the consistency and success of adipose grafting techniques, addressing the limitations of current methods.
Materials and reagents
Reagents
Polylactide-co-glycolide acid (PLGA) 82:18 lactide:glycolide (Corbion Purac, catalog number: PLG8218)
Hexafluoroisopropanol (Oakwood Chemical, catalog number: 3409)
0.9% sodium chloride solution (saline) (Henry Shein, catalog number: 1047098)
Ethanol, 200 Proof (100%) (Decon Labs, catalog number: 2701)
Betadine (Henry Schein, catalog number: 67618–150-09)
Isopropyl alcohol (Fisher Scientific, catalog number: A416P-4)
Isoflurane (Primal Health Care, catalog number: NDC 66794–017-10)
Artificial tear ointment (Henry Schein, catalog number: 1338333)
Nairing cream (Nair, catalog number: 339823)
Solutions
70% ethanol solution (see Recipes)
Recipes
70% ethanol solution
Mix 35 mL of 100% ethanol with 15 mL of sterile water, to create 50 mL of 70% ethanol solution.
Laboratory supplies
0.5 mm sieve (IKA, model: 2939000)
0.25 mm sieve (IKA, model: 2938900)
2 mL Eppendorf Tubes (VWR, catalog number: 87003-298)
50 mL Falcon tubes (VWR, catalog number: 89039-660)
Alcohol swabs (Henry Shein, catalog number: HS1007)
Cotton tip applicator (McKesson, Q-Tip, catalog number: 785468)
Liquid glue (3M Vetbond, catalog number: B07Q39FL9M)
6–0, C-22 black braided silk sutures (Henry Schein, catalog number: 101–2636)
Surgical tape (3M, catalog number: 1527–0)
18-gauge needle (Central Infusion Alliance, catalog number: BD 305196)
3 mL syringe (Fisherbrand, catalog number: 14–955-457)
Parafilm (Bemis, catalog number: PM999)
Equipment
Electrospinner (Glassman High Voltage, model: PS/FJS0R02.4)
Micro-miller (IKA Mini-Mill, model: 2836001)
Cold centrifuge (VWR, Eppendorf, model: 5430 R, catalog number: 76458-526)
Weigh scale (VWR, model: VWR-6001E, catalog number: 10204-996)
Analytical balance (VWR, model: VWR-310AC/CAL, catalog number: 89422-676)
Slide warmer with temperature control (4MD Medical, catalog number: CASCXH-2001)
Somnosuite low-flow anesthesia system (Kent Scientific, catalog number: 13-005-111)
Induction chamber (Kent Scientific, catalog number: Somno-0705)
Warming pad (Kent Scientific, catalog number: RT-0501)
Electric razor with vacuum (Remmington, catalog number: VPG-6530)
Tenotomy scissors (Fine Science Tools, catalog number: 14066–11)
Ridge-less forceps (Fine Science Tools, catalog number: 11016-17)
Germinator 500 (Braintree Scientific, catalog number: GER5287120V)
Procedure
Pulverized electrospun PLGA fiber fabrication
For fiber fabrication, electrospinning was used to create ultra-fine fibers from poly(lactic-co-glycolic acid) (PLGA) polymer. The process begins with the preparation of a solution where PLGA is dissolved in an organic solvent (hexafluoroisopropanol), resulting in a homogeneous solution. This solution is then loaded into a syringe fitted with a needle, and a high-intensity electric field is applied between the needle tip and a grounded collector plate at a constant flow rate. The electric field induces the formation of a liquid droplet at the needle's tip, which elongates into a jet and is ejected toward the collector. As the jet travels, the solvent evaporates, and the polymer solidifies into very thin fibers, forming a non-woven mat. This process is driven by the interaction between electrostatic forces and the surface tension of the polymer solution, and it depends on factors such as the solution's viscosity, the strength of the electric field, and the distance between the needle and the collector. Pulverization of PLGA is then performed by breaking down the electrospun fiber mat into fine fragments. This is achieved through mechanical grinding, where the mat is subjected to shear and impact forces, reducing it to smaller fragments. Milled and un-milled fibers were obtained from Nanofibers Solutions (Columbus, OH) as outlined below:
Dissolve PLGA, at a ratio of 82:18 lactide to glycolide, in hexafluoroisopropanol at 6% w/w (Figure 1).
Figure 1. Pulverized electrospun poly(lactic-co-glycolic acid) (PLGA) fibers. A. PLGA at a lactide to glycolide ratio of 82:18. B. Electrospinning to get an electrospun fiber mat. C. The resulting PLGA injectable fiber fragments. Scanning electron microscopy (SEM) micrographs of the PLGA before and after the milling process.
Mix at room temperature for at least 24 h.
Electrospin the polymer solution at +12 kV/-3 kV at a flow rate of 5 mL/h until reaching a 0.1–0.3 mm thick nanofiber sheet at room temperature and humidity with a tip-to-collector distance of 20 cm using an 18-gauge needle.
Evaporate the solvent from the nanofiber sheet for ~16 h at room temperature.
Cut the electrospun mat into 1 cm wide strips and feed into the mini mill. This will grind the fibers into smaller fragments.
Perform milling with the 60-mesh sieve for the first pass. Then, re-run the fibers that passed through that sieve through the 40-mesh sieve mill. Then, run the fibers that passed through that sieve through the 20-mesh sieve to achieve the final desired particle size.
Co-injectable nanofiber and adipose tissue preparation
Adipose tissue was sourced from Cosmetic & Plastic Surgery of Columbus via standard minimally invasive procedures such as single port cannula liposuction; no further characterization of the tissue was done (See General Note 1).
Note: Preparation should be done in a sterile environment for best results.
Transport the tissue on ice and place it in appropriate containers (See General Note 2).
Centrifuge at 195× g for 1 min at 4 °C (Figure 2).
Note: This step will separate the tissue from oils and anesthetics that can permeate the tissue during liposuction surgery.
Figure 2. Co-injectable nanofiber and adipose tissue. A. Adipose tissue collected from patients using standard liposuction procedures. B. The tissue is centrifuged to separate fat oils and anesthesia from viable cells. C. Pulverized fibers (50 mg) are combined with (D) adipose tissue (1 mL) to form an injectable solution (50 mg/mL).
Discard excess oils and debris.
Place 50 mg of fibers into a 2 mL tube.
Add 1 mL of tissue to the fiber-containing tube (See General Note 3).
Mix tissue and fibers into a 3 mL syringe and cover the end with parafilm for surgical use.
Fat engraftment surgery preparation
Sterilize all surgical tools, surgical trays, and materials before surgery.
Note: Between same-day animal surgeries, clean instruments with 70% ethanol solution and sterilize them using the germinator prior to the next surgery.
Set and maintain the surgical warming pads at 37 °C during surgery.
Place mouse cages onto a slide warmer set to 37 °C.
Note: Each mouse should be placed in individual cages to prevent agitation of the wound site.
Weigh each mouse and record baseline weights.
Note: Baseline weight will be compared with post-surgery values to evaluate mouse recovery in the form of post-implantation weight loss.
Completely remove hair from the incision area 24 h prior to surgery. Shave hair with the razor, apply Nairing cream directly to the skin with a cotton tip applicator, and leave it for 10 s. Remove the cream entirely with warm water and cotton balls.
Note: It is crucial to remove the Nairing cream thoroughly to prevent serious skin burns.
In vivo fat engraftment surgery
Place the mouse into the induction chamber (see General Note 4).
Set the Somnosuite low-flow anesthesia system at 5% isoflurane in room air at ~500 mL/min and induce flow into the induction chamber to anesthetize the mouse (see General Note 5).
At full anesthetic depth, move the mouse to the nose cone on the warming pad.
Note: Confirm the anesthetic depth of the mouse by testing the toe-pinch reflex.
Quickly set the anesthesia system at 1.5%–2% isoflurane at ~250 mL/min and induce flow into the nose cone for the duration of the procedure.
Apply artificial tear ointment to the mouse’s eyes to maintain moisture.
Apply betadine followed by isopropyl to the incision area using cotton tip applicators (repeat this step three times).
Create a 1-inch incision horizontally on the upper and lower dorsal area using ridge-less forceps to gently pull the skin taut, allowing for an easier cut (Figure 3).
Figure 3. Fat engraftment surgery. Prepare the back of the athymic nude mice for injection. Create a 1-inch incision horizontally on the upper and lower dorsal area for the injection of adipose + fibers and their internal control with only adipose tissue, respectively.
Form a cavity under the mouse’s skin by detaching the surrounding fascia using tenotomy scissors.
Push approximately 100–120 μL of warm saline into each side of the incision.
Connect a 3 mL syringe loaded with adipose tissue and fibers to a 4 mm cannula.
Note: Inject adipose tissue alone as a control, as opposed to the adipose tissue–fiber combination treatment.
Push 1 mL from the syringe load into the cavity (see General Note 6).
Note: Larger syringes (>3 mL) may be required to push the adipose tissue and fibers through the 4 mm cannula, as this can facilitate the displacement of larger volumes while generating less pressure, minimizing the risk of clogging. The syringe may have to be pumped back and forth to ensure that any remaining contents are injected, and it may require readjusting the cannula and extension of the syringe to push in all of the load.
Close the cavity by suturing the skin with 6–0 braided sutures along the incision in a single interrupted suture pattern.
Clean the skin of the mouse using an alcohol pad.
Seal the incision line using liquid glue/bandage.
Remove the mouse from the nose cone, weigh the mouse, and record post-surgery weight.
Place the mouse into a new, clean cage and allow the mouse to recover.
Monitor the status of the mouse until fully awake (1–5 min).
Keep mice on the slide warmer at 37 °C throughout the duration of the study.
Place mash (i.e., wetted food) and hydrogel in Petri dishes into each mouse cage to facilitate fluid/food intake.
Data analysis
Scanning electron microscopy (SEM) is widely used to characterize the physical properties of electrospun fiber scaffolds before and after milling, evaluating the effects on porosity, pore size, and fiber diameter, which impact cellular responses.
To assess the biocompatibility of the milled fibers with adipose tissue, in vitro cultures of adipocytes with the fibers can be used to run viability assays (e.g., live/dead kit) and fluorescence microscopy for quantifying live cells. Similarly, in vivo biocompatibility can be evaluated through subcutaneous implantation in immunocompetent mice, followed by histological analyses (e.g., hematoxylin and eosin staining) to detect cytoarchitecture differences and immunohistochemistry to analyze immune cell infiltration and expression of inflammatory mediators, as previously described [19].
For evaluating volume retention after adipose tissue implantation, width and length measurements are commonly used to assess graft volume [20–22] (Figure 4). Additional methods include water displacement, magnetic resonance imaging (MRI), computed tomography (CT), and ultrasound. Immunostaining is employed to verify graft vascularity using endothelial markers (e.g., CD31, VEGF, bFGF, lectin) [23,24]. Laser speckle imaging (LSI) can be used to evaluate graft perfusion, while flow cytometry, MRI, ultrasound, and micro-CT provide further insights into vascularization by measuring endothelial marker expression at the single-cell level (see General Note 7). When using this model, sample sizes should be estimated based on a power analysis. It is recommended to include at least four biological and technical replicates per group in each experiment. Additionally, to minimize bias, blinding of the animals or experimental groups is advised.
Figure 4. Fat engraftment outcomes. Monitor the volume retention regularly between the day of engraftment and the final day post-engraftment. Remove the graft and evaluate volume retention, displacement, vascularization, and perfusion.
Validation of protocol
This protocol has been used and validated in the following research article:
Das, et al. [19]. Injectable pulverized electrospun poly(lactic-co-glycolic acid) fibers improve human adipose tissue engraftment and volume retention. Journal of Biomedical Materials Research Part A (Figures 1 and 3).
General notes and troubleshooting
General notes
General note 1: Engraftment outcomes can be affected by the source of adipose tissue, with decreased cell proliferation being linked to factors such as increasing donor age, higher body mass index, diabetes mellitus, and other comorbidities [25].
General note 2: To ensure maximum cell viability, keep tissue samples on ice unless otherwise specified.
General note 3: While this study focused on the application of PLGA fibers, the protocol can be adapted to substitute or incorporate other materials, allowing for modified properties.
General note 4: Consider mouse characteristics such as age, sex, and housing status (single vs. group) as these factors may influence outcomes. This procedure is most commonly performed on mice aged 6–9 weeks. It is important to evaluate the immunoreactivity profile using immunocompetent mice. In this study, we used wildtype C57/BL6 (Jackson Laboratories, stock #000664) mice to test the immunoreactivity of PLGA fibers via subcutaneous implantation. Once the immunoreactivity was evaluated, male athymic nude mice (Jackson Laboratories, strain #002019) were used due to their immunodeficient nature, which allows for the study of human tissue engraftment without immune response interference [26]. Evaluating the immune response is crucial since it could lead to cell death, inflammation, and calcification of the engrafted fat tissue, complicating the study of fat engraftment.
General note 5: While the duration of the procedure and recovery period may vary, we recommend anticipating approximately 21 min from the initial induction of the mouse with isoflurane (step D2) until the completion of the procedure (step D16). Post-procedure, the mouse should be awake and alert within 3–5 min of the postoperative recovery period (step D18).
General note 6: In this study, human adipose tissue was engrafted into a mouse model. It is important to note that outcomes may vary as different adipose tissue sources or engraftment models are used, as the results are specific to human adipose tissue engraftment in mouse models.
General note 7: One limitation of this study is the evaluation period of adipose tissue engraftments. In this study, evaluations were conducted over 21 days, but a longer period can fully assess the host's response.
Acknowledgments
Schematics were created with BioRender.com. Funding for this work was partly provided by NIH grants (DP1DK126199, DP2 EB028110–01 to D.G.P).
Competing interests
Jed Johnson is a co-founder of Nanofiber Solutions, LLC. (Columbus, OH).
Ethical considerations
All animal experiments were performed in accordance with protocols approved by the Laboratory Animal Care and Use Committee at The Ohio State University (IACUC # 2016A00000074-R1 and IACUC # 2009A0006-R3).
All studies involving human adipose tissue were completed in accordance with a protocol that was approved by the Institutional Review Board (IRB) no. 180822–2. Informed consent was obtained by all human subjects prior to study.
References
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Article Information
Publication history
Received: Jul 22, 2024
Accepted: Sep 23, 2024
Available online: Oct 17, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,118 | https://bio-protocol.org/en/bpdetail?id=5118&type=0 | # Bio-Protocol Content
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Utilizing the Planar Lipid Bilayer Technique to Investigate Drosophila melanogaster dMpv17 Channel Activity
SC Samantha Corrà
YK Yevheniia Kravenska
VC Vanessa Checchetto
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5118 Views: 293
Reviewed by: Philipp A.M. Schmidpeter Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in iScience Oct 2023
Abstract
The planar lipid bilayer (PLB) technique represents a highly effective method for the study of membrane protein properties in a controlled environment. The PLB method was employed to investigate the role of mitochondrial inner membrane protein 17 (MPV17), whose mutations are associated with a hepatocerebral form of mitochondrial DNA depletion syndrome (MDS). This protocol presents a comprehensive, step-by-step guide to the assembly and utilization of a PLB system. The procedure comprises the formation of a lipid bilayer over an aperture, the reconstitution of the target protein, and the utilization of electrophysiological recording techniques to monitor channel activity. Furthermore, recommendations are provided for optimizing experimental conditions and overcoming common challenges encountered in PLB experiments. Overall, this protocol highlights the versatility of the PLB technique in advancing our understanding of membrane protein function and its broad application in various fields of research.
Key features
• This protocol leverages the planar lipid bilayer (PLB) technique to study the Drosophila Mpv17 protein (WT and mutant forms), revealing its role in forming ion channels.
• The use of a PLB apparatus and advanced electrophysiological recording equipment ensures precise and accurate measurement of ion channel activity.
• The PLB allows for the study of ion channel mechanics and regulation without interference from other proteins, ensuring the purity and accuracy of the results.
Keywords: Electrophysiology Planar lipid bilayer Ion channels dMpv17
Graphical overview
Background
Electrophysiological methods are of pivotal importance in the study of the dynamic functions and physical properties of ion channels, which are essential for cellular signaling and homeostasis [1–5]. Among these methods, the planar lipid bilayer (PLB) technique is held in high regard for its precise analysis of individual ion channel characteristics in a controlled artificial environment [6–8]. This method allows for a detailed investigation of ion channel mechanics at the molecular level and the direct assessment of the effects of various chemicals on these channels, without interference from regulatory proteins present in natural membranes.
The principal advantage of PLB experiments is their capacity to monitor and quantify ion transport across membrane channels, a crucial aspect of cellular physiology. These experiments are conducted in a specially designed chamber comprising two distinct compartments, the cis- and trans-compartments. The aperture between the compartments typically ranges from 50 to 250 μm in diameter, which is necessary for forming a planar bilayer membrane through which lipid-soluble ion channel proteins are integrated. This integration can occur directly from solutions or via fusion with liposomes. Once integrated, the system can measure ionic currents and membrane potentials generated by an electrochemical driving force, providing critical insights into ion conductance and channel activity.
While the PLB method is invaluable for elucidating the intricate relationship between molecular components and physical factors that regulate mammalian ion channels, it does present certain challenges (Table 1). The aperture size can result in prolonged voltage response times and elevated noise levels due to the extensive bilayer area. Nevertheless, contemporary PLB systems with low-noise, high-bandwidth recordings are gradually alleviating these issues, thereby enhancing measurement resolution and precision. Furthermore, the development of sophisticated electrophysiological setups and high-quality synthetic lipids has simplified the formation of artificial membranes and improved reproducibility. However, the variability in membrane proteins, which constitute the ion channels, remains a significant challenge, often determining the success of experimental outcomes.
In a previous study, we investigated the role of the Drosophila melanogaster ortholog of MPV17 (dMpv17) [9]. In humans, mutations in MPV17 are a prominent cause of pediatric-onset hepatocerebral form of mitochondrial DNA depletion syndromes (MDS) (OMIM: 266810) [10–14], including Navajo neurohepatopathy (NHH) [15]. MPV17-dependent MDS is characterized by the rapid deterioration of hepatic function and severe early-onset hypoglycemia, later complicated by neurological impairment. NHH is particularly prevalent in the Navajo population of North America, being caused by the founder R50Q mutation. NHH is characterized by a combination of peripheral neuropathy, liver disease, and other systemic dysfunctions resulting from mitochondrial DNA depletion. The syndrome can manifest in infancy, childhood, or adulthood but is generally associated with longer survival [16]. However, the precise role of MPV17 remains elusive despite decades of intensive research.
Using the planar lipid bilayer (PLB) technique, we investigated the ion channel activity of dMpv17. We showed that dMpv17 exhibited a conductance of 330 pico Siemens (pS) and displayed a preference for cations over anions, like the human counterpart. Furthermore, we demonstrated that the channel facilitated the translocation of uridine but not orotate. Finally, we showed that disease-associated mutations differently impact on channel activity.
This study provides a detailed protocol for PLB setup, expression and reconstitution of ion channel proteins on prepared membranes, ion translocation recordings, and channel current analysis.
Table 1. Key features, benefits, and limitations of PLB. PLB offers customizable membrane compositions and allows the reconstitution of purified proteins and channels, enabling detailed electrophysiological measurements. They provide easy shineaccess to both sides of the bilayer for various assays, making them versatile for a wide range of studies. However, precise control of lipid composition and protein reconstitution can be challenging. PLBs are sensitive to electrical noise and artifacts, and their fragility can lead to experimental failures. Additionally, they require specialized equipment and expertise.
Feature Benefits Limitations
Membrane composition Customizable to study various lipid compositions Requires precise control of lipid composition
Reconstitution of proteins Allows incorporation of purified proteins Protein reconstitution can be challenging
Electrophysiological measurements Enables detailed ion channel activity analysis Sensitive to electrical noise and artifacts
Accessibility Easy to access both sides of the bilayer for assays Bilayer fragility can lead to experimental failure
Experimental versatility Suitable for a wide range of electrophysiological studies Requires specialized equipment and expertise
Materials and reagents
L-α-phosphatidylcholine (Sigma-Aldrich, catalog number: P5638)
1,2-diphytanoyl-sn-glycero-3-phosphocholine (4ME 16:0 PC) (Avanti Polar Lipids, catalog number: 850356P)
Decane (Sigma-Aldrich, catalog number: 457116)
Chloroform (Sigma-Aldrich, catalog number: 372978)
Octane (Sigma-Aldrich, catalog number: 296988)
Acetone (Sigma-Aldrich, catalog number: 179124)
Insulin syringes (Biosigma, catalog number: BSS133/A)
Amber glass screw top vials (1.8 mL) (DWK Life Sciences, catalog number: 224750)
Nitrogen stream
Sterile syringe filter with 0.22 μm (Starlab, catalog number: E4780-1226)
Wheat Germ CECF Kit (Biotechrabbit, catalog number: BR1402501)
pIVEX 1.3 WG vector (Biotechrabbit, catalog number: BR1401301)
GenScript SurePAGE, Bis-Tris (GenScript, catalog number: M00658) and Tris-MOPS-SDS running buffer (GenScript, catalog number: M00138)
Bio-Rad Mini-Protean Tetra Cell System
In-Fusion® HD Cloning kit (Takara, catalog number: 639650)
KCl (Sigma-Aldrich, catalog number: P9541)
Tris (Sigma-Aldrich, catalog number: T1503)
KOH (Sigma-Aldrich, catalog number: P5958)
Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: 05030)
N,N-Dimethyl-n-dodecylamine N-oxide (LDAO) (Sigma-Aldrich, catalog number: 40236)
Uridine (Sigma-Aldrich, catalog number: U3750)
Sodium hypochlorite (Sigma-Aldrich, catalog number: 1056142500)
Primers (Thermo Fisher):
Forward: CCACAACAGCTTGTCGAACCATGAAGAGACTTAAAGCGTA
Reverse: TGATGATGAGAACCCCCCCCGCTATTAAGTATCATGGAGAG
QuickChange II Site-Directed Mutagenesis Kit (Agilent, catalog number: 200523)
R41Q: GAAACGGAGGGTTTGCCCCGCATCCCA and TGGGATGCGGGGCAAACCCTCCGTTTC
S163F: CCCGCTATTAAGTATCATGAAGAGGTAGCAGTTCCATAC and GTATGGAACTGCTACCTCTTCATGATACTTAATAGCGGG
Proteins
WT dMpv17
S163F dMpv17
R41Q dMpv17
Solutions
Bathing solution (pH 7.4) (see Recipes)
10% SDS (see Recipes)
10% LDAO (see Recipes)
1 M KCl (see Recipes)
Uridine (see Recipes)
NaClO (see Recipes)
Recipes
Note: See General note 1.
Bathing solution (pH 7.4)
Reagents Concentration of working solution Amount Final volume Comment
KCl 150 mM 1.68 g 150 mL Final volume of solution in the PLB cuvette is 3 mL
Tris 10 mM 0.18 g 150 mL
KOH 100% Some drops to adjust pH
10% SDS
Reagents Concentration of stock solution Amount Volume of stock solution Comment
SDS 10% 1 g 10 mL The final concentration in the experiment is 2%
10% LDAO
Reagents Concentration of stock solution Amount Final volume of stock solution Comment
LDAO 10% 1 g 10 mL The final concentration in the experiment is 2%
1 M KCl
Reagents Final concentration Amount Total with the final volume
KCl 1 M 37.275 g 500 mL
Uridine
Reagents Concentration of stock solution Amount Final volume of stock solution Comment
Uridine 200 mM 1.221 g 25 mL The final concentration in the experiment is 10 mM
NaClO
Reagents Final concentration
Sodium hypochlorite 100%
Equipment
General equipment
50 mL tubes (Corning, catalog number: 430828)
Refrigerated tabletop centrifuge (Hettich, model: MIKRO 220/220R centrifuge)
Vacuum system and vacuum gas pump (VWR Mini Laboratory Pump, VP 86, catalog number: 181-0067P)
Thermomixer (Eppendorf ThermoMixer C, catalog number: EP5382000031)
PLB Equipment (Figure 1)
Figure 1. Bilayer workstation. (A) The bilayer workstation used for dMpv17 experiments. (B) Graphical representation of bilayer workstation. The planar lipid bilayer workstation consists of (1) a Faraday cage positioned on an anti-vibration isolation table (2), (3) a bilayer clamp amplifier (e.g., BC-525, Warner Instruments Corporation), (4) a digitizer (e.g., Digidata 1322A, Axon Instruments), a 16-bit device connected to the current and voltage outputs and to a data acquisition system, and (5) a personal computer with an acquisition program (e.g., pCLAMP program sets) for data acquisition. The central part of the workstation is the headstage holder system (6), where the chamber and cuvette system are placed on a magnetic stirrer. Created with BioRender.com.
Faraday cage with vibration isolation table (see General note 2) (Figure 2)
Figure 2. Planar lipid bilayer chamber and cuvette. (A) The lipid bilayer chamber and cuvette, which is inserted into the trans-cavity. The small hole visible on the cuvette is the aperture where the lipid bilayers are formed. Lipid bilayers are painted on the aperture in the center of the cuvette. (B) Graphical representation of the lipid bilayer chamber with the inserted cuvette. The labeled parts are (1) input, (2) reference, (3) salt bridges, (4) Ag-AgCl electrodes, (5) trans side, (6) cis side, (7) magnetic stir bar. Schematic representation of the formation of a lipid bilayer using the "lipid painting" technique. The pipette deposits lipids onto an aperture between two compartments, cis and trans, facilitating the formation of the lipid bilayer. An air bubble aids in the even distribution of the lipids. The close-up shows the structure of the lipid bilayer, with hydrophilic heads facing outward and hydrophobic tails inward. Created with BioRender.com.
Bilayer clamp amplifier (Warner Instruments Corporation, model: BC-525)
Digitizer (Axon Instruments, model: Digidata 1322A, 16-bit) (see General note 3)
Bilayer clamp probe (see General note 4) (Figure 2) (Warner Instruments)
Bilayer chamber classic 22 mm chamber (3 mL volume) BCH-M22 (Warner Instruments, catalog number: W4 64-0453)
Classic 22 mm cuvettes Delrin with 250 μm aperture CD22A-250 (Warner Instruments, catalog number: W4 64-0411 (see General note 5)
Homemade salt bridges (see General note 6)
Ag-AgCl electrodes (Warner Instruments, catalog number: WA 10-5, 64-1327)
Magnetic stir bar and magnetic micro stirrer (VELP SCIENTIFICA, catalog number: F203A0161)
5 mm Teflon coated magnets (Warner Instruments, catalog number: 64-0420)
Personal computer equipped with software for data acquisition and analyses
Software and datasets
Clampfit 8 and/or 10.7.0.3 (Molecular Devices LLC)
ORIGIN 6.1 (OriginLab)
Graph Pad Prism 9.0.0 (GraphPad Software LLC)
Procedure
Lipid solution preparation
In the dMpv17 studies, experiments were conducted using two types of lipids: L-α-phosphatidylcholine and 4ME 16:0 PC. L-α-phosphatidylcholine and 4ME 16:0 PC were selected as lipid sources for PLB formation due to their biophysical properties, which closely resemble natural biological membranes. L-α-phosphatidylcholine, a major component of cell membranes, provides a stable lipid bilayer that is essential for effective channel reconstitution and functional analysis. Additionally, 4ME 16:0 PC was chosen to facilitate the formation of a more uniform and consistent planar lipid bilayer, which is critical for accurate ion channel measurements. These lipid choices help to create experimental conditions that closely replicate the natural environment of mitochondrial membranes, leading to more physiologically relevant results.
L-α-Phosphatidylcholine
Purification: Partially purify L-α-phosphatidylcholine by precipitating it with cold acetone from a chloroform solution. Make sure to perform this step in a fume hood to manage the organic solvents safely. Store at 4 °C for a maximum of one month.
i. Dissolve phosphatidylcholine in a glass vial using a minimal amount of chloroform. Use glass vials, as plastic can release plasticizers into apolar solvents like chloroform, which may contaminate your sample.
ii. Prepare a tube with 40 mL of cold acetone.
iii. Slowly add the phosphatidylcholine solution drop-by-drop into the acetone while stirring.
iv. Centrifuge the mixture for 5 min.
v. Discard the supernatant and redissolve the pellet in the same volume of chloroform used initially.
vi. Add the phosphatidylcholine solution to a second tube filled with acetone.
vii. Continue stirring and centrifuge for another 5 min; then, discard the supernatant.
viii. Spread the clean phosphatidylcholine on the tube walls, stirring under a nitrogen flow for 5 min to remove most of the acetone.
ix. Cover the tube with aluminum foil and dry it overnight using a vacuum pump.
x. Scrape the solid lipid from the tube walls using a glass scraper.
xi. Seal the purified phosphatidylcholine in glass vials under a nitrogen atmosphere.
xii. Store the lipid solutions at 4 °C and degas with a nitrogen stream each time the vial is opened. The solutions are stable for up to one month.
Dissolving
i. Dissolve the purified L-α-phosphatidylcholine in a mixture of n-decane and chloroform (in a 100:1 ratio) to achieve a final concentration of 10 mg/mL. Use a clean, dry pipette to add the solvents.
ii. Gently dry down the solution using nitrogen or air in a fume hood to avoid contamination.
Storage
i. Transfer the dissolved lipids into amber glass vials to protect from the light.
ii. Ensure that the vials are sealed tightly to prevent evaporation.
iii. Keep the dissolved lipids at 4 °C for a maximum of three days.
4ME 16:0 PC Lipids
Dissolving: Dissolve pure 4ME 16:0 PC lipids in n-octane at a final concentration of 10 mg/mL. Always conduct the handling and mixing of organic solvents in a fume hood to avoid exposure to harmful vapors.
Storage
i. Store the dissolved lipids in amber glass vials.
ii. Ensure vials are well-sealed.
iii. Keep at -20 °C. Under these conditions, stability can be maintained for several months, up to six months or more, provided that the container is well-sealed to prevent solvent evaporation.
dMPV17 protein preparation
Clone the dMpv17 coding sequence into the pIVEX 1.3 WG vector using the In-Fusion® HD Cloning Kit and following the manufacturer’s instructions (see General note 7).
Introduce the different point mutations in the dMpv17 sequence using the QuickChange II Site-Directed Mutagenesis Kit, following the manufacturer’s instructions.
Perform in vitro expression using an RTS100 Wheat Germ CECF Kit as described in [16–18].
Solubilize the expressed protein with 2% LDAO for 90 min at 30 °C with shaking at 1,400 rpm.
After solubilization, centrifuge the mixture at 20,000× g for 20 min at room temperature.
To control protein expression and solubilization, use 1–5 μL of the supernatant for analysis on SDS-polyacrylamide gels.
Perform western blotting with an anti-His antibody (see General note 8).
Apply the resulting supernatant to the planar bilayers to evaluate channel activity (see General note 9). Pipette the resulting supernatant containing dMpv17 protein directly into the experimental chamber, which contains the preformed planar lipid bilayer. Take care not to disturb the bilayer during the pipetting process to ensure the integrity of the experimental setup. Freshly prepared dMpv17 protein can be stored at 4 °C for up to four days without significant loss of activity.
PLB formation
Remove the vial containing the lipid solution from the refrigerator and allow it to reach room temperature.
Dry the white cuvette using a stream of air or nitrogen for a few minutes, ensuring not to touch or contaminate the aperture (Figure 2A).
Using a micropipette, pretreat the aperture by applying 0.3 μL of lipid solution directly onto the outer side of the aperture and allow it to dry for 5 min.
Repeat this process on the inner side, performing the procedure three times for each side.
Insert the clean white cuvette into the clean black chamber and place the chamber holder inside the Faraday cage.
Secure the chamber in the holder.
Ground the trans chamber.
Attach the electrodes. Ag-AgCl electrodes are composed of a silver wire coated with a thin layer of silver chloride. The electrodes are then placed in the experimental chambers (cis and trans) and connected to the recording apparatus via agarose bridges filled with 1 M KCl. It is important to ensure that the electrodes are correctly positioned and that they have good contact with the solution. The electrodes are then chloritized by immersing them in a solution of NaClO. When the electrodes are not in use, they should be stored in a chloride-containing solution.
Slowly add the working medium to both the cis and trans chambers (see General note 10).
To form the bilayer, a small air bubble is created using a pipette and used to paint, meaning to distribute the lipid film across the surface of the aperture. The movement of the air bubble over the aperture is crucial as it ensures the lipid film is spread evenly over the entire surface (Figure 2B). This process is essential for forming a thin and stable lipid bilayer, which is fundamental for ensuring the proper functionality of the system and the reproducibility of electrophysiological measurements.
Monitor the bilayer formation by measuring the capacitance.
Check the quality of the created membrane (see General note 11).
Once the bilayer is formed, apply a transmembrane potential using Ag/AgCl electrodes connected to the bath via agarose bridges filled with 1 M KCl (for additional information, refer to Section D) (Figure 2B).
Verify the stability of the membrane and the absence of impurities by applying both negative and positive voltages (from ± 20 to ± 100 mV) for at least 5 min before proceeding.
Carefully add the protein-containing supernatant to the cis side of the bilayer chamber.
Allow sufficient time for the protein to incorporate into the bilayer and assess its activity by measuring ionic currents or other relevant parameters. The insertion of the protein into the bilayer can take from a few seconds to several minutes. In many cases, signs of protein activity can be observed within 60 s to 15 min after adding the protein. It is recommended to place the magnetic micro stirrer beneath the two-part bilayer chamber to facilitate effective stirring. To activate the micro stirrer to maintain continuous agitation of the solutions, ensure consistent and uniform mixing in both compartments. If no activity is observed within this time frame, adjustments to the protein concentration or stirring conditions might be necessary to optimize incorporation.
Preparation of curved agar bridges
Bend the glass capillaries (with diameters from 1.0 to 2.0 mm) at a right angle by applying a flame at approximately one-third of the length from the end.
Weigh 1 g of agarose for every 100 mL of solution, place it in a 200 mL PYREX glass container, and melt it in the microwave. Heat the solution until it begins to bubble. After heating, add KCl. Do not place KCl directly in the microwave, as this could cause sparks.
Once the 1 M KCl in 1% agarose is heated, inject the solution into the capillaries using a syringe, ensuring no bubbles are present in the bridges.
Allow the capillaries to dry.
Trim the ends of the tubes with a diamond cutter.
Store the agar bridges in 1 M KCl at 4 °C.
Channel activity recording
Open the Clampfit program.
Access the File menu and select Open Data from the toolbar.
Choose the appropriate file and its saving location.
Set the appropriate filter and the sampling rate.
Prepare the membrane and check for noise and/or extraneous signals (see General note 12).
Ensure the membrane is intact and unbroken.
Add the purified protein to the cis compartment and allow it to integrate into the planar bilayer.
Stir the contents of both chambers using magnetic stir bars.
Record channel activity continuously at various voltages (from -100 to +100 mV) at room temperature.
Perform single-channel recordings under symmetrical K+ conditions (150 mM KCl in both cis and trans sides) or under asymmetrical K+ conditions (400 mM KCl in the cis side and 135 mM KCl in the trans side).
Measurement of dMpv17 activity in the presence of uridine
Fill both compartments (cis and trans) of the bilayer chamber with the bathing solution.
Place a magnetic micro-stirrer beneath the chamber to ensure effective stirring throughout the experiment.
Add the prepared and solubilized dMpv17 sample to the cis compartment.
Measure the channel activity at a specific potential (e.g., -80 mV).
Add uridine to the cis compartment to reach a final concentration of 10 mM in the cuvette. The uridine used in the experiment is a dilution from a more concentrated stock solution.
Record the channel activity after adding uridine.
Collect a sample from the trans compartment to analyze the uridine translocation process.
Measure the concentration of uridine in the trans compartment using mass spectrometry (see General note 13).
Data analysis
See General note 14.
Open Clampfit 8 or 10.7.0.3 software.
Filter the recordings using the Filter function in the Analyze menu of the Clampfit toolbar.
Navigate to the Single-Channel Search in the Event Detection tab.
Verify that the baseline of the recordings is strictly horizontal and consistent at both the beginning and end of the recording.
Ensure that the recording duration is adequate for precise data analysis. For dwell time values, maintain channel activity for at least 60 s. If the channel shows extended periods of closure, increase the recording time. For Po values, a recording duration of a few tens of seconds may be sufficient, as long as there are no prolonged closure periods.
Single-channel search in Clampfit
The Single-Channel Search function allows to identify and analyze the activity of individual ion channels in a bilayer lipid membrane or similar experimental setups.
Import the data
Choose the Single-Channel Search option located in the Event Detection menu of the Clampfit.
Choose File > Open Data. Open the file containing the single-channel recording data.
Preprocess the data
Filter the data: Apply any necessary filters to remove noise and baseline drift. This can be done using the Filter option under the Analyze menu.
Perform single-channel search (Figure 3A)
Navigate to Analyze > Single-Channel Search. The Single-Channel Search dialog box will appear. The range of interest is marked with vertical cursors 1 and 2, ensuring segments of equal length are selected for all processed signals. The moving horizontal cursor 0 is set to the baseline, representing the complete closure level of all channels on the signal. Each subsequent line (numbered 1, 2, 3, etc.) is formed using the Make function on the left menu and set to the open level of each corresponding channel. This method allows the amplitude in Amperes (the current) of each channel to be immediately visible in the Level window.
Run the search (Figure 3B)
The search starts using the double arrow Nonstop on the top toolbar, and the results are displayed in the Event Statistics window in the Event Detection button of the toolbar. By controlling the up and down arrows, all necessary information for all levels (channels) is obtained.
Review and analyze events
After the search is complete, an event list will be generated showing all detected single-channel events. Review the event list to ensure that all significant events have been correctly identified (Figure 3B).
Navigate to Analyze > Histogram to generate histograms and analyze the dwell-time distributions. Use Clampfit's fitting functions to fit models to the dwell-time data.
Save and export results
Use File > Save Event List to save the list of detected events.
Export the analyzed data and histograms using File > Export.
Figure 3. Interface of the Clampfit 10.7.0.3 program, single-channel search mode. (A) The print screen demonstrates preparation for the Single-Channel Search function on the exemplary dMpv17 activity recording. Vertical cursors 1 and 2 limit the segment of interest. The baseline is set to the closed state of the channel, and “level 1” line is set to the open one. The left menu shows the signal amplitude in nanoampere (Level 1), and selected options for baseline and Search Region. This recording represents a single-channel activity. (B) Print screen of the single-channel search result on the dMpv17 activity recording. The Single-Channel Statistics window displays the search data. In the right-hand corner, the Event Viewer window (in green) makes it possible to evaluate the quality of the search for single events and take further actions with them. Directly on the recording, the program denotes detected channel closures with blue lines, and its openings with red lines.
Open probability in Clampfit
Open probability value for each level can be taken from the Event Statistics window as was shown above (see point A). Analyzing multiple channels with the same conductance, the parameter NPo (the number of open channels multiplied by the open probability) is used. The open probability for each channel can be determined using the formula:
P
o
=
N
P
o
N
where:
NPo is the average number of open channels multiplied by the open probability.
N is the total number of channels.
The primary sources of information and solutions to technical issues are the PDF Manual and the Clampfit Help option found in the Help tab on the toolbar.
Plotting the I-V graph
To plot a current-voltage (I-V) graph of a lipid bilayer, the current-voltage relationship for this type of structure must first be understood. In lipid bilayers, the current is generally determined by the movement of ions through ion channels or pores formed in the bilayer. The I-V relationship can be linear or nonlinear, depending on the specific characteristics of the bilayer and the ion channels present. Under symmetrical K+ solutions (see step E10), the expected current through the dMpv17 is about -20 pA at -60 mV. Accordingly, the channel conductance is 330 pS. Under asymmetrical K+ solutions, the I-V plot is expected to show a current range from around -40 to +70 pA at a voltage of -100 to +100 mV [9].
Measure the current (the amplitude of the signal, see point A). The amplitude should preferably be calculated manually to avoid distortion from automatic calculations, which may include short openings or substates. To ensure consistency, all available records should be processed in the same manner.
Record the data: Note the current value for each applied voltage.
Plot the graph: The collected data are used to plot a graph with voltage (V) on the x-axis and current (I) on the y-axis.
The obtained values are statistically analyzed using built-in functions in ORIGIN 6.1 and GraphPad Prism. These programs are also used for creating graphs and figures. Each data group should be obtained from three or more independent experiments to ensure reliability.
Validation of protocol
In our recent publication, we applied this step-by-step protocol to investigate the role of the MPV17 of Drosophila melanogaster [9]. We investigated the ion channel activity of dMpv17 using the PLB technique, exploiting recombinant, LDAO-solubilized in vitro–expressed dMpv17 protein with a His-tag at the C-terminus. In the presence of K+, channel opening was detected with a predominant single-channel conductance of 330 picoSiemens (pS). Like the human protein, dMpv17 exhibited a slight preference for cations over anions and demonstrated a linear current-voltage relationship.
Additionally, we showed that the reconstituted channel translocates uridine but not orotate, suggesting a possible role for this protein in the translocation of key metabolites across the mitochondrial membrane.
Lastly, we examined whether pathogenic mutations have a significant impact on the channel's slope conductance. We selected the conserved Ser-170 (S163F in Drosophila), one of the three predicted phosphorylation sites whose mutation causes MDS, and Arg-50 (R41Q in Drosophila), associated with NHH syndrome, as candidate amino acids for further analysis. The S163F mutation prevented channel opening under physiological conditions, while the R41Q mutation led to a significant reduction in conductance but did not completely abolish it.
General notes and troubleshooting
General notes
All solutions must be prepared using ultrapure water and analytical-grade reagents. It is imperative that solutions are freshly prepared and stored in a refrigerator or at -20 °C. To reduce the risk of contamination, it is recommended that all buffers be filtered through a 0.22 μm filter.
It is recommended that the system be isolated from external electromagnetic fields using a Faraday cage. It is imperative that the Faraday cage is of sufficient size to encompass all electronic components and that it is properly grounded.
The digitizer should be connected to the amplifier. It is essential to ensure that the digitizer is correctly configured to align with the amplifier's output, as it is responsible for transforming the analog signals from the amplifier into digital data for subsequent computer analysis.
The headstage should then be secured in the holder system. It is essential to position the headstage correctly and to connect it to the amplifier, as this component is integral to the configuration and serves as the interface with the sample.
The PLB chamber and cuvette are to be assembled using a black Delrin chamber and a white polystyrene cuvette with a precision-machined aperture. For further details on the cleaning process, please refer to the comprehensive instructions provided by Warner Instruments at www.warneronline.com.
The preparation of salt bridges entails the dissolution of 1 M KCl in agarose, which is then allowed to solidify in a suitable mold. It is imperative that the salt bridges be handled with the utmost care to prevent contamination and ensure that they are devoid of any air bubbles. The salt bridges should be positioned between the electrodes and the experimental chambers (cis and trans), ensuring correct placement and optimal contact with the solution.
The dMpv17 coding sequence should be cloned into the pIVEX 1.3 WG vector using the In-Fusion HD Cloning Kit, in accordance with the instructions provided by the manufacturer.
It is crucial to perform a western blot or other quality control methods to assess the quantity and quality of the expressed and solubilized protein. The precise amount required may depend on the efficiency of protein expression and purification processes.
The protein concentration used in bilayer experiments can vary depending on the type of protein and the specific aim of the experiment. Typically, the concentrations of channel proteins used in PBL experiments are in the nanomolar (nM) or picomolar (pM) range. A typical concentration may be between 0.1 and 1 µg/mL, but this can be adjusted depending on the size, behavior, and activity of the specific protein being studied. To achieve optimal results, it is common to perform preliminary experiments to optimize the protein concentration so that channels form in the bilayer without oversaturating the system.
It is imperative that all voltages from the cis chamber are duly recorded. In our experiments, zero is assigned to the trans (grounded) side. It is recommended that positive currents be considered to indicate the flow of cations from the cis compartment to the trans compartment.
It is recommended that approximately 150–200 pF capacity be used for bilayer experiments. The measured capacitance of 150–200 pF aligns with the aperture sizes used in our experimental setup. Specifically, larger apertures, approaching 250 µm, tend to yield higher capacitance due to the increased surface area available for forming the lipid bilayer. Since the capacitance of the lipid bilayer is directly proportional to the bilayer's surface area, larger apertures result in higher capacitance values, as observed in our study. We believe that this range of capacitance accurately reflects our experimental conditions and is representative of the aperture sizes employed in our setup.
It is essential to eliminate any extraneous signals that may impair the quality of the graphics and impede the normal analysis of channel activity. If the noise amplitude exceeds 10% of the anticipated channel activity, it is imperative to implement measures to eliminate the noise during the recording process (for further details, please refer to General Note 2).
Samples were centrifuged at 12,000× g for 10 min (Hettich Mikro 120 Benchtop Centrifuge) and directly analyzed. The UHPLC-HRMS (ultra-high performance liquid chromatography–high resolution mass spectrometry) system was equipped with an Agilent 1260 Infinity II LC liquid chromatographer coupled to an Agilent 6545 LC/Q-TOF mass analyzer (Agilent Technol-ogies, Palo Alto, CA, USA). The analytical column was an Atlantis Premier BEH C18 AX 2.5 μm, 150 × 2.1 mm (Waters, Milano, Italy), kept at 25 °C. The mobile phase components A and B were water containing 15 mM acetic acid and 50 mM ammonium acetate and acetonitrile, respectively. The eluent flow rate was 0.25 mL/min. The mobile phase gradient profile was as follows (t in min): t0–3 0% B; t3–18 0–100% B, t18–20 100% B; t20–30 0% B. The MS conditions were: electrospray (ESI) ionization in negative mode, gas temp 225 °C, drying gas 13 L/min, nebulizer 35 psi, sheath gas temp 350 °C, sheath gas flow 12 L/min, VCap 3,500 V, nozzle voltage 0 V, and fragmentor 125 V. Centroid full scan mass spectra were recorded in the range 100–1,000 m/z with a scan rate of 1 spectrum/s. MS/MS spectra were obtained by selecting the precursor with an isolation window of 1.3 m/z and collision energy of 30 eV. MS and MS/MS data were analyzed using the Mass Hunter Qualitative Analysis software (Agilent Technologies, Palo Alto, CA, USA).
The identity of each analyte was confirmed by MS/MS data, while the chromatographic peak integration was performed by selecting the extracted ion chromatogram (EIC) of the [M-H]- species related to analytes of interest with a window of 5 ppm. Quantification of analytes was carried out by external calibration using a five-point calibration curve obtained by spiking the buffer solution with each analyte in the range of 10–100 nM. Linearity showed R2 > 0.98 and the limit of detection of the method was assessed as approximately 5 nM. Samples were analyzed in duplicate, and results were reported as mean values. Relative standard deviation (RSD) of reported data was always lower than 20%.
For further information and solutions to technical issues, please refer to the PDF manual and the Clampfit Help option, which can be found in the Help tab on the Clampfit toolbar.
Acknowledgments
This protocol has been used and validated in the following research article: Corrà et al. [9], Drosophila Mpv17 forms an ion channel and regulates energy metabolism. 428 iScience, 2023. 26(10): p. 107955. https://doi.org/10.1016/j.isci.2023.107955. Research on MPV17 by the Checchetto Group is supported by PRIN 2022 (grant number 2022ZY7ATN). S.C. expresses gratitude for the fellowship provided by Prof. Massimo Zeviani (Telethon grant number GMR23T1065). Y.K. expresses gratitude for her grant provided by the Seal of Excellence (MSCA), grant number SZAB_MSCASOE21_01.
We extend our sincere thanks to Prof. Ildikò Szabò for her invaluable assistance with data analysis and for granting us access to her PLB setups, which were crucial for conducting all experiments.
Competing interests
The authors declare no competing interests.
References
Zoratti, M. and Szabó, I. (1994). Electrophysiology of the inner mitochondrial membrane. J Bioenerg Biomembr. 26(5): 543–553.
Paggio, A., Checchetto, V., Campo, A., Menabò, R., Di Marco, G., Di Lisa, F., Szabo, I., Rizzuto, R. and De Stefani, D. (2019). Identification of an ATP-sensitive potassium channel in mitochondria. Nature. 572(7771): 609–613.
Raffaello, A., De Stefani, D., Sabbadin, D., Teardo, E., Merli, G., Picard, A., Checchetto, V., Moro, S., Szabò, I. and Rizzuto, R. (2013). The mitochondrial calcium uniporter is a multimer that can include a dominant-negative pore-forming subunit. EMBO J. 32(17): 2362–2376.
De Stefani, D., Raffaello, A., Teardo, E., Szabò, I. and Rizzuto, R. (2011). A forty-kilodalton protein of the inner membrane is the mitochondrial calcium uniporter. Nature. 476(7360): 336–340.
Patron, M., Checchetto, V., Raffaello, A., Teardo, E., Vecellio Reane, D., Mantoan, M., Granatiero, V., Szabò, I., De Stefani, D. and Rizzuto, R. (2014). MICU1 and MICU2 Finely Tune the Mitochondrial Ca2+ Uniporter by Exerting Opposite Effects on MCU Activity. Mol Cell. 53(5): 726–737.
Zakharian, E., (2013). Recording of Ion Channel Activity in Planar Lipid Bilayer Experiments. Methods Mol Biol. 109–118.
Polak, A., Mulej, B. and Kramar, P. (2012). System for Measuring Planar Lipid Bilayer Properties. J Membr Biol. 245(10): 625–632.
Harsman, A., Bartsch, P., Hemmis, B., Krüger, V. and Wagner, R. (2011). Exploring protein import pores of cellular organelles at the single molecule level using the planar lipid bilayer technique. Eur J Cell Biol. 90(9): 721–730.
Corrà, S., Checchetto, V., Brischigliaro, M., Rampazzo, C., Bottani, E., Gagliani, C., Cortese, K., De Pittà, C., Roverso, M. and De Stefani, D. (2023). Drosophila Mpv17 forms an ion channel and regulates energy metabolism. iScience. 26(10): 107955.
Spinazzola, A., Invernizzi, F., Carrara, F., Lamantea, E., Donati, A., DiRocco, M., Giordano, I., M., Meznaric‐Petrusa, Baruffini, E. and Ferrero, I. (2008). Clinical and molecular features of mitochondrial DNA depletion syndromes. J Inherited Metab Dis. 32(2): 143–158.
Spinazzola, A., Viscomi, C., Fernandez-Vizarra, E., Carrara, F., D'Adamo, P., Calvo, S., Marsano, R. M., Donnini, C., Weiher, H. and Strisciuglio, P. (2006). MPV17 encodes an inner mitochondrial membrane protein and is mutated in infantile hepatic mitochondrial DNA depletion. Nat Genet. 38(5): 570–575.
AlSaman, A., Tomoum, H., Invernizzi, F. and Zeviani, M. (2012). Hepatocerebral form of mitochondrial DNA depletion syndrome due to mutation in MPV17 gene. Saudi J Gastroenterol. 18(4): 285.
El-Hattab, A. W., Wang, J., Dai, H., Almannai, M., Staufner, C., Alfadhel, M., Gambello, M. J., Prasun, P., Raza, S. and Lyons, H. J. (2018). MPV17-related mitochondrial DNA maintenance defect: New cases and review of clinical, biochemical, and molecular aspects. Hum Mutat. 39(4): 461–470.
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Article Information
Publication history
Received: Jul 29, 2024
Accepted: Sep 17, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biophysics > Electrophysiology
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Display of Native SARS-CoV-2 Spike on Mammalian Cells to Measure Antibody Affinity and ADCC
RW Rebecca E. Wilen
AN Annalee W. Nguyen
AQ Ahlam N. Qerqez
JM Jennifer A. Maynard
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5119 Views: 439
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Original Research Article:
The authors used this protocol in eLIFE Mar 2023
Abstract
The COVID-19 pandemic led to the rapid development of antibody-based therapeutics and vaccines targeting the SARS-CoV-2 spike protein. Several antibodies have been instrumental in protecting vulnerable populations, but their utility was limited by the emergence of spike variants with diminished susceptibility to antibody binding and neutralization. Moreover, these spike variants exhibited reduced neutralization by polyclonal antibodies in vaccinated individuals. Accordingly, the characterization of antibody binding to spike variants is critical to define antibody potency and understand the impact of amino acid changes. A key challenge in this effort is poor spike stability, with most current methods assessing antibody binding using individual domains instead of the intact spike or variants with stabilizing amino acid changes in the ectodomain (e.g., 2P or HexaPro). The use of non-native spike may not accurately predict antibody binding if changes lie within the epitope or alter epitope accessibility by altering spike dynamics. Here, we present methods to characterize antibody affinity for and activity against unmodified SARS-CoV-2 spike protein variants displayed on a mammalian cell membrane that recapitulates the native spike environment on infected cells. These include a flow cytometry–based method to determine the effective antibody binding affinity (KD) and an antibody-dependent cellular cytotoxicity (ADCC) assay to assess Fc-mediated activities. These methods can readily evaluate antibody activity across a panel of spike variants and contribute to our understanding of spike/antibody co-evolution.
Key features
• Allows rapid characterization of antibody binding to native SARS-CoV-2 spike on the mammalian cell surface
• Describes analysis of antibody binding to multiple native spike variants without stabilizing mutations
• Describes analysis of Fc-mediated antibody-dependent cellular cytotoxicity
• Requires transient transfection of Expi293F and 293T cells to assess antibody binding and ADCC, a flow cytometer for antibody binding, and a plate reader for ADCC
• Protocol is readily adaptable to other viral fusogens and membrane proteins
Keywords: SARS-CoV-2 Antibody affinity KD Flow cytometry Spike protein Native spike variants ADCC
Graphical overview
Background
There is widespread interest in therapeutics to treat COVID-19 by targeting the spike protein of SARS-CoV-2 (hereafter referred to as “spike”). This type I viral fusogen mediates fusion between the virion and host cell membranes, necessary for infection, and is the primary target of the SARS-CoV-2 antibody response [1]. Between 2020 and 2023, six different spike-binding antibody therapeutics received emergency use authorization from the FDA but exhibited reduced activity against subsequent spike variants whose residue changes altered antibody binding and neutralization [2]. These variants were also less susceptible to neutralization by polyclonal antibodies elicited by vaccination with the original Wuhan-Hu-1 spike variant [3]. Antibody binding affinity and ability to recruit Fc-dependent immune responses against different spike variants can determine antibody potency, the biochemical impact of spike changes, and implications for protection.
However, analyses of antibody/spike interactions are complicated by the structure-function relationships of this large, homotrimeric protein. Each spike protomer comprises a conserved S2 stalk and a more variable receptor-binding S1 subunit. Host cell infection is initiated when the receptor binding domain (RBD) within S1 engages an ACE2 receptor on a host cell. Since the RBDs are primarily present in a “down” position, which shields the ACE2 binding site from immunological surveillance, this interaction requires an RBD to transiently sample the “up” state in order to be accessible for receptor binding [4]. The RBD/ACE2 interaction triggers a cascade of spike conformational changes: The S2’ cleavage site in the S2 subunit is exposed, allowing cell surface proteases (typically TMPRSS2 and cathepsin L) access to cleavage sites. Proteolysis releases the S1 subunit, freeing the hydrophobic fusion peptide to embed itself in the target cell membrane. The S2 domain then collapses from a meta-stable pre-fusion state into a thermodynamically more stable post-fusion conformation, resulting in the fusion of the viral and target cell membranes [5]. These structural rearrangements present challenges for the analysis of antibody/spike interactions as epitopes can be occluded or exposed depending on the immediate state of the spike protein.
Biochemical and immunological characterization of antibody/spike interactions is key to understanding spike antigenicity, elucidating mechanisms of protection and supporting the development of potent antibody therapeutics. However, the spike variant used, experimental conditions, and any changes to the spike sequence (e.g., stabilizing mutations, truncations to support soluble expression, or glycosylation differences) can impact the relevance of experimental measurements to human infection. Mutations throughout the spike protein can directly impact the ratio of spike found in “open” or “RBD-up” forms vs. “closed” or “RBD-down” forms, the ratio of pre- vs. post-fusion spike conformations, and antibody access to epitopes buried deeply in S2 [6–8]. In particular, low endosomal pH values (pH ~5–6) drive spike toward a more “closed” conformation, which can shield neutralizing epitopes [7,9]. Storage temperature can also influence spike stability—hydrogen-deuterium exchange mass spectrometry data indicate that incubation at 4 °C induces a reversible conformation shift toward an “open” state that exposes portions of the S2 interface [6,10]. The range of spike conformations sampled during evaluation can impact antibody binding and interpretation of mechanisms of antibody-mediated protection.
Unfortunately, the native spike protein is unstable when expressed solubly, resulting in very low yields of aggregation-prone protein and presenting challenges for biochemical analyses. The introduction of two stabilizing proline substitutions (K986P and V987P, also called the “2P” variant), was essential for determining the first cryo-EM structure of SARS-CoV-2 spike [11]. These 2P changes were subsequently included in multiple approved vaccines and vaccine candidates since they dramatically improve protein yield and storage stability [11,12]. A second-generation spike called HexaPro comprises the 2P changes plus four additional proline substitutions to further increase the yield and stability of the pre-fusion spike [13]. Additionally, most forms of soluble spike protein replace the native transmembrane region with a foldon domain, which may impact global protein dynamics [14]. While stabilized spike variants have been pivotal for enabling spike research, they also introduce non-native mutations. Since antibody binding is a result of both direct epitope–paratope interactions and indirect epitope accessibility, these stabilizing changes can impact effective antibody affinity by indirectly altering global spike dynamics as well as directly altering the epitope sequence. As a result, antibody binding results with stabilized spike variants may not accurately reflect interactions with native spike.
Strategies to characterize spike-binding antibodies have primarily used isolated spike subunits or stabilized spike variants. These approaches fuse the native RBD to the surface of yeast [15,16] or display truncated spike variants (such as the isolated RBD) or stabilized intact spike variants on mammalian cells [17,18], which allows researchers to measure antibody binding in a high throughput manner. However, these data reflect antibody binding to fully exposed RBDs, which may not predict binding to intact spike on the viral or infected cell surface. Other reports fused stabilized spike trimers to non-native transmembrane domains including the PDGFR homodimer, which may also impact antibody binding behavior [18]. Lentiviral pseudovirus assays using native spike on the surface of VSV-G-based pseudoviruses and ACE2 expressing 293T or Vero E6 target cells have been successfully used to evaluate antibodies [19–21]; however, these assays are complex and require increased safety precautions.
In addition to antibody binding, Fc-mediated effector functions are increasingly recognized for their contributions to protection. The antibody-dependent cellular cytotoxicity (ADCC) activity of anti-spike antibodies has been reported with similar methods that rely on different reporter or target cells than those used in this protocol [22,23]. Chen et al. used T-REx-293 cells stably expressing EGFP, luciferase, and codon-optimized native spike with the same NK-92 cell line used below. Hong et al. used Expi293F cells stably expressing red fluorescent protein and native spike with engineered Jurkat-luciferase NFAT-CD16 cells. While these protocols are effective in monitoring ADCC activity, they may require special user-created, engineered stable cell lines. As an alternative, this protocol demonstrates ADCC differences between anti-spike antibodies using two well-characterized human cell lines (293T and NK-92 cells) and transient transfection.
We recently described antibodies that exhibit cross-reactive binding across SARS-CoV-1, SARS-CoV-2, and MERS-stabilized spikes [24]. The epitope recognized, spanning residues 985–1001 near the HR1/CH hinge in the pre-fusion spike S2 domain, is shielded in closed spike conformations and overlaps with the stabilizing “2P” changes. The antibodies analyzed include 3A3, which binds stabilized but not native SARS-CoV-2 Wuhan-Hu-1, SARS-CoV-1, and MERS spikes, and its engineered derivative RAY53, which binds both stabilized and native spike from SARS-CoV-2 Wuhan-Hu-1 and MERS and the stabilized form of SARS-CoV-1. Although both antibodies bind soluble, stabilized spike from these three β-coronaviruses with <10 nM affinity, the cryptic location of this epitope at the S2 apex overlapping the 2P residues suggests that the stabilizing changes alter antibody binding. To understand antibody interactions with this epitope in the context of native spike, we expressed spike with its native transmembrane domain and no stabilizing changes on the surface of Expi293F or 293T cells. In this protocol, native spike-expressing cells are used to measure the effective binding affinity, adapted from previously described protocols [25,26], and to assess susceptibility to Fc-mediated ADCC, using a cytotoxic assay that evaluates antibody ability to clear infected cells (Figure 1).
Figure 1. Overview of method. IA) Expi293F cells are transfected with EGFP- and spike-expression plasmids and incubated for two days at 37 °C. IB) Cells are washed and stained with anti-spike antibodies bearing human constant domains followed by detection with anti-human-Fc fluorescent antibody. IC) Cells are analyzed on a flow cytometer to assess anti-spike antibody binding by geometric mean fluorescence intensity (GMFI). IIA) 293T cells are grown, transfected with EGFP and spike-expressing plasmids, and incubated for 2 days at 37 °C. IIB) NK-92 cells are grown with recombinant human IL-2. IIC) Spike-expressing 293T cells are loaded with calcein-AM dye and then co-incubated with NK-92 cells and anti-spike antibodies at 37 °C. IID) After a 4 h co-incubation, the release of fluorescent calcein into the media from lysed target cells is measured using a plate reader. Created in BioRender. Wilen, R. (2024) BioRender.com/s02t921
Materials and reagents
Biological materials
Expi293F cells (Thermo Fisher Scientific, catalog number: A14527)
293T cells (Millipore Sigma, catalog number: 12022001)
NK-92 cells expressing the high-affinity CD16a allele V158 (Brink Biologics, catalog number: haNK CD16-V158.NK-92.05; formerly, ATCC, catalog number: PTA-8836)
Reagents
Expi293F Transfection Kit (Thermo Fisher Scientific, catalog number: A14524). Includes ExpiFectamine 293 reagent
pHDM vector SARS-CoV-2 Wuhan-Hu-1 Spike glycoprotein (BEI Resources, catalog number: NR-52514); referred to here as pWT-SARS-2
pEGFP plasmid as described in Nguyen et al. [27]
Control anti-spike antibody S309 with human constant domains (constructed based on Pinto et al. [28]; also called Sotrovimab and commercially available from Thermo Fisher, catalog number: MA5-42316)
Note: Select a control antibody binding a highly accessible epitope available both in the “open” and “closed” spike conformations, such as S309/Sotrovimab, which strongly binds SARS-CoV-1 and SARS-CoV-2 variants Wuhan-Hu-1 through Omicron XBB.1.5 and BQ.1.1. However, S309/Sotrovimab poorly binds spike variants starting after variants CH.1.1 and CA.3.1 [29].
Anti-human-Fc-PE antibody or equivalent anti-human-Fc secondary antibody (Jackson ImmunoResearch, catalog number: C840K53)
Lipofectamine 3000 kit (LifeTech, catalog number: L3000008)
Recombinant human IL-2 (Millipore Sigma, catalog number: SRP3085-50UG)
Dulbecco’s phosphate buffered saline (PBS) (Millipore Sigma, catalog number: D8537)
Fetal bovine serum (FBS) (Gibco, catalog number: A5267-01)
Penicillin-Streptomycin (Millipore Sigma, catalog number: P4458)
Inositol (Millipore Sigma, catalog number: I7508)
2-mercaptoethanol (Millipore Sigma, catalog number: M6250)
Folic acid (Millipore Sigma, catalog number: F8758)
Heat-inactivated horse serum (Millipore Sigma, catalog number: H1270)
Triton-X (Fisher Scientific, catalog number: BP151-100)
Sodium dodecyl sulfate (SDS) (Fisher Scientific, catalog number: BP166-500)
NaCl (Millipore Sigma, catalog number: S9888-10KG)
EDTA, disodium salt dihydrate (Fisher Scientific, catalog number: S311)
Trypan blue (MP Biomedicals, catalog number: 02195532-CF)
Expi293F expression media (Thermo Fisher Scientific, catalog number: A1435101)
OptiMEM (Gibco, catalog number: 31985070)
DMEM media (Millipore Sigma, catalog number: D5796)
Alpha minimum essential media (Thermo Fisher Scientific, catalog number: 12561056)
Expi293F Transfection Kit protocol (Thermo Fisher Scientific, catalog number: A14524, publication number: MAN0007814)
Solutions
Staining buffer (see Recipes)
293T growth media (see Recipes)
Serum-free 293T growth media (see Recipes)
NK-92 growth media (see Recipes)
Lysis buffer (see Recipes)
Recipes
Staining buffer
500 mL of PBS
1% v/v FBS
293T growth media
440 mL of DMEM
10 mL of penicillin-streptomycin
50 mL of FBS
Serum-free 293T growth media
490 mL of DMEM
10 mL of penicillin-streptomycin
NK-92 growth media
500 mL of Alpha minimum essential medium without ribonucleosides and deoxyribonucleosides
0.2 mM inositol
0.1 mM 2-mercaptoethanol
0.02 mM folic acid
200 U/mL human IL-2 to start cultures, 100 U/mL for maintenance media
12.5% heat-inactivated horse serum
12.5% heat-inactivated FBS
Lysis buffer
2% Triton-X
1% SDS
100 mM NaCl
1 mM EDTA
Laboratory supplies
96-well, U-bottom plate (Fisher Scientific, catalog number: 07-200-95)
Note: Can be replaced by flow tubes or microcentrifuge tubes.
96-well, black plate with a flat, clear bottom (Fisher Scientific, catalog number: 07-200-625)
1.5 mL microcentrifuge tubes (Fisher Scientific, catalog number: 05-408-129)
Pipette tips (1000, 200, and 10 μL) (Fisher Scientific, catalog number: 02-707-401, 02-708-416, 02-707-437)
Hemocytometer (Sigma-Aldrich, catalog number: Z375257)
125 mL shaking flask for Expi293F (Fisher Scientific, catalog number: PBV12-5)
Micropipettes (Fisher Scientific, catalog number: F167380)
Equipment
Biosafety cabinet (Thermo Scientific, catalog number: 1326122CON)
Incubator (37 °C, 8% CO2, shaking) for Expi293F (Infors HT, catalog number: I80002)
Incubator (37 °C, 5% CO2, stationary) for 293T and NK-92 (Eppendorf, catalog number: 6734010015)
Microscope (Olympus, catalog number: CKX41SF)
Tabletop centrifuge (Eppendorf, catalog number: 5405000441)
Vacuum trap system (BrandTech, catalog number: 20727403PM)
BD Fortessa LSR Cytometer or equivalent with at least two-color capability (BD Biosciences)
Microplate reader capable of reading fluorescence (excitation: 488, emission: 515) (Agilent BioTek, catalog number: SH1M-SN)
Software and datasets
FlowJo or similar FSC analysis software (version 10.7.1)
GraphPad Prism or similar graphing/analysis software such as Microsoft Excel (version 9.5.0)
Procedure
Part I. Determine effective antibody KD
Transfect Expi293F cells for spike display (day 0–1)
Stabilization of the membrane-bound spike protein enhances overall stability and expression but also alters the spike conformational dynamics. Here, we describe the expression of native spike protein on the mammalian cell surface, which better mimics the dynamics of native protein on SARS-CoV-2-infected cells. Expi293F cells are transiently transfected with a plasmid encoding the entire spike ectodomain and its native transmembrane domain. Although the plasmid was originally designed for use in packaging lentivirus with SARS-CoV-2 spike on the viral surface, it works well for direct transient expression in mammalian cells. Because liposomal transfection results in multiple plasmids entering each cell, co-transfection with an EGFP-expressing plasmid is used to identify successfully transfected cells. After two days of expression, cells can be stained with anti-spike antibodies and fluorescent secondary antibodies to analyze antibody binding (section B).
Note: This protocol is adapted from the Expi293F Transfection Kit protocol.
Thaw and grow Expi293F cells for at least one week after thawing.
Count cells and determine viability.
Dilute a sample of cells 10-fold with PBS (i.e., 100 μL cells + 900 μL PBS).
Undiluted cells should be approximately 3–5 × 106 cells/mL so diluted cells will be approximately 3–5 × 105 cells/mL, a more manageable amount to manually count.
Mix 10 μL of diluted cells with 10 μL trypan blue.
Use a hemocytometer and inverted microscope or automated cell counter to determine cell count and viability.
Note: Cell viability should be >95%. If viability is lower, re-seed cells for maintenance and proceed at a later date.
Determine the number of cells required for the experiment and seed 1.5× that number at 2.5 × 106 cells/mL.
We recommend staining 3 × 105 cells/sample in microcentrifuge tubes or per well of a 96-well plate.
Example: 3 × 105 cells/sample × (6 concentrations + 3 controls) × 2 replicates = 5.4 × 106 cells needed × 1.5 overage = at least 8.1 × 106 mock-transfected and spike-transfected cells to seed at 2.5 × 106 cells/mL.
We recommend transfecting 1.5× the number of cells needed for the experiment to account for excess; however, the exact number transfected may need to be adjusted to account for available tissue culture plates.
Example: 18 samples require 3.25 mL of cells at 2.5 × 106 cells/mL, which can be rounded up to 4 mL to fit available tissue culture plates (i.e., 2 wells with 2 mL each in a 6-well plate)
Incubate cells at 37 °C and 8% CO2 with shaking at 125 rpm.
Next day, repeat step A2 to determine cell count and viability.
Note: Viability should be >95%. If cell viability is less than 95%, do not proceed and repeat steps A1–4.
Dilute cells to 2.5 × 106 cells/mL and seed the appropriate number of cells calculated in step A3.
Example: 18 samples calculated above require approximately 4 mL of culture at 2.5 × 106 cells/mL.
Calculate the amount of DNA and other reagents necessary for the experiment.
pEGFP plasmid: 0.5 μg/mL of culture to transfect.
pWT-SARS-2 plasmid: 0.5 μg/mL of culture to transfect.
OptiMEM: 60 μL/mL of culture to transfect.
Fresh OptiMEM is needed to dilute both the DNA and ExpiFectamine, so two aliquots of OptiMEM should be prepared per transfection.
ExpiFectamine 293: 3.2 μL/mL of culture to transfect.
Example: For 4 mL of cell culture to be transfected, use 2 μg of pEGFP plasmid, 2 μg of pWT-SARS-2 plasmid, 240 μL of OptiMEM for DNA dilution, 240 μL of OptiMEM for ExpiFectamine dilution, 12.8 μL of ExpiFectamine 293.
Combine pEGFP plasmid and pWT-SARS-2 plasmid in OptiMEM. Use the EGFP plasmid only for a mock transfection control lacking spike expression to evaluate antibody specificity.
Mix by gently pipetting up and down, inverting the tube, or swirling.
Note: DNA should be at a concentration ≥ 1 μg/mL, treated with an endotoxin removal column to remove residual endotoxin from plasmid purification, and sterile-filtered (0.2 μm) to prevent contamination of the Expi293F cultures.
Dilute ExpiFectamine 293 reagent in fresh OptiMEM.
Mix by gently pipetting up and down.
Incubate at room temperature for 5 min.
Note: The ExpiFectamine dilution can be pooled into a single tube at this step and aliquoted to each transfection (mock, SARS-CoV-2 spike, other spike variants) in the next step.
Add the diluted ExpiFectamine (step A10) to the diluted DNA (step A9) and mix gently by pipetting, swirling, or inverting the tube. Incubate the mixture for 10–20 min at room temperature.
Add DNA–ExpiFectamine 293 mixture dropwise to the diluted cells from step A6 and gently swirl to mix.
Incubate cells in an incubator at 37 °C and 8% CO2 with shaking at 125 rpm for 48 h.
Note: Expi293F transfection typically includes adding enhancers to the cell culture the day following transfection. This addition is optional. We have not noticed that the addition of enhancers results in a difference in surface spike expression levels two days after transfection.
Flow cytometry staining (day 3)
The Expi293F cells expressing native spike protein (Step A) can be used to characterize anti-spike antibodies. Anti-spike antibodies are serially diluted and incubated with spike-expressing Expi293F cells. These are then washed, incubated with a secondary detection antibody, washed again, and analyzed by flow cytometry as described in section C. We recommend several key controls including (i) no primary antibody to assess non-specific binding by the secondary antibody used (hereafter referred to as “secondary only”), (ii) a negative isotype control antibody with the same constant domains as the test antibody but different variable regions such that it does not bind spike, and (iii) a positive control antibody that is known to bind spike. It is important to choose a positive control antibody (such as S309) that binds an epitope available on various coronaviruses as well as an epitope that is on a portion of the SARS-CoV-2 spike accessible in various conformations (“up” vs. “down” vs. “open”).
The Langmuir isotherm analysis used to measure the effective KD requires that the soluble binding partner (here, antibody) always be present in excess. This allows the experiment to comply with the model assumption that the free antibody concentration remains constant throughout the experiment, regardless of the amount of antibody/ligand complex formed. At low antibody concentrations, this may require the use of large volumes to provide a greater number of antibodies than spike proteins. Appropriate volumes can be determined from a simple experiment: Stain an equal number of cells with a single, low antibody concentration in different final volumes (e.g., 10 nM antibody in 50 μL, 100 μL, 500 μL, 1 mL, and 5 mL) before staining with secondary antibody and flow cytometric analysis. If the number of antibody molecules limits antibody/spike complex formation, the GMFI of the stained cells will increase with increasing volume. The smallest volume resulting in maximal observed GMFI should be used for affinity measurements.
The protocol below references the example plate layout in Figure 2. Briefly, antibody dilutions are prepared in the preparative Plate A at 2× final concentration [sufficient volume for at least 4 wells (>100 μL): two replicates each of two cell types at 25 μL of antibody solution per well]. In the assay Plate B, 25 μL of cells are aliquoted into each well, and then 25 μL of serially diluted antibody from Plate A is added for a 1:1 dilution to the final cell and antibody concentrations. Typical final cell concentration is 6 × 106 mL at 50 μL per well, while the antibody range will depend on antibody affinity; however, 300 nM to ~1 nM is recommended for the example antibody used here, RAY53.
On day 2 post-transfection, count the transfected or mock-transfected cells using a hemocytometer or automated cell counter.
Determine the number of cells needed for the experiment and transfer 1.5× the calculated number to a 15 mL conical tube.
We recommend staining 3 × 105 cells per sample (50 μL of final volume at 6 × 106 cells/mL final concentration) in a 96-well plate; however, this may need to be optimized depending on the flow cytometer used.
Example: 3 × 105 cells per sample × (6 concentrations + 3 controls) = 5.4 × 106 cells needed × 1.5 overage = at least 8.1 × 106 cells transferred to a conical tube.
Wash twice with staining buffer.
Centrifuge at 250× g for 5 min.
Aspirate media.
Resuspend cells in at least ~600 μL of staining buffer per 106 cells.
Example: 8.3 × 106 cells should be resuspended in at least ~5,000 μL of staining buffer.
Repeat step B3a.
Note: Washing can lead to cell losses. The recommended resuspension volume should lead to a cell concentration of ~1.6 × 107 cells/mL with no cell losses, which is above the final concentration required, allowing the user to dilute the cells to their final concentration and reducing the need for additional spins to prepare the cells.
Count cells and dilute to 1.2 × 107 cells/mL in staining buffer.
Dilute cells 20-fold (i.e., 5 μL of cells + 95 μL of PBS) to ~8 × 105 cells/mL.
Mix 10 μL of diluted cells with 10 μL of trypan blue.
Use a hemocytometer and inverted microscope or automated cell counter to determine cell count and viability.
Dilute test antibody and controls in a fresh dilution plate (Figure 2A, Plate A).
Prepare enough antibody solution for at least two replicates at 25 μL per replicate plus 20 μL excess.
Note: Multiply the number of replicates by the number of cell types tested. For example, two replicates and two cell types should be treated as four replicates worth of antibody needed. The following protocol describes two replicates and two cell types (four replicates worth of antibody) for each antibody concentration.
For four replicates: Start with 120 μL of staining buffer per well (25 μL × 4 replicates + 20 μL excess = 120 μL). For a 5-fold serial dilution, 30 μL will be added, mixed, and transferred to the subsequent dilution.
Prepare the highest antibody concentration at 600 nM (2× the final desired highest concentration) and serially dilute 5-fold at least six times.
i. Add 120 μL of staining buffer to wells A2–A7 of Plate A (repeat in additional row per antibody testing).
In the example in Figure 2A, Antibody 1 is diluted in row A. Antibody 2 is diluted in row C, so 120 μL of staining buffer was also added to wells C2–C7.
ii. Add 150 μL of 600 nM antibody to well A1.
In the example in Figure 2A, 150 μL of 600 nM Antibody 2 should be added to well C1.
iii. Transfer 30 μL from A1 to A2 and mix thoroughly >6 times.
In the example in Figure 2A, 30 μL from C1 should also be transferred to C2 and mixed thoroughly.
iv. Repeat, transferring 30 μL of A2 to A3 and mixing thoroughly, 30 μL of A3 to A4, etc., until A7.
In the example in Figure 2A, this should be repeated by serially diluting from C2 to C7.
Note: Assay controls include:
i. Positive control antibody to confirm spike display: Samples stained with S309 or other positive control antibody to detect spike on the cell surface (25 μL per replicate well at 10 nM).
ii. Secondary-only control: Sample not stained with an anti-spike antibody (no antibody control/secondary only control) (25 μL of staining buffer).
iii. Cells-only control: Sample lacking both anti-spike primary antibody and secondary (25 μL of staining buffer).
Note: Concentrations used may need to be optimized for different antibodies. We recommend initial tests starting 10–50× above the expected KD. If the affinity is unknown, we recommend starting at 300 nM final concentration. The dilution factor can also be adjusted for each antibody to encompass the full curve.
Aliquot 25 μL of diluted cells (3 × 105 total cells) per sample in a fresh 96-well plate (Figure 2B, Plate B).
Transfer 25 μL of stain from the stain dilution plate (step B5, Plate A) to appropriate wells of cells (step B6, Plate B), mix gently by pipetting, and incubate for 1 h on ice. In this example, the stain prepared in wells A1–A9 in plate A is added to wells A1–A9, B1–B9, E1–E9, and F1–F9 in plate B. The stain prepared in wells C1–C8 in plate A is added to wells C1–C8, D1–D8, G1–G8, and H1–H8 in plate B.
Wash twice with staining buffer.
Centrifuge at 250× g for 5 min.
Aspirate media.
Resuspend cells in 200 μL of staining buffer.
Repeat steps B8a and B8b.
Incubate cells with 50 μL of 1:250 goat anti-human-Fc-PE in staining buffer for 1 h on ice.
Note:
If using a different secondary antibody, the dilution factor may need to be optimized.
For cells-only control, it is appropriate to replace the secondary stain with 50 μL of staining buffer.
Repeat step B8 with the final resuspension in 200 μL of staining buffer or the volume required for analysis on the flow cytometer used.
Figure 2. Example plate layouts for antibody staining for Part I, section B (antibody KD, a–b) and Part II, section C (ADCC, c–d). A. The antibody is diluted in a fresh plate (Plate A) to create 2× concentrated master mixes with enough volume for all necessary replicates (25 μL per replicate plus excess to allow for pipetting). B. 3 × 105 total cells (25 μL, 1.2 × 107 cells/mL) are mixed with 25 μL of antibody master mix from Plate A in the staining plate (Plate B). C. Cells and antibodies for antibody-dependent cellular cytotoxicity (ADCC) are diluted in tubes to create concentrated master mixes with enough volume for all necessary replicates (4× concentrated for antibodies, 1 × 105 cells/mL for target cells, 2 × 106 cells/mL for effector cells). D. 1 × 104 target cells (100 μL, 1 × 105 cells/mL), 1 × 105 NK-92 cells (50 μL, 2 × 106 cells/mL), and 4× concentrated antibody (50 μL) are combined. Created in BioRender. Wilen, R. (2024) BioRender.com/c65x019
Note: Plates or tubes can be used in both panel A and panel C. Tubes may be preferred for simplicity; however, as the number of antibody samples increases, a plate allows for better organization.
Flow cytometry data acquisition (day 3)
Expi293F cells expressing native spike and stained with anti-spike antibodies are analyzed by multi-color flow cytometry. Cells are first gated by size (FSC vs. SSC), then for singlets (FSC-A vs. FSC-H), and finally for EGFP-positive cells to identify the transfected population. These EGFP-expressing cells are then assessed for antibody binding, which is monitored by fluorescence from the secondary antibody. Key controls include secondary antibody only (negative control), an isotype control antibody (negative control), and a cross-reactive anti-spike antibody (positive control) in addition to the antibody of interest. Confocal imaging confirms GFP expression and S309 binding to Wuhan-Hu-1 spike on the cell surface (Supplemental Figure 1).
Collect data on a flow cytometer.
Gating strategy:
i. Cells: FSC A vs. SSC A, gate main cell population.
ii. Singlets: FSC A vs. FSC H, gate only cells on the main diagonal; cells off the diagonal may be doublets or larger cell clumps.
iii. EGFP+ cells: EGFP vs. FSC A, gate only EGFP+ cells.
Collect data for 10,000 EGFP+ cells.
Note: Depending on individual instrument and filter settings, compensation may be necessary. Adjust voltages so that negative and positive samples appear on the screen and are not cut off by the cytometry software. Make sure to acquire EGFP and PE (or appropriate secondary fluorophore) fluorescence in addition to FSC and SSC channels.
Example gating strategy is shown in Figure 3.
Export either geometric mean fluorescence intensity (GMFI, used here) or median fluorescence intensity (MFI) for analysis to account for a non-normal distribution of points.
Proceed to the data analysis section for details on analysis to determine the effective KD.
Figure 3. Gating strategy to isolate spike-displaying cells. Example results and gating strategy to determine the effective KD. A) Example gating for secondary-only control (top), RAY53 staining of Expi293F cells only expressing EGFP (middle), and RAY53 staining of Expi293F cells expressing both EGFP and spike (bottom). The sample is first gated for cell size (FSC vs. SSC), singlets (FSC-A vs. FSC-H), and EGFP expression; then, the PE fluorescence (Ab binding) of the final gated population is analyzed. The reported PE GMFI over several concentrations can be used to directly determine the effective antibody KD. B) Example shown of positive and negative control histogram after gating. Negative control (secondary only) in black, and positive control (S309 antibody) in red.
Part II: Antibody-dependent cellular cytotoxicity
Transfection of 293T cells with spike and EGFP plasmids (day 0–1)
In a similar protocol to Part I, section A above (Expi293F transfection), adherent 293T cells are transfected with both a plasmid for native spike protein expression and a plasmid for EGFP expression using Lipofectamine 3000. 293T and Expi293F cells are variants of HEK 293 cells that have different levels of protein expression, and the 293T cell variant used for both the following and previous protocols may need to be optimized. Cell media is replaced the day after transfection to remove transfection reagents. Passage the 293T cells before they reach confluency, as these cells can easily form clumps that are difficult to break up into single cells. As in Part I, we recommend including a “mock” transfection control lacking DNA to assess the specificity of the ADCC response.
Passage 293T cells in 293T growth media as described [30].
Note: It is important not to let the cells overgrow (>90% confluency), as this can cause clumps that can cause high error in this assay.
Seed cells such that there is a single flask prepared on the day of transfection at 70%–90% confluency for each spike variant or control tested.
Example: To assess ADCC with cells expressing only SARS-CoV-2 Wuhan-Hu-1 spike, two flasks should be prepared—one for transfecting with pWT-SARS-2 plasmid and one for mock transfection as a negative control.
Calculate the number of target cells required for each spike variant and scale the transfection to at least five times that number to ensure sufficient cell numbers. The 293T cell line typically reaches 2–3 × 105/cm2 at 100% confluence. For each spike variant or control tested, triplicate wells containing 104 transfected 293T per well for each control (target only, spontaneous release, maximum lysis) and each antibody concentration will be required.
Example: To assess ADCC of SARS-CoV-2 Wuhan-Hu-1 spike transfected cells incubated with 3A3 or RAY53 at a single concentration, 15 wells of spike-expressing 293T cells will be required (three replicates each of 3A3 experimental well, RAY53 experimental well, target-only control, spontaneous release control, and maximum lysis control) and a minimum of 7.5 × 105 cells must be transfected. Although transfection of 70% confluent 293T cells in a single well of a 24-well plate (surface area ~2 cm2) would be sufficient, the recommended minimum transfection scale is a single well of a 6-well plate (~10 cm2) or a T25 flask (25 cm2) for ease of handling. We describe the transfection of a T25 flask of 293T cells here.
For a T25 flask with 4 mL of culture volume, seed 5 × 105 cells per milliliter of 293T cells in 4 mL of 293T growth media the day before transfection.
Dilute 1 μg of DNA per milliliter of culture volume and 2 μL of P3000 reagent per milliliter of culture volume into 50 μL of OptiMEM per milliliter of culture volume.
For a T25 flask with 4 mL of culture volume, combine 4 μg of DNA, 8 μL of P3000 reagent, and 200 μL of OptiMEM in a microcentrifuge tube.
Note: DNA stock should be at a concentration ≥ 1 μg/mL, treated with an endotoxin removal column to remove residual endotoxin from plasmid purification, and sterile-filtered (0.2 μm) to prevent contamination of the 293T cultures.
Dilute 2.5 μL of Lipofectamine 3000 reagent per milliliter of culture volume into 50 μL of OptiMEM per milliliter of culture volume.
For a T25 flask with 4 mL of culture volume, combine 10 μL of Lipofectamine 3000 and 200 μL of OptiMEM in a microcentrifuge tube.
Gently pipette diluted DNA/P3000 mixture into the tube containing diluted Lipofectamine 3000, mix gently by pipetting, swirling, or inverting the tube, and let sit for 10–15 min.
Slowly add the DNA/Lipofectamine 3000 mixture in a dropwise manner to the prepared flask of 293T cells.
Incubate at 37 °C overnight.
The next day, remove media and replace with fresh 293T growth medium.
Preparation of NK-92 cells (day 1–2)
Approximately 90% of human NK cells express the classical Fc receptor CD16A, which is activated by clustered antibodies on target cells to initiate ADCC activities, but initial attempts at immortalizing NK-92 cells resulted in loss of CD16A expression [31,32]. Accordingly, NK-92 cell lines engineered to stably express the high (V158) or low (F158) alleles of CD16A are a popular option to assess ADCC in vitro [31,33,34]. These lines require human IL-2 to support growth in vitro and constitutively express EGFP. Before seeding into the assay plates, cells are washed to remove residual IL-2 and bovine IgG from the media. Bovine IgG may bind to CD16 on the NK cells, reducing available receptors and limiting the final ADCC response.
Passage NK-92 cells using NK growth media described above, ensuring that sufficient healthy NK-92 cells will be available for the planned assay. Each well planned for the experiment will require 104 NK-92 cells. Plan to have at least three times the number required available on the day of the assay. Achieving adequate numbers of NK-92 cells can be a limiting factor in this experiment, so incorporating the NK-92 growth timeline in the experimental plan is essential.
Supplement media with 500 U human IL-2/mL initially upon thaw, followed by 250 U human IL-2/mL for subsequent passages.
Note: IL-2 is sensitive to freeze-thaw and should be aliquoted into small aliquots after reconstitution. Once thawed, the IL-2 aliquots can be stored at 4 °C for up to one week.
Grow NK-92 cells with the T-flask placed vertically.
During passaging, cell densities should range between ~8 × 104 cells/mL and ~3 × 105 cells/mL.
Cells may be clumpy during passaging. Pipetting up and down with a serological pipette of appropriate volume may gently disperse clumps as cells trapped in clumps will be poorly responsive in the assay.
One day after transfecting the 293T cells (day 1), centrifuge the NK-92 cells (250× g for 5 min) and aspirate the media.
Replace with fresh NK-92 media, add fresh IL-2 (250 U human IL-2/mL), and incubate overnight at 37 °C.
On the day of the assay (day 2), count the cells (Part I, section A2) and resuspend them in NK-92 growth media without FBS to a concentration of 2 × 106 cells/mL.
Note: Viability should be >95%. If cell viability is <95%, do not proceed and repeat Part II, section B1–3.
Setup of ADCC (day 2)
To measure ADCC in terms of target cell lysis, spike-expressing 293T target cells are loaded with acetoxymethyl ester calcein (calcein-AM) dye. This hydrophobic dye diffuses across the cell membrane where it is cleaved by intracellular esterases to its fluorescent calcein form. Upon lysis, cells release fluorescent calcein into the media. After removing the cells and debris, media are transferred to a new plate and the fluorescence is measured by a plate reader. When preparing the spike-expressing 293T target cells, gently pipette to dissociate cell clumps, which can reduce calcein-AM loading into cells and subsequent release by ADCC. The procedure described below uses a 10:1 effector:target cell ratio; however, this can be optimized as needed. While edge effects from evaporation are unlikely during the short incubation, common practice is to avoid using the outer wells of the plate (column 1, column 12, row A, and row H) and fill these wells with PBS or excess media. Key controls include (i) a spontaneous release control with target cells, NK-92 cells, and no antibody present, (ii) an NK-92-only control, (iii) a target cell–only control, and (iv) a maximum lysis control with target cells only and lysis buffer added ~10–15 min before reading the plate absorbance.
On day 2, remove media from 293T cells and wash with one culture volume of PBS.
For a T25, add 4 mL; for a T75, use 8–12 mL.
Repeat 2–3 times to remove non-adherent and dead cells.
Remove PBS, add one culture volume of PBS with 10 mM EDTA, and incubate for approximately 10 min or until cells detach from the flask surface. Gentle tapping on the side of the flask can help release the cells from the surface of the flask.
Note: Use EDTA rather than trypsin, as trypsin can cleave the spike proteins from the cell surface.
When cells are released from the plate (confirm by looking at the flask under an inverted microscope), add 2–3 mL of complete 293T growth media per milliliter of original culture and transfer to a 15 mL conical tube.
Spin cells at 250× g for 5 min, remove supernatant, and resuspend in serum-free 293T growth media.
Repeat steps IIC2–3 times to remove excess dead cells. Resuspend in serum-free 293T growth media after the final wash.
Note: Use enough media to resuspend the cells at ~1 × 106 cells/mL based on confluency and flask size. For example, one well of a 6-well plate at ~80% confluency should have approximately 1.6–2.4 × 106 cells, so the pellet should be resuspended in ~2 mL of 293T growth media.
Pipette cells up and down vigorously ~10 times with a 1 mL Pipetman or similar with a small opening to disrupt cell clumps.
Note: The presence of cell clumps causes several issues for this assay. First, they will result in incorrect cell counting and inaccurate E:T ratios. Second, calcein-AM staining may be inconsistent across cell clumps. Third, clumpy cells will not properly engage target cells. Together, these issues impact assay reproducibility.
Count cells, determine viability, and dilute in serum-free 293T growth media to a concentration of 0.5–1 × 106 cells/mL in a conical tube.
Note: If a large fraction of dead cells appears at this stage (greater than 10%), the likelihood of a successful assay is low; repeat or optimize steps IIC1–6.
Add calcein-AM to the transfected cells to a final concentration of 2 μM, invert the tube several times to mix, wrap in foil, and incubate at 37 °C for 30 min, inverting every ~10 min.
Wash transfected 293T cells three times with at least double the original culture volume of serum-free 293T growth media. For a T25, wash with at least 8 mL of complete serum-free 293T growth media.
Note: The cell pellet will appear green due to calcein.
Resuspend transfected 293T cells in 1 mL of serum-free 293T growth media, count, and dilute cells to 1×105 cells/mL.
If cells are less than 1 × 105 cells/mL, spin cells one more time and resuspend in appropriate volume.
Prepare 4× final concentration antibody stock in microcentrifuge tubes (Figure 2C).
For example, if a final concentration of 50 nM is required, the stock should be diluted to 200 nM.
To each well in a 96-well plate (Figure 2D, Plate D):
Add 100 μL of transfected 293T cells (at 1 × 105 cells/mL, 1 × 104 cells/well).
Use 100 μL of media only for the NK-92 only control wells.
Add 50 μL of antibody from the dilution plate to the assay plate.
Replace with media for spontaneous release control, but do not add media or antibody to max lysis control.
Add 50 μL of NK-92 cells (at 2 × 106 cells/mL, 1 × 105 cells/well for an E:T ratio of 10:1).
Do not add to max lysis control.
Max lysis control: 100 μL of transfected 293T cells only (1 × 104 cells/well). These wells will have half the volume of the other wells during this incubation step.
NK-92-only control: 50 μL of NK-92 cells (1 × 105 cells/well) + 150 μL of serum-free 293T growth media.
Spontaneous lysis control: 100 μL of transfected 293T cells (1× 104 cells/well) + 50 μL of NK-92 cells (1 × 105 cells/well) + 50 μL of serum-free 293T growth media.
In the final 200 μL of assay volume, antibody concentrations will be 50 nM, with an E:T ratio of 10:1 and 1.1 × 105 total cells.
Incubate plate (Plate D) at 37 °C for 4 h covered in foil.
ADCC readout and analysis (day 2)
After 4 h, the cells are centrifuged, and the media is transferred to a black plate. Calcein released into the media is detected via fluorescence on a plate reader.
Remove the plate from the incubator and carefully add 100 μL of lysis buffer to the maximum lysis control wells to normalize its volume to that of experimental wells. Any contamination of other wells with the lysis buffer will impact the results of the assay. Allow the plate to sit for 5–10 min at room temperature in the dark.
Spin the plate at 400× g for 10 min and transfer 100 μL of supernatant to a fresh black clear-bottom 96-well plate.
Using a plate reader, measure the fluorescence intensity at an excitation wavelength of 488 nm and emission wavelength of 515 nm.
Data analysis
Part I:
Upload your FSC files into FlowJo or an equivalent flow cytometry analysis software. Follow the gating scheme described above (Figure 3; Part 1, section C) to isolate the EGFP-expressing cell population. First, check for spike expression by analyzing positive and negative controls (S309 and secondary-only, respectively). Confirm controls are as expected (no shift for negative control and large shift for positive control) (Figure 3B); if not, do not proceed. An exception is in the case of a control known not to bind a particular spike variant; for example, antibody S309 can bind several SARS lineage β-coronavirus spikes but is known not to bind to the MERS spike.
To determine the effective antibody KD, export the GMFI for each sample into Microsoft Excel or another spreadsheet software. For each antibody concentration, normalize each sample by subtracting the average GMFI of the pEGFP-only transfected wells stained with the same concentration of antibody. Upload the resulting normalized GMFIs with their corresponding antibody concentrations into GraphPad Prism or other plotting software. Visualize the Langmuir isotherm plot: antibody concentration vs. PE GMFI of EGFP positive cells, Equation 1. The plot should include the full curve, with multiple points in the initial increase phase as well as at least one point showing maximum binding. If there are one or fewer points in each phase, repeat the experiment with more points (Figure 4).
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Where Ceq is the concentration of complex (antibody bound to receptor, in this case sample GMFI), Cmax is the maximum concentration of complex (here, the GMFI of our top concentration, ideally saturated), L0 is the concentration of ligand (here, the antibody), and KD is the equilibrium binding affinity. A key assumption of the Langmuir analysis is that the antibody is always present in excess of the number of spike proteins and thus is constant regardless of the number of complex molecules formed; the testing outlined in the description of Part I, section B determines whether this assumption is met.
Next, create a semi-log plot with the log of antibody concentration on the x-axis and the PE GMFI of EGFP positive cells on the y-axis using GraphPad Prism. Under Analysis, click Analyze and then Nonlinear regression (curve fit), Dose-response – Stimulation, and choose [Agonist] vs response (three parameters). This program uses a slight variation of the Langmuir isotherm (Equation 1) that accounts for a non-zero minimum GMFI, listed below,
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Y
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where minimum is the GMFI of the no-antibody control, maximum is the GMFI of the top concentration (Cmax), X is the concentration of antibody (L0), Y is the GMFI of the sample (Ceq), and EC50 is the effective equilibrium KD. The calculated KD will have the same units as the antibody concentration.
Figure 4. Determining effective KD from flow cytometry data. Data derived from the gating described in Figure 3 is transformed and plotted to determine data fits and effective KD of anti-spike antibodies. A) After gating the cytometer data as shown in Figure 3, example histograms show a fluorescence shift (GMFI) corresponding to different RAY53 antibody concentrations against cells displaying Wuhan-Hu-1 spike. B) The population GMFI from the histograms in A is plotted against antibody concentration to form a Langmuir isotherm. C) Data can also be plotted log-transformed and used to determine effective KD as described above. Plot taken from Silva et al. [24].
To examine antibody cross-reactivity and epitope accessibility, two comparisons are made. First, if the epitope is known, the epitope amino acid sequence for each variant can be compared. Second, the binding histogram and GMFI should be compared to the positive control (S309) for each variant and compared between the variant and Wuhan-Hu-1 spike (Figure 5A). Ideally, choose a positive-control antibody whose epitope is available in most spike conformations to measure spike expression level. We also highly recommend choosing a control antibody well-characterized in the literature and known to bind various spike variants.
Before testing experimental samples, confirm the expression of each spike variant with the positive-control antibody. The respective binding (GMFI) of controls to variant spikes can be compared to known affinities in the literature. For example, S309 is known to bind SARS-1 and SARS-2 but not MERS [28]. Each experimental sample can then be compared to the positive control for each respective spike variant (Figure 5B). Large differences in transfection efficiency and display (≥10%) can impact comparisons across spikes and transfection should be optimized to minimize these differences if needed.
If there is no direct change in the sequence of the antibody epitope, changes in binding are most likely due to conformational differences in the spike variants. If there are identified mutations in the epitope, it may be difficult to confidently distinguish between changes in epitope affinity and epitope accessibility. Conclusions that may be drawn from various results are described below in Table 1.
Figure 5. Antibody binding to cell surface–displayed spike variants. Example data comparing antibody binding to a panel of native coronavirus spike proteins. Each spike protein was expressed on the surface of Expi293F cells, stained with antibodies, and qualitatively characterized by flow cytometry. A) A histogram shows antibody binding to Wuhan-Hu-1 spike by the secondary-only negative control and S309 positive control and test antibodies 3A3 and RAY53. B) Binding of the 3A3 and RAY53 antibodies to cells displaying the indicated spike variants is compared by histograms of singlet cells. Differences in RAY53 binding and epitope accessibility can be evaluated by examining the loss of binding to spike with mutations in the epitope as described in Table 1.
Table 1. General guide to aid in determining the possible causes of antibody binding changes between spike variants.
Sequence comparison (variant vs. Wuhan-Hu-1) Binding comparison (variant vs. Wuhan-Hu-1) Cross-reactivity and epitope accessibility
No change No change Cross-reactive
No change Significantly reduced, non-zero Reduced epitope accessibility
No change No binding Severe or complete reduction in epitope accessibility
No change Significantly improved Cross-reactive, improved epitope accessibility
Mutation(s) No change Cross-reactive*
Mutation(s) Significantly reduced, non-zero Cross-reactive, potential loss of affinity*
Mutation(s) No binding Not cross-reactive*
Mutation(s) Significantly improved Cross-reactive*
*This analysis works best in cases where there is no direct change to the antibody epitope. In this case, observed changes in antibody binding can be attributed to differences in spike protein conformation and epitope accessibility. One caveat in evaluating cases with epitope changes is that an affinity loss can be offset by increased accessibility, or vice versa. This can make it difficult to discern whether changes are primarily due to affinity or epitope accessibility.
Part II:
In Microsoft Excel or another equivalent software, upload your fluorescence measurements from the plate reader. To calculate the percentage of target cells that were lysed through ADCC, use Equation 3 below:
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100
Experimental: Sample of interest (i.e., target cells + NK cells + 10 nM antibody).
Spontaneous release: Sample containing target cells and NK cells but no antibody.
Max release: Sample containing target cells and NK cells with lysis buffer added (no antibody).
Values should be between 0% and 100%, although some non-effective samples will result in minor negative values. Values ranging beyond 0%–100% imply issues with non-specific activity in the negative control or low responses in the positive control, while large error among replicates can be due to high experimental variation among replicates (e.g., due to pipetting errors) or a small dose-response range, which is very sensitive to small differences. Additional factors that can contribute include evaporation causing edge effects, incomplete lysis of the control lysis wells, or cell clumps leading to inconsistencies among wells. The calculated percentage lysis values can be graphed as in Figure 6, to visualize comparisons and facilitate statistical comparisons.
Figure 6. Analysis of antibody-dependent cellular cytotoxicity (ADCC) activity. ADCC activity (percentage target cell lysis) is compared between mock and spike-expressing target cells for an isotype antibody (grey), control antibody CR3022 (green), and RAY53 (blue). Two-way ANOVA was performed to compare samples with spike-displaying cells or mock-transfected cells for each antibody. ** denotes p < 0.1; ns, not significant. Figure from (Silva et al., 2023).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Silva et al. [24]. Identification of a conserved S2 epitope present on spike proteins from all highly pathogenic coronaviruses. eLife. Effective antibody KD: Figure 4, panel e, ADCC: Figure 5, panel d.
Each point was performed with 2–3 technical duplicates and each experiment was repeated 2–3 times. Antibody binding controls included a single stain with the control antibody S309 at 10 nM (maximum staining, data not shown) to determine spike display level.
Validation of spike expression was performed using S309, also known as Sotrovimab, a well-characterized anti-spike antibody that is cross-reactive to SARS-CoV-2 variants and is known to have reduced binding to Omicron variants and no binding to MERS spike protein (Pinto et al. [28]).
General notes and troubleshooting
General notes
Make sure to test the transfected cells with control antibodies to confirm that the spike is displayed. We recommend using S309. See Figure 3B for an example.
This protocol can be modified to test for antibody binding to various SARS-CoV-2 variant spike proteins by substituting the transfection plasmid DNA to include desired amino acid changes. The resulting antibody binding can be evaluated by a full dilution curve as above or by single antibody concentrations within the dose-response range to various spike variants. See Figure 5B for an example.
This protocol has been tested using other cell lines for spike expression, including 293F and ExpiCHO. While spike expression was detected, we also observed a higher percentage of dead cells than when using Expi293F. While dead cells can be gated out during analysis, they introduce potential sources of error, especially in cases of “sticky” antibodies.
Troubleshooting
Part I: Effective KD by flow cytometry
Problem 1: No antibody staining observed.
Possible cause A: Antibody has a weak affinity.
Solution A: Increase the highest concentration of antibody used.
Possible cause B: Secondary antibody is inactive or incompatible with the primary antibody used.
Solution B: Check the compatibility of the primary antibody and the secondary antibody used in an ELISA or similar assay.
Problem 2: High percentage of dead cells.
Possible cause A: Contamination in DNA preparation, transfection reagents, or media.
Solution A: Filter-sterilize all reagents.
Possible cause B: Transfecting too much DNA.
Solution B: Repeat using lower DNA mass (or re-measure DNA concentration).
Possible cause C: Spike expression is toxic for the cells.
Solution C: Repeat using lower DNA mass.
Problem 3: High staining found in the negative controls (isotype antibody and/or secondary antibody–only controls).
Possible cause A: Non-specific binding by the isotype control or secondary antibody.
Solution A: Isotype and secondary antibodies can exhibit low non-specific binding to one cell line but be problematic for another. Accordingly, we first screen secondary antibodies, including those from different vendors and polyclonal vs. monoclonal options, anti-human-Fc, anti-human kappa, anti-human H+L, and F(ab)2 preparation of anti-human Fc to identify one with low non-specific binding to cells, similar to unstained cells. We then screen several candidate isotype control antibodies to find one that exhibits minimal non-specific binding. Extra washes and or different antibody concentrations may also be required.
Possible cause B: High percentage of dead cells.
Solution B: Dead cells will often bind to antibodies indiscriminately. Staining of untransfected cells with high viability can help determine if this is the issue. Follow the suggestions in Problem 2 to reduce the fraction of dead cells in your samples.
Problem 4: Staining by the experimental antibody is low or high across all concentrations tested.
Possible cause: Antibody concentrations used are outside the dose-response range.
Solution: If the signal is low, increase concentration. If the signal is high, decrease concentration. Perform a pilot experiment using a wide range of concentrations with 5-fold or 10-fold dilution steps to define those corresponding to maximum and minimum signals. Repeat the experiment using smaller dilution steps across the confirmed dose-response range.
Part II: Antibody-dependent cellular cytotoxicity
Problem 1: Final ADCC values above 100%.
Possible cause: Incomplete lysis in maximum lysis control or inconsistent cell numbers per well.
Solution: Make a new lysis buffer or optimize co-culture incubation time for complete lysis. Avoid clumps in initial 293T passaging, and pipette gently but thoroughly before and during cell seeding into plates.
Problem 2: Final ADCC values below 0%.
Possible cause: High spontaneous lysis or inconsistent cell numbers per well.
Solution: Make sure cells have good viability before seeding. To avoid edge effects, do not use the outer wells of the plate or fill the space between wells with PBS. Avoid clumps in initial 293T passaging, and pipette vigorously before seeding in plates.
Problem 3: High error between replicates.
Possible cause A: Inconsistent cell number per well or inconsistent calcein-AM loading due to clumps in target cells.
Solution A: Avoid clumps in initial 293T passaging, and pipette vigorously before seeding in plates. Additional parameters that may need to be optimized include E:T ratio, total number of cells, incubation times, and antibody concentration.
Possible cause B: Issues with the multi-channel pipette.
Solution B: Visually inspect multi-channel during pipetting to determine whether equal volumes are being administered by all channels.
Acknowledgments
The following reagent was contributed by David Veesler for distribution through BEI Resources, NIAID, NIH: Vector pcDNA3.1(-) containing the SARS-Related Coronavirus 2, Wuhan-Hu-1 Spike Glycoprotein Gene, NR-52420 [35]. Flow cytometry and confocal microscopy were performed at the Center for Biomedical Research Support Microscopy and Imaging Facility at UT Austin (RRID# SCR_021756).
The graphical abstract was created in BioRender. Wilen, R. (2024) BioRender.com/l51i973.
This work was supported by the Bill & Melinda Gates Foundation INV-017592 (J.A.M.); Welch Foundation grant F-1767 (J.A.M.); NSF RAPID 2027066 (J.A.M.); and a Texas Biologics grant (J.A.M.).
Competing interests
A.W.N. and J.A.M. are inventors on U.S. patent application no. 63/135,913 (“Cross-reactive antibodies recognizing the coronavirus spike S2 domain”). J.A.M., is an inventor on patent no. WO/2021/243122 and PCT/US2021/034713 (“Engineered Coronavirus Spike (S) Protein and Methods of Use Thereof”). JAM is on the scientific advisory boards of Janux and Releviate. and previously received research funding from Synthetic Biologics and Dynavax. This work was supported in part by a grant from the University of Texas Biologics to J.A.M.
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Article Information
Publication history
Received: Apr 15, 2024
Accepted: Sep 28, 2024
Available online: Oct 30, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
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Immunology > Antibody analysis > Antibody-antigen interaction
Molecular Biology > Protein > Expression
Immunology > Antibody analysis > Antibody modification
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5,120 | https://bio-protocol.org/en/bpdetail?id=5120&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
A High-Throughput Droplet-based Method to Facilitate Microbial Conjugation
MC Monica J. Chu *
JW Jose A. Wippold *§
RR Rebecca Renberg
MH Margaret Hurley
BA Bryn L. Adams
AH Arum Han
(*contributed equally to this work, § Technical contact)
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5120 Views: 235
Reviewed by: Alba BlesaMercedes SanchezFernando A Gonzales-Zubiate
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Original Research Article:
The authors used this protocol in New Biotechnology Jul 2024
Abstract
Droplet microfluidic platforms have been broadly used to facilitate DNA transfer in mammalian and bacterial hosts via methods such as transformation, transfection, and conjugation, as introduced in our previous work. Herein, we recapitulate our method for conjugal DNA transfer between Bacillus subtilis strains in a droplet for increased conjugation efficiency and throughput of an otherwise laborious protocol. By co-incubating the donor and recipient strains in droplets, our method confines cells into close proximity allowing for increased cell-to-cell interactions. This methodology is advantageous in its potential to automate and accelerate the genetic modification of undomesticated organisms that may be difficult to cultivate. This device is also designed for modularity and can be integrated into a variety of experimental workflows in which fine-tuning of donor-to-recipient cell ratios, growth rates, and media substrate concentrations may be necessary.
Key features
• Builds on previous Bacillus-conjugation methods introduced by Brophy et al. [1] increasing the throughput by flowing the donors and recipients into a droplet microfluidic chip.
• Experiments performed on this chip increase conjugation efficiency as compared to conjugation performed traditionally by co-incubating the cells in free culture.
• This platform enables fine-tuning of experimental parameters, e.g., donor-to-recipient cell ratios, induction concentration, and incubation times, all critical factors in engineering undomesticated organisms.
• Adaptable for upstream automation of bacterial cultivation and downstream analysis of transconjugants encapsulated in droplets.
Keywords: Droplet microfluidics Bacterial conjugation Automation Synthetic biology Genetic engineering
Graphical overview
Background
Droplet microfluidic platforms have been broadly used to enable facile transformation of both mammalian and microbial cells in a high-throughput manner. This has been demonstrated through microfluidic electroporation of droplet-encapsulated cells [2,3], chemically-mediated transformation of cells (in some instances with heat shock added) [4,5], as well as transfection through means of mechanoporation [6], which has even been demonstrated in difficult-to-transfect primary T lymphocytes [7], moving toward CAR-T applications.
As evidenced, there exists a wide array of droplet-based microfluidic tools and techniques developed to streamline transformation and transfection, yet conjugative methods of DNA transfer have been scarcely explored. While conjugation is a modality of DNA transfer exclusive to microbial species, it is a powerful method commonly used to introduce heterologous DNA into microbial hosts that may be less amenable to modification. A canonical example of this is E. coli-based conjugation between F+ and F- cells. In this case, E. coli containing an F plasmid are able to synthesize a pilus structure allowing for DNA to be transferred during cell-to-cell contact [8]. Conversely, Agrobacterium-mediated DNA transfer, which leverages its native pathogenic functions to transfer DNA to a microbial or plant host, is able to do so through a Type IV secretion system [9]. Lastly, a unique example of conjugation is that of some strains of B. subtilis, which involves the transfer of integrative conjugative elements (ICE) from one host to the other. The XPORT strain [1], used in this protocol, is an engineered donor strain with a modified ICE. In the aforementioned work, this strain was used to transfer DNA via conjugation to a broad array of undomesticated strains of Bacillus and other related strains. Moreover, XPORT has demonstrated survivability and successful conjugation in soil environments where cell growth is uncontrolled. While the biological components of the system are robust, we have designed a microfluidic platform (previously introduced in Wippold et al. [10]) that offers more experimental control and complements the workflow described by Brophy et al. [1], expediting an otherwise laborious protocol and enabling easy optimization of experimental parameters such as donor-to-recipient ratios.
Although there have been other efforts demonstrating in-droplet conjugation, such as that introduced by Lam et al. [11], we propose two advantages to our protocol: (1) The lower surfactant concentration, which yields higher cell viability and enables longer incubation times if needed, and (2) the additional micro-vessel (on the secondary chip), which allows for accurate timing over co-incubation times with the first-in, first-out system. We also acknowledge several disadvantages to this device: (1) The current system does not allow for in-line sorting of successful transconjugants from unconjugated recipients, (2) there is no option for facile, on-chip incubation besides placing the entire device in an incubated chamber, and (3) the current tandem (two-chip) system introduces more potential points of failure, such as leakage between the fluidic tubing connections and the increased potential for air bubble introduction at the tubing-chip interfaces. For example, a downstream reagent inlet stream can be incorporated into the microfluidic platform that allows for the injection of a microdroplet breaking reagent (such as the commercial PicoBreak solution or 1H,1H,2H,2H-perfluoro-1-octanol PFO) to controllably release the inner contents of the droplets. Another possibility is to pair a commercial droplet sorting and plating system (such as the one marketed by On-Chip Biotechnologies or Atrandi Biosciences) to enable a mechanism to detect, sort, and place desired droplets into known areas.
Overall, we envision several areas of application for this device beyond the scope of what is presented in this work, which solely leverages B. subtilis to B. subtilis conjugation. This device can potentially be used with other donor organisms designed for conjugation (e.g., Agrobacterium tumefaciens AGL1) or perhaps even expanded to a tri-parental mating system, such as that described in Banta et al. [12] and Heinze et al. [13]. Moreover, with the addition of upstream and downstream modules, one may also automate the culturing steps leading up to the conjugation as well as the breakage of droplets and subsequent plating steps.
Materials and reagents
Biological materials
Donor strain JAB981 (from Brophy et al. [1]), B. subtilis JH642 Δ(ydcS-yddM)::[(Pspank-GFPmut2) tet(M) cat] thrC::[(int-yddJ) ΔnicK mls] alrA::[(Psweet-rapI) spc]
Recipient strain JAB545 (from Brophy et al. [1]), B. subtilis PY79 cured of ICEBs1, ΔcomC:: aphA-3
Reagents for bacterial culture
LB Miller (Fisher Scientific, catalog number: BP1426-2)
Agar (Fisher Scientific, catalog number: BP1423-500)
Phosphate buffered saline (PBS) (VWR, catalog number: 97062-336)
D-alanine (Sigma-Aldrich, catalog number: A7377)
Tetracycline hydrochloride (Calbiochem, catalog number: 583411; may alternatively use Sigma-Aldrich, catalog number: T7660)
Isopropyl β-d-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, catalog number: I5502)
NH4Cl (Sigma-Aldrich, catalog number: A9434)
MgCl2 (Sigma-Aldrich, catalog number: M8266)
K2HPO4·3H2O (Sigma-Aldrich, catalog number: P5504)
Materials for chip fabrication and use
SU-8 2075 photoresist (Kayaku, formerly Microchem Corp)
SU-8 2050 photoresist (Kayaku, formerly Microchem Corp)
Microposit EBR 10A (Kayaku, formerly Microchem Corp)
IP-S (Nanoscribe GmbH)
Tridecafluoro-1,1,2,2 tetrahydrooctyl trichlorosilane (United Chemical Technologies LLC, catalog number: T2492)
Poly(dimethylsiloxane) (PDMS) (Dow Corning Corp, PDMS Sylgard 184, catalog number: 761028-5EA)
3-(trimethoxysilyl)propyl methacrylate (Sigma-Aldrich, catalog number: 440159)
Propylene glycol monomethyl ether acetate (PGMEA) (Sigma-Aldrich, catalog number: 484431)
99% isopropyl alcohol (ULINE, catalog number: S20735)
EBR-10 A (Microposit Kayaku Advanced Materials)
3M Novec 7500 Engineered Fluid (3M, catalog number: 7100025016)
PicoSurf (Sphere Fluidics, catalog number: C021)
Reagents for confirmation assays
Q5 polymerase (NEB, catalog number: M0491L)
dNTPs (NEB, catalog number: N04475)
Q5 reaction buffer (NEB, catalog number: B9027S)
10 μM primers (IDT, exact sequences are noted in the protocol)
Sheath fluid for flow cytometry (Thermo Scientific, NERL Blood Bank Saline, catalog number: 8504)
ReadyLyze Lysozyme (LGC Biosearch Technologies, Lucigen, catalog number: R1804M)
MasterPure Total Nucleic Acid Isolation Kit (LGC Biosearch Technologies, Lucigen, catalog number: MC85200)
Solutions
Tris-Spizizen salts (TSS) buffer (see Recipes)
Carrier (oil) phase for ENTRAP device (see Recipes)
LB agar (see Recipes)
Recipes
Tris-Spizizen salts (TSS) buffer
Reagent Final concentration Quantity or Volume
NH4Cl 150 mM 2 g
K2HPO4·3H2O 20 mM 0.35 g
Tris base (pH 7.5) 49.5 mM 6 g
MgCl2 125 mM 11.9 g
H2O n/a 1,000 mL
Carrier (oil) phase for ENTRAP device
Reagent Final concentration Quantity or Volume
PicoSurf (5% in Novec 7500) 1.25% (v/v) 1 mL
Novec 7500 n/a 4 mL
Total n/a 5 mL
LB agar (250 mL)
Reagent Final concentration Quantity or Volume
LB 2.5% (w/v) 6.25 g
Agar 1.5% (w/v) 3.75 g
H2O n/a 250 mL
Laboratory supplies
15 mL culture tubes (Corning, Falcon, catalog number: 352059)
5 mL plastic syringe (BD, catalog number: 309646)
1 mL plastic syringe (BD, catalog number: 309628)
0.2 μm Whatman Puradisc 13 syringe filter (Sigma-Aldrich, catalog number: WHA67801302)
Microtube-130 AFA fiber for sonication (Covaris, catalog number: 520216)
Short Read Eliminator XS kit (Circulomics (PacBio))
Non-DEHP tubing, 30 gauge (Saint-Gobain Performance Plastics, Tygon ND-100-80, catalog number: VWR 89404-300)
Micropipettes 100–1,000 μL (VWR, Eppendorf, catalog number: 89125-306)
Pipette tips 1,000 μL (USA Scientific, TipOne, catalog number: 1122-1830)
96-well microplate (Corning, catalog number: 353072)
Glass slides, 75 × 50 (Corning, catalog number: 2947)
Equipment
Microcentrifuge (Eppendorf, model: Centrifuge 5420)
Shaking incubator (New Brunswick, Innova 42R, catalog number: M1335-0080)
Spectrophotometer (VWR, model: Biowave CO8000 Cell Density Meter)
Stationary incubator (Thermo Fisher Scientific, Heratherm, catalog number: 50125590)
Focused ultrasonicator for genome extractions (Covaris E220 Evolution, catalog number: 500429)
Desiccator (Zoro, catalog number: G4655463)
Spin coater (Laurell Technologies Corporation, model: H6-23)
Photolithography mask aligner (EVG Group, model: EVG 610)
Syringe pump (Chemyx, Fusion 200-X, catalog number: 0720X)
Plasma cleaner (Harrick Plasma, catalog number: PDC-32G)
Optical microscope (Zeiss, model: Colibri Axiovert 200)
Nanoscribe Photonics Professional GT (Nanoscribe GmbH)
Software and datasets
Geneious 2023 (paid software for genetic mapping)
Alternative genetic mapping software: Benchling (free online interface for gene editing)
AutoCAD (paid software for designing master molds)
Alternative CAD software for silicon mold design: Blender, FreeCAD
Filtlong v0.2.1 (https://github.com/rrwick/Filtlong.git)
Raven v1.7.0 (https://github.com/lbcb-sci/raven.git)
medaka v1.4.2 (https://github.com/nanoporetech/medaka.git)
prokka v1.14.5 (https://github.com/tseemann/prokka.git)
Procedure
Device fabrication
Option 1: Traditional photolithography method to fabricate master mold on silicon wafer
Spin coat SU-8 2050 onto a silicon wafer at 2,100 rpm. See General note 1.
Apply the first soft baking step at 65 °C for 24 h.
Apply the second soft baking step at 95 °C for 20 min.
Expose the coated Si wafer in the photolithography mask aligner using a mask design (either screen printed or using chromium mask processing) wafer using standard i-line (365 nm) and a total dose of 1,000 mJ/cm2 (Figure 1).
Figure 1. Mask designs used for the photolithography-based master mold process during the microfluidic device fabrication workflow. Left: DNA ENTRAP mask designs. Right: Microdroplet observation chamber device.
Apply a post-exposure baking step at 65 °C for 40 min, then 95 °C for 20 min.
Remove the unexposed photoresist using EBR-10A (commercial Edge Bead Remover solution engineered to eliminate any unexposed negative photoresist).
Using a conventional desiccator, apply a coating of tridecafluoro-1,1,2,2 tetrahydrooctyl trichlorosilane for 60 min. See General note 2.
For this, orient all wafers to be silanized feature-side up. Apply 2–4 drops of trichlorosilane (using a plastic pipettor; roughly 100 μL total) to a small weigh boat.
Option 2: Two-photon lithography method to fabricate master mold on silicon wafer.
For the devices used in this work, two approaches can be used for the fabrication of the microfluidic master molds that are used to conduct the soft lithography steps (i.e., the replica molding step to create the PDMS chips). We used a typical SU-8 photolithography approach along with a two-photon polymerization approach, depending on the overall geometry and footprint of the microfluidic chips that were designed. See General note 3.
Silanizing the silicon wafer
i. Make a bath of 1% solution of coupling agent [3-(trimethoxysilyl)propyl methacrylate] in ethanol.
ii. Submerge substrates overnight. Submerging can be expedited using only a 4 h period, if timing is an issue.
iii. Remove the substrate from the chemical bath, rinse with distilled water, and dry using a N2 gun.
iv. Store wafers in an airtight container at room temperature for up to 3 months. See General note 4.
Adding IP-S photoresist
i. Apply a volume of IP-S resist sufficient to cover the intended footprint of the microfluidic geometry to be fabricated.
ii. Gently spin the photoresist (500 pm) for 60 s to ensure an even spreading.
Running Nanoscribe
i. Run fabrication using Nanoscribe 2PP tool.
Upload the .STL file to the two-photon polymerization software tool: DeScribe.
Fabricate the master mold pattern on the Nanoscribe Photonics Professional GT (Nanoscribe GmbH, Germany) using the pretreated silicon wafer as the substrate and negative photoresist (IP-S, Nanoscribe GmbH, Germany) using a power scaling of 1.0, tetrahedron inner scaffold, base scan speeds of 50,000, base laser power of 60%, shell/scaffold scan speeds of 100,000, and shell/scaffold laser power intensities set at 70%. For more details regarding this method, see Wippold et al. [14].
Developing
i. Following the fabrication run, remove the wafers from the tool and develop in propylene glycol monomethyl ether acetate (PGMEA) for 6 min.
ii. For fine development, place the wafers containing the master mold in 99% isopropyl alcohol (IPA) for 10 min.
iii. Following the IPA bath, dry the masters gently with nitrogen gas and inspect under a microscope for quality assurance [14].
Post-fabrication silanization of microfabricated structures
Following the inspection, coat the patterned wafers with tridecafluoro-1,1,2,2 tetrahydrooctyl trichlorosilane for 20 min to prevent pattern removal during PDMS replication [14].
Generating the ENTRAP device using PDMS
Mix ~22 g of PDMS (2 g of curing agent and 20 g of polymer material; 1:10 ratio as specified by the manufacturer). See Figure 2.
Figure 2. Stepwise procedure from weighing PDMS pre-polymer to mixing the pre-polymer with the curing agent
Place the silicon wafer mold (fabricated in steps A1–2) into a Petri dish or any clean container large enough to fit the silicon wafer. Pour 20 g of material over the mold (Figure 3). See General note 5.
Figure 3. Stepwise procedure for pouring mixed PDMS onto a silicon master mold attached via Kapton tape to a plastic Petri dish
For ease of removal of the PDMS post-curing, secure the Si-SU-8 master mold to the Petri dish using Kapton tape along the edges of the mold. Place the Petri dish containing the PDMS-covered wafer into a desiccator and apply vacuum pressure to remove bubbles. Bubbles may also be popped with a pipette tip (Figure 4).
Figure 4. Stepwise procedure for de-gassing the mixed PDMS and removing all bubbles
Allow PDMS to cure over the mold either at room temperature (~25 °C) overnight on a level surface or at 65 °C for ~1–4 h in an oven (Figure 5).
Figure 5. Stepwise procedure for oven-curing a PDMS mold
Fabricating the bottom surface of the channels
Using the PDMS made in step A3, coat a clean glass slide with a thin layer of PDMS (30 μm thickness) by spin-coating at 3,000 rpm for 30 s.
Allow this PDMS to cure under the same conditions as in step A3c. See General note 6.
Bonding the ENTRAP device to the PDMS-coated glass slide and preparing for experimentation
Carefully, using a scalpel or razor, cut a large rectangle in the PDMS around the channel patterns (Figure 6).
Figure 6. Stepwise procedure for removing a PDMS microfluidic molded device from the silicon master mold wafer
Remove the PDMS rectangle (which contains the channels) from the silicon wafer and place it on the PDMS-coated glass slide. See General note 7.
Introduce tubing for the inlet and outlets of the device
Use a needle to poke holes in the device where the inlets and outlets are designed (Figure 7).
Figure 7. Stepwise procedure for preparing a PDMS microfluidic molded device for oxygen plasma bonding (final step)
Oxygen plasma bonding
Place the device in the plasma cleaner and apply a 90 s plasma treatment at a power of 150 W and a pressure of 150 mTorr (Figure 8).
Figure 8. Bonded microfluidic chip (two identical experimental patterns) following successful oxygen plasma bonding
Place Tygon tubing in the holes for each of the inlets (one for the oil inlet, one for the cells) and one for the outlet.
Connect the outlet tube to the second chip containing the micro-vessel system for incubation of the cells.
Cell culture preparation
Streak strains from glycerol stock onto LB agar plates with selection antibiotics. B. subtilis donor strain JAB981 should be struck onto LB agar supplemented with tetracycline (10 μg/mL) and D-alanine (100 μg/mL). B. subtilis recipient strain JAB545 should be struck onto LB agar; supplementation with chloramphenicol (10 μg/mL) is optional.
Incubate these streaked agar plates overnight (>14 h) at 37 °C. Pause point.
Ensure that at least 25 mL of LB is available for the entirety of the culturing and dilution process described ahead. Pick a single large colony from the plates and suspend this colony in 3 mL of LB liquid media supplemented with the same selection antibiotics that were on the agar media. See General note 8.
Culture the donor and recipient strain to an OD600 between 0.8 and 1.2, as read on the spectrophotometer.
At this point, subculture these strains into fresh media with the same selection antibiotics (culture volumes: 3–5 mL).
Allow the donor strain to grow to an OD600 of 0.2; at this point, xylose should be added to the culture to a final concentration of 1%.
Critical: This step serves to induce the conjugation machinery in the donor strain. No conjugation will occur without this.
Culture both strains to an OD600 between 0.8 and 1.0. See General note 9.
Pellet cultures by centrifugation at 3,000× g for 3 min.
Wash pellets with 1 mL of PBS and re-centrifuge at 3,000× g for 3 min. Repeat twice.
Resuspend both pellets in either TSS media (Recipe 1) or LB media supplemented with D-alanine (100 μg/mL) to a final OD600 of 0.6. See General note 10.
Conjugation of cells in the ENTRAP device
Prior to experimentation, the fully fabricated PDMS DNA ENTRAP microdevices should be baked at 80 °C for 4 h to ensure a state of ensured hydrophobicity.
Combine the donor cells with recipient cells at a 1:1 ratio. This step is optional, should you wish to keep the donor and recipient cells separate and use separate inlets for each. In such a case, use the chip designed with two reagent inlets. See General note 11.
Load at least 500 μL of the donor/recipient cell mix into a 1 mL BD plastic syringe.
Prepare the oil phase solution according to Recipe 2. Filter the solution through a 0.2 μm filter before use.
Prime all microfluidic channels and devices with the oil phase solution prior to operation.
Load 5 mL of the carrier oil/surfactant mixture (Novec 7500/PicoSurf) into a 5 mL BD plastic syringe.
Place the loaded syringes into separate Chemyx Fusion 200 pumps. For the syringe containing the cell suspension, set the syringe pump at a flow rate of 400 μL/h. For the syringe containing the carrier phase (oil + surfactant), set the syringe pump to a flow rate of 2,000 μL/h. See General note 12. Also, see Figure 9 for a deconstructed view of the tubing arrangement.
Figure 9. Tubing layout of ENTRAP device. Left image: DNA ENTRAP front-end microfluidic system with labeled reagent syringes. Right image: Close-up of the left image. Green tag: oil and surfactant phase. Orange tag: inlet for cell A. Pink tag: inlet for cell B. Red tag: inlet for inducer reagent. White tag: outlet.
Run both syringe pumps to begin droplet formation. Ensure that the outlet tube of the ENTRAP chip is connected to the second chip containing the micro-vessel system (Figure 10), which enables accurately timed incubation periods. Allow the droplets to accumulate in the micro-vessel system.
Figure 10. Microfluidic vessel for droplet collection. Top: One example of a microdroplet collection example. Bottom: Example of the microfabricated micro-vessel chamber (droplets filled with blue dye).
While droplets are being formed, you may perform imaging under an inverted optical microscope (Figure 11). For the Zeiss AxioObserver Colibri, use the following parameters:
For brightfield imaging, use the 10× objective with a phase contrast (Ph1) ring and 10 ms exposure time at 9 V total light power.
For fluorescent imaging, the corresponding LED (to excite the fluorophore or fluorescent protein) should be used in combination with an appropriate dichroic mirror/beam splitter and emission filter (to remove signal not corresponding to the excited target). For example, with GFP imaging, the BP 469/38 LED should be used to excite the fluorophore, while paired with a triple band pass splitter 405+493+610. LED power should be set to 95% and exposure times ranging from 300 to 3,000 ms. Select the appropriate exposure time that provides satisfactory signal-to-noise feedback. Keep the exposure time consistent across all experimental runs.
Figure 11. Experimental setup using blue dye–filled droplets for illustrative purposes
You may stop the syringe pumps when you have achieved the desired volume (or expected droplet number). Clip the end of the tube (stopping flow) by placing ~0.5 cm of the outlet tube into a microcentrifuge tube and closing the lid.
Incubate droplets for at least 1 h at ambient temperatures or 37 °C (depending on your recipient strain’s culture constraints). See Figure 12.
Figure 12. Microdroplet incubation vessel placed in a shake incubator
After the desired co-incubation period, flow the droplets out from the micro-vessel chip.
Pipette 50 μL of the droplets that are in oil suspension within the micro-vessel chambers used for conjugation onto a selection agar plate (Figure 13), which may be any media appropriate for transconjugant growth supplemented with 10 μg/mL tetracycline. Spread the droplets (breaking them) using a sterile L-spreader. For the recipient strain, JAB 545, we use LB Miller as the base media. See General note 13.
Figure 13. Plating of microdroplets on agar media for transconjugant selection. Top: Plating microdroplets from micro-vessel type 1. Bottom: Plating microdroplets from micro-vessel type 2.
For controls and conjugation efficiency calculations, plate the same volume of droplets onto (1) selection agar containing 100 μg/mL D-alanine and 10 μg/mL tetracycline, which will capture both donor and transconjugant, and (2) agar media with no antibiotic or D-alanine supplementation, which will capture untransformed recipients and transconjugants while preventing JAB981 donors from growing due to their auxotrophy.
Incubate the plates (~16 h) in a 37 °C stationary incubator or at the preferred temperature of the recipient strain. Most of the Bacillus recipient strains we have used the XPORT donor with can tolerate 37 °C, but one should take heed to the culture conditions of the recipient when plating for transconjugants. Pause point.
Transconjugant selection and evaluation
After the incubation period, inspect the transconjugant plates (plates with 10 μg/mL tetracycline) for colonies. If colonies have not yet grown, you may need to incubate for longer.
Pick several colonies from the transconjugant plate. Inoculate 3 mL cultures, supplemented with 10 μg/mL tetracycline. After reaching the mid-log phase, sub-culture these strains for improved consistency (across replicates) in expression.
When grown to an OD600 of ~0.2, induce cells with 1 mM IPTG, then shake for 1 h at 37 °C (or whatever temperature suits your recipient strain). Be sure to aliquot a portion of the cultures out as a negative (untreated) control. See General note 14.
After this incubation period, you may choose to perform various analyses to confirm the successful conjugation of DNA into the recipient strain.
Option 1: Flow cytometry. For this type of analysis, prepare a sample by diluting the induced sample and uninduced control in sheath fluid (100-fold dilution). Run the samples using the Sony SA3800 Spectral Analyzer with the following parameters: Threshold FSC value, 0.5%; FSC gain, 17; SSC voltage, 20%; fluorescence PMT voltage, 69.8%; acquisition of 10,000 events.
Option 2: Plate reader for fluorescence measurement. For this type of analysis, prepare the following dilutions of your samples: undiluted, 2-, 4- and 8-fold dilutions. Pipette 200 µL of each sample into a 96-well plate. Read absorbance at 600 nm and fluorescence in accordance with the excitation/emission parameters of the expressed fluorophore. In this work, we measure GFPmut2, which has an excitation/emission of 485 and 508 nm, respectively. Divide the relative fluorescence units (RFUs) by the absorbance at 600 nm of the same sample in order to normalize the difference in cell densities across samples.
Option 3: Fluorescence microscopy. One may use the sample parameters as stated above in step C11 of the above protocol.
Option 4: Colony PCR. Use the following primer sequences, flanking the insertion site, to confirm conjugation success: 5'-AAAGTCTTTTATTCTGCGCCG-3' (forward primer) and 5'-ATCATTTAGTAAGGCAGCTGCTA-3' (reverse primer). After obtaining the template DNA by boiling colonies for 3 min, set up the PCR reactions using Q5 polymerase and the aforementioned primers. Apply an annealing temperature of 64 °C and an extension time of 8.5 min. Sequence the resulting amplicon.
Option 5: Whole-genome sequencing. Obtain a cell pellet of the transconjugant. Isolate genomic DNA from the pellet using ReadyLyze lysozyme and MasterPure Total Nucleic Acid Isolation Kit on the Covaris E220 Evolution with a microtube-130 AFA Fiber. After isolating gDNA, treat with the Short Read Eliminator XS kit (Circulomics (PacBio)) to remove fragments smaller than 10 kb. Because the ICE insertion is larger than 10 kb, this procedure helps to ensure that DNA fragments corresponding to the ICE will contain at least one or both of the flanking sequences (upstream/downstream) of the integration site. Apply the Oxford Nanopore Ligation protocol (SQK-LSK109) to pre-treat the gDNA, then sequence the fragments using the GridION (Oxford Nanopore Technologies) on an R9.4.1 MinION flow cell.
Data analysis
To calculate conjugation efficiency, count the number of colonies on the tetracycline/D-alanine plate (i.e., number of donors + transconjugants) and then count the number of colonies on the tetracycline plate (i.e., number of transconjugants). Subtract the latter from the former; this number estimates the total number of donor CFUs. Divide the number of transconjugants by the number of donors to obtain the conjugation efficiency. Be sure to use at least three biological replicates for this calculation.
Sequences may be analyzed with the following protocol:
Filtering with filtlong (https://github.com/rrwick/Filtlong.git) to a minimum length of 1000
Assembly with raven (https://github.com/lbcb-sci/raven.git)
Polishing with medaka (https://github.com/nanoporetech/medaka.git)
Annotation with prokka (https://github.com/tseemann/prokka.git)
Resulting annotations can then be assessed with Geneious. For further sequencing confirmation of transconjugants, whole-genome sequencing on the transconjugants should be performed with the fragments de novo assembled into a genome. The de novo–assembled transconjugant genome can then be aligned with the original genome (wild type). Gaps where the transconjugant sequence does not match the wild-type genome should reveal any potential site where the ICE could have been inserted. Alternatively, the long-read fragments can be mapped to the theoretical sequence of the transconjugant genome to confirm the presence of the ICE.
For flow cytometry analysis, be sure to run at least three biological replicates and strive to induce all samples at the same point in their growth phase; this ensures better consistency in expression across samples. For the data acquisition, be sure to run at least 10,000 events. If there is enough sample volume and/or unusually heterogeneous population, an acquisition of 50,000 events is appropriate. Calculate the mean fluorescence of each sample and the standard deviation across three biological replicates. For multiple sample groups, one may apply Tukey’s multiple comparisons (post-hoc) test to determine whether or not there is any significant difference in expression between the controls and the experimental samples. See General note 15.
Validation of protocol
Validation of this original experimental work and data was previously presented in Wippold et al. [10].
General notes and troubleshooting
General notes
SU-8 2050 is used to obtain 65 μm channels (height). However, for 100 μm channels, use SU-8 2075 following manufacturer guidelines.
It is important to note that the layout of materials is consequential within the desiccator. The wafers to be silanized should be placed between the port connected to a vacuum source and the weigh boat with the trichlorosilane. This will enable a more thorough silanization of the wafers. If desired, rotate the wafer 180º after 30 min within the desiccators to ensure an even coating.
Both traditional photolithography and two-photon photolithography were used during the timeline of experimentation for this work. This was the result of the sponsoring laboratory’s cleanroom installing an upgraded system. Traditional photolithography is quicker, yet typically constrained to a 2.5D fabrication. The height of a resultant structure is typically set at a single level and achieved through spin-coat a wafer to a desired thickness (i.e., channel height). For two-photon photolithography (2PP), the channel height is fully dimensioned into the CAD drawing of the microchannel system. Thus, through 2PP, you are able to achieve multiple heights without the need for multiple spin-expose-develop-spin steps as required through a traditional approach. The drawback of the 2PP approach when compared to a traditional approach is the overall fabrication time needed to create the master mold. In a traditional approach, exposure of the entire device usually takes <5 min. In 2PP, the overall footprint of a microfluidic device master mold can take anywhere between 6 and 36 h depending on the total volume needing to be polymerized. For this experiment, a 2PP approach was used for devices containing complex geometry (i.e., sub-10 micrometer feature sizes or multiple height critical geometries).
Wafers are stable for up to three months. After three months of shelf storage, the authors cannot ascertain whether the surface modification will be strong enough to ensure full attachment of the photoresist.
Be careful not to introduce any bubbles at this step. If there are any bubbles left over from the previous step, you may centrifuge the PDMS mixture for 30 s at 2,000× g to remove them.
It should be noted that this step is optional. PDMS can readily bond to an uncoated glass slide.
When placing the PDMS down on the glass slide, be sure that the PDMS is flush against the slide with no bubbles. Check for cracks and any channel defects in the PDMS under the microscope.
Alternatively, instead of pulling a colony from a plate, liquid cultures (>3 mL) may be started from the previous night (shaken at 22 °C) and sub-cultured the next morning by performing no less than a 250-fold dilution of the overnight culture into fresh media. If you choose this alternative step, you may skip steps B4 and 5 and use this culture as the final culture to be used in the ENTRAP chip.
There is some flexibility in this portion of the protocol. While the OD600 range is somewhat arbitrary, this fixed range is proposed to ensure that the cells are being collected before the late exponential growth phase, where B. subtilis cells may begin to enter a vegetative state or begin to sporulate. Every spectrometer is different, so a growth curve analysis on your preferred instrument is recommended.
This resuspension media is the media that will be used for co-incubation. Alternative media may be used in this step if the donor and recipients used in this protocol differ from JAB981 and JAB545. For example, if the donor being used in this platform is an E. coli S17 and the recipient is another E. coli or Pseudomonas spp., then any compatible media to sustain both strains (e.g., LB or M9) may be used.
Pre-mixing cells can add bias to the results as one cannot ascertain whether conjugation occurs in the culture tube prior to microfluidic injection or in the droplet system. Therefore, it is best practice to avoid any mixing of cell types until the droplet generation point. Additionally, pre-mixing cells will result in an in-droplet distribution that will require additional optimization steps to ensure target in-droplet cell concentrations and ratios are favorable for the conducted experiments.
For dual reagent (donor and recipient separate) experiments, each injection input syringe is controlled via a distinct syringe pump operated at 200 μL/h. Variable flow rates can be applied for different donor:recipient ratios. In general, as dictated in part by droplet geometry, a 1:5–1:10 ratio between the input phase(s) and the carrier phase is optimal for generating droplets at a stable diameter.
PicoBreak may be used to disperse the droplets, but it is usually not necessary as the carrier phase is volatile and will volatilize once plated onto agar.
Inducing at OD600 ~0.2 is an arbitrary value. When developing this protocol, we aimed to induce the cells at the early log phase, which was OD600 = 0.2. However, the operator should consider the growth behavior of their particular strain. Inducing any time during the early log phase is recommended.
Given that [10] is the first XPORT conjugation-based droplet microfluidic work, there is little known regarding the expected confidence and p-values. However, as a potential frame of reference for the statistical analysis of conjugation results, please refer to Brophy et al. [1].
Troubleshooting
What to do if there are bubbles in the channel or if the droplets are merging?
Perform a surface treatment of the channels in the device. Prior to experimentation, flow aquapel through the DNA ENTRAP droplet generator device in reverse flow (from the out to the reagent inlets). Apply a short (~5 mm) tubing into each of the inlets. After initial filling, sequentially clamp reagent inlets while leaving one open. This will ensure aquapel flow is directed fully through the inlet channel. Repeat this process for all the inlet by releasing the clamp and replacing a clamp on the previously open inlet. In this case, 500 μL Eppendorf tubes were used as clamps. Allow aquapel to react with the PDMS/glass for 60 s. Remove the syringe from the outlet and air purge with an air-filled, hand-actuated 10 mL plastic BD syringe. Repeat the aquapel filling process another time to ensure full coating of the channel surfaces to enable a maximum hydrophobic treatment. Following this second aquapel treatment, repeat the air purge above. Next, replace the aquapel syringe with a 5 mL syringe with the carrier oil (0.2 μm filtered Novec 7500). Using the carrier oil, hand actuate the syringe to purge the channel of any residual aquapel from the channels with 500 mL of solution, again repeating the inlet clamp/release method to ensure all inlets are cleared. Finally, repeat the air purge to empty the channels. Inspect the microfluidic device under a microscope using a 10× objective to complete a quality control step to identify any particles that have infiltrated the channels. If any particles are seen, repeat the carrier oil purge to dislodge and remove the particle. At this point, one can engage flow from any of the available inlet ports or again through the outlet port. If no particles are seen, apply Kapton tape to the inlet and outlet ports to protect the microfluidic device from airborne particles. Set aside for later use. The aquapel-treated device will be stable at room temperature for one year. Immediately prior to experimental use, pre-fill all channels with the carrier oil. Once the channels are filled, begin engaging the reagent syringe pumps to inject the cell, reagent, and carrier oil + surfactant phases into the microfluidic device.
Acknowledgments
This project was supported in part by the United States Combat Capabilities Development Command - Army Research Laboratory (DEVCOM ARL) cooperative agreement with Texas A&M Engineering Experiment Station (TEES) (W911NF-17-2-0144). J. Wippold was supported through the Department of Defense Graduate SMART Scholarship program. The authors would like to thank the TEES AggieFab Nanofabrication Facility, the U.S. ARL SEMASC micro- and nanofabrication facilities for the microfabrication support, and the DoD High Performance Computing Modernization Program for computer resources. Validation of this experimental work and data was presented by Wippold et al. [10].
Competing interests
Authors of this work have applied for patent protection of the developed technology.
References
Brophy, J. A. N., Triassi, A. J., Adams, B. L., Renberg, R. L., Stratis-Cullum, D. N., Grossman, A. D. and Voigt, C. A. (2018). Engineered integrative and conjugative elements for efficient and inducible DNA transfer to undomesticated bacteria. Nat Microbiol. 3(9): 1043–1053.
Zhan, Y., Wang, J., Bao, N. and Lu, C. (2009). Electroporation of Cells in Microfluidic Droplets. Anal Chem. 81(5): 2027–2031.
Im, D. J. and Jeong, S. N. (2017). Transfection of Jurkat T cells by droplet electroporation. Biochem Eng J. 122: 133–140.
Pérez-Sosa, C., Sanluis-Verdes, A., Waisman, A., Lombardi, A., Rosero, G., Greca, A. L., Bhansali, S., Bourguignon, N., Luzzani, C., Pérez, M. S., et al. (2022). Single cell transfection of human-induced pluripotent stem cells using a droplet-based microfluidic system. R Soc Open Sci. 9(1): e211510.
Sha, J., Wang, Y., Wang, J., Liu, W., Tu, Q., Liu, A., Wang, L. and Wang, J. (2011). Heat-shock transformation of Escherichia coli in nanolitre droplets formed in a capillary-composited microfluidic device. Anal Methods. 3(9): 1988.
Joo, B., Hur, J., Kim, G. B., Yun, S. G. and Chung, A. J. (2021). Highly Efficient Transfection of Human Primary T Lymphocytes Using Droplet-Enabled Mechanoporation. ACS Nano. 15(8): 12888–12898.
Hur, J., Kim, H., Kim, U., Kim, G. B., Kim, J., Joo, B., Cho, D., Lee, D. S. and Chung, A. J. (2023). Genetically Stable and Scalable Nanoengineering of Human Primary T Cells via Cell Mechanoporation. Nano Lett. 23(16): 7341–7349.
Ippin-Ihler, K. (1986). The Conjugation System of F, the Fertility Factor of Escherichia coli. Annu Rev Genet. 20(1): 593–624.
Christie, P. J. and Vogel, J. P. (2000). Bacterial type IV secretion: conjugation systems adapted to deliver effector molecules to host cells. Trends Microbiol. 8(8): 354–360.
Wippold, J. A., Chu, M., Renberg, R., Li, Y., Adams, B. and Han, A. (2024). XPORT ENTRAP: A droplet microfluidic platform for enhanced DNA transfer between microbial species. New Biotechnol. 81: 10–19.
Lam, T., Maienschein-Cline, M., Eddington, D. T. and Morrison, D. A. (2019). Multiplex gene transfer by genetic transformation between isolated S. pneumoniae cells confined in microfluidic droplets. Integr Biol. 11(12): 415–424.
Banta, A. B., Ward, R. D., Tran, J. S., Bacon, E. E. and Peters, J. M. (2020). Programmable Gene Knockdown in Diverse Bacteria Using Mobile‐CRISPRi. Curr Protoc Microbiol. 59(1): e130.
Heinze, S., Kornberger, P., Grätz, C., Schwarz, W. H., Zverlov, V. V. and Liebl, W. (2018). Transmating: conjugative transfer of a new broad host range expression vector to various Bacillus species using a single protocol. BMC Microbiol. 18(1): 1–10.
Wippold, J. A., Huang, C., Stratis-Cullum, D. and Han, A. (2020). Enhancing droplet transition capabilities using sloped microfluidic channel geometry for stable droplet operation. Biomed Microdevices. 22(1): 1–5.
Article Information
Publication history
Received: Jul 6, 2024
Accepted: Sep 8, 2024
Available online: Oct 16, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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A Simple, Rapid, and Cost-Effective Method for Assessing Carbohydrate Partitioning in Microalgae and Arabidopsis thaliana
AB Araceli N. Bader §
LR Lara Sánchez Rizza
MM María A. De Marco
AL Ana P. Lando
GM Giselle M. A. Martínez-Noël
VC Verónica F. Consolo
LC Leonardo Curatti
(§ Technical contact)
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5121 Views: 273
Reviewed by: Abhilash PadavannilWalmik Karbhari GaikwadJoyce Chiu
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Abstract
Carbohydrates serve crucial functions in most living cells, encompassing structural and metabolic roles. Within the realms of plant and algal biology, carbohydrate biosynthesis and partitioning play pivotal roles in growth, development, stress physiology, and various practical applications. These applications span diverse fields, including the food and feed industry, bioenergetics (biofuels), and environmental management. However, existing methods for carbohydrate determination tend to be costly and time-intensive. In response to that, we propose a novel approach to assess carbohydrate partitioning from small samples. This method leverages the differential solubility of various fractions, including soluble sugars, starch, and structural polymers (such as cellulose). After fractionation, a straightforward spectrophotometric analysis allows for the quantification of sugars.
Key features
• We developed a cost-effective method to assess carbohydrate distribution in small samples based on differential solubility and spectrophotometry.
• Efficient carbohydrate partitioning methods reduce time and effort, especially for large sample sets.
• Cost-effective carbohydrate analysis method reduces expenses, promotes accessibility, and encourages adoption in research and quality control.
• Analysis of small samples from microalgal and A. thaliana seedlings has wide applicability in scientific and technological research.
Keywords: Sugars Starch Structural carbohydrates Arabidopsis thaliana Microalgae Carbohydrate partitioning method
Graphical overview
Description of the complete process. A total of 10 mg of dry sample or an equivalent quantity of fresh biomass is sufficient to carry out the extraction process. The initial step involves disruption of the samples through freezing and thawing. This is followed by the sequential extraction of simple sugars through an alcoholic extraction, which is then subjected to centrifugation to retain the supernatant. The resulting pellet is retained for starch extraction/hydrolysis by cold perchloric acid. The resulting insoluble polysaccharides from this fraction are extracted/hydrolyzed by hot sulfuric acid. Finally, sugars are determined by the anthrone method in every fraction collected, which corresponds to sugars, starch, and structural carbohydrates. The basic principles and methods for sugar extraction with ethanol, starch extraction/hydrolysis with perchloric acid, and polysaccharides hydrolysis with sulfuric acid have been described before [1,2].
Background
Carbohydrates play essential roles in most living cells, serving structural and metabolic functions. In algae and plants, soluble sugars are crucial for various metabolic processes, including osmotic regulation, energy translocation, and signal transduction. When carbon assimilation exceeds the demand for growth and maintenance, the excess is converted into starch. Moreover, starch accumulates within cells during unfavorable environmental conditions, allowing algae and plants to withstand severe challenges. Polysaccharides, in conjunction with proteins, are vital components of cell walls, influencing their conformation, resistance, and stability. While microalgae cell walls exhibit extensive diversity, they may contain cellulose, pectin, agar, or alginate [3]. Conversely, in the model plant A. thaliana, the primary cell wall primarily consists of pectin, hemicellulose, and cellulose [4].
Several techniques are currently available for the quantitative identification and determination of carbohydrates, namely chromatography methods (such as HPLC and GC–MS), enzymatic assays, colorimetric methods, and physical measurements based on a specific gravity or refractive index [5]. While these methods are highly accurate, they often require expensive equipment and high-quality reagents and can be time-consuming. Additionally, methods for determining multiple carbohydrates from the same sample remain scarce.
To address this gap, we have developed an easy, rapid, and cost-effective method for assessing carbohydrate partitioning in microalgae and plants. This approach allows high-throughput data acquisition. A modified protocol for smaller sample sizes and reduced reaction volumes is provided in the Supplementary information.
Materials and reagents
Biological materials
Chlamydomonas reinhardtii represents a valuable research organism with a sequenced genome, simple life cycle, and ability to grow in multiple metabolic states. It is a model organism for studies in areas like flagella structure, chloroplast biogenesis, photosynthesis, and cell–cell recognition. Its cell wall is composed of glycoproteins and polysaccharides in a multi-layered structure. Strain CC125 represents a popular wild-type (WT) alga for the species; strain cw15 [6] is a cell wall–deficient mutant that facilitates genetic manipulation and cellular process studies; and sta6 is a mutant strain deficient in starch accumulation, which is very useful for studies on carbon partitioning and lipid accumulation [7].
Scenedesmus obliquus and Chlorella sorokiniana are thoroughly used for biotechnological applications such as the production of biofuels and food/feed ingredients and wastewater treatment. They show very robust and prolific growth in cultivation facilities. In contrast to C. reinhardtii, they currently offer limited possibilities for genetic modification and reduced availability of mutant strains. While the S. obliquus cell wall comprises a resistant outer layer of algaenan and an inner layer of cellulosic material, the C. sorokiniana cell wall presents a thin trilaminate layer of hemicellulose and a glucosamine-based residue and produces mucilage [8].
A. thaliana, a model organism in plant biology, boasts features like a small genome, rapid life cycle, and easy genetic modification. Its cell wall, composed of cellulose, hemicellulose, and pectin, provides strength, flexibility, and porosity. Some cells even develop a secondary cell wall with lignin for rigidity. Structural proteins, enzymes, and glycoproteins contribute to cell signaling and growth. Remodeling occurs during growth and in response to environmental stress [9].
All of these organisms produce starch as a carbon and energy reserve, and sugars with different physiological roles.
Organisms' cultivation conditions
The microalgal strains C. reinhardtii CC125 (WT), cw15 (cell wall–deficient), and sta6 (starchless mutant) were kindly provided by C. Benning (Michigan State University), and strains C. sorokiniana RP and S. obliquus C1S were isolated [10] and characterized [11] by our research group. These algal strains belong to the INBIOTEC culture collection.
The microalgal strains were cultivated and induced for carbohydrate accumulation as previously described in Bader et al. [12] under nitrogen-deficient conditions.
The A. thaliana used in this study was a wild type of Columbia (Col-O) stored in our laboratory. Seeds were sown in pots and grown under 16/8 h light/dark (light 120 μmol photons m-2·s-1) at 23 °C in a growth room. Fourteen-day-old A. thaliana plants were treated with 150 mM NaCl for 14 days before leaves were collected for carbohydrate analysis.
Reagents
Distilled water (dH2O)
Double-distilled water (ddH2O) for anthrone solution
Ethanol 96% (C2H6O) (e.g., Alco Protect, Porta, Argentina)
Perchloric acid 60% (HClO4) (Fluka, catalog number: 77232)
Sulfuric acid 96%–98% (H2SO4) (Biopack, Brand, catalog number: 0303A0036)
Glucose analytical grade (C6H12O6) (Glc) (Sigma-Aldrich, catalog number: G5767)
Anthrone (C14H10O) [Fluka, catalog number: A 866 (0740)]
Thiourea (CH4N2S) (Sigma-Aldrich, catalog number: T7875-500G)
Note: Both sulfuric acid and perchloric acid are hazardous to touch and inhalation; therefore, it is necessary to work with caution and use personal protective equipment (gloves, lab coat, closed-toe shoes, and safety goggles).
Solutions
For carbohydrate quantification, the anthrone method was used, which has been widely validated and allows for the quantification of a broad range of carbohydrates, hexoses, and aldopentoses [13]. In the presence of concentrated sulfuric acid, carbohydrates are dehydrated into furfurals (or hydroxymethylfurfurals), which condense with anthrone (10-keto-9,10-dihydroxyanthracene) (Figure 1) to produce a blue-green complex. The intensity of the color is quantified by absorbance at 620 nm.
Figure 1. Reaction of the anthrone assay. Modified from Gerwig [14]. The extinction coefficient of the complex between 5-hydroxy methyl furfural and anthrone at 620 nm is 76,000 M-1·cm-1.
67% sulfuric acid solution (see Recipes)
Anthrone solution (72% sulfuric acid final concentration) (see Recipes)
10 mM glucose solution (see Recipes)
Recipes
67% sulfuric acid solution (10 mL)
Conduct the process under a fume hood and using personal protective equipment (gloves, lab coat, closed-toe shoes, and safety goggles). In a glass container, pour 3.3 mL of ddH2O. Place the container on ice and then slowly add 6.7 mL of sulfuric acid to prevent acid splashes upon contact with water.
Anthrone solution (100 mL)
Conduct the process under a fume hood and using personal protective equipment (gloves, lab coat, closed-toe shoes, and safety goggles). In a glass container, pour 28 mL of ddH2O. Place the container on ice and then slowly add 35 mL of sulfuric acid to prevent acid splashes upon contact with water, while stirring with a magnetic stir bar. Add 50 mg of anthrone and 1 g of thiourea. Complete with 37 mL of sulfuric acid. Leave to stir for 16 h in darkness.
10 mM Glc solution (10 mL)
Prepare by adding 18 mg of Glc to 10 mL of ddH2O.
Equipment
General lab supplies: microtubes, pipettes, tips, tube racks, etc.
Block heater (Thermolyne, model: Type 17600 Dri-Bath)
Bench-top vortex (Labnet, model: VX100)
Benchtop centrifuge for 2 mL tubes at room temperature and 4 °C (Thermo Scientific, model: Sorvall Legend Micro 21R)
Fume Hood (Esco, model: EFA-4UDRVW-8)
5 mL glass test tubes
Analytical balance (Sartorius, model: Entris2241-1S)
UV/VIS spectrometer (Shimadzu Corp., serial number A114548, 05789)
1 cm Pathlength quartz cuvettes; disposable or glass cuvettes can also be used
Gas mask (3M, model: 6899B, filters 3M 6003) for measuring samples with anthrone
Thermostatic water bath for laboratory (Daglef Patz, model: Hor, 18.3867)
Magnetic stirring bar and magnetic stirrer (Thermolyne, model: Nuova II)
Procedure
Fractionated carbohydrate extraction
Sample preparation
Use approximately 5–10 mg of dry and ground sample or equivalent of fresh sample. To calculate the amount of fresh biomass necessary to carry out the entire process, the pellet obtained from 1.5 mL of culture was placed in a pre-weighed microtube and centrifuged at 6,200× g for 10 min. After centrifugation, the supernatant was discarded, and the tube was weighed to calculate the fresh weight. Then, the pellet was dried at 60 °C and weighed until a constant weight was reached. The quantification was performed in triplicate, and the average was used to calculate the dry weight to fresh weight ratio and to determine the amount of fresh biomass equivalent to 10 mg of dry biomass. For a smaller sample size, reduce proportionally the volumes of extraction solvents; for example, if 1 mL of extraction volume is used for 10 mg of biomass, 200 μL would be used for 2 mg of biomass.
Resuspend the sample in 250 μL of dH2O.
For microalgae, freeze and thaw the sample three times to break the cells. For plant samples, cell disruption is performed by maceration or by grinding with nitrogen in the case of fresh biomass.
Notes:
Cell disruption by freeze and thaw: Once water is added to the biomass, place the suspension in the microtubes in a freezer at -20 °C for a few minutes until frozen. Then, thaw at room temperature, vortex, and repeat the process two more times.
The material can be macerated directly in the microtube using a Teflon tip.
Extraction of sugars
Add 250 μL of 96% EtOH preheated in a water bath at 50 °C to extract sugars. Vortex for 30 s.
Centrifuge at 2,400× g for 10 min and keep the supernatant for sugar quantitation. Repeat step 1 three times (final volume of 1 mL).
Extraction of starch
Dry the pellet in a thermal block at 50 °C for 15 min to evaporate the EtOH.
Add 250 μL of dH2O, homogenize, and then add 250 μL of 60% HClO4. Vortex for 30 s.
Incubate on ice for 20 min to extract starch.
Centrifuge at 2,400× g for 10 min and keep the supernatant for starch quantification.
Add 250 μL of 60% HClO4, centrifuge (same as step 4), and keep the supernatant for starch quantitation. Repeat one more time (final extraction volume 1 mL).
Extraction of structural carbohydrates
Add 1 mL of dH2O to the pellet to rinse the HClO4. Vortex and centrifuge at 6,200× g for 10 min.
Add 500 μL of 67% H2SO4. Vortex for 30 s and let stand for 1 h. Vortex periodically three or four times during this period (since it is a thick suspension, it is very important to homogenize the sample well; otherwise, significant variation between replicates in the determination is observed).
Centrifuge at 16,200× g for 10 min and keep the supernatant for structural carbohydrates quantitation.
Note: Samples can be stored at -20 °C until quantitation.
Carbohydrate quantitation
Prepare a 1:10 dilution of the extracts (it is necessary to dilute the samples at least 1:5 to avoid interference of HClO4 with anthrone).
Take 200 μL of dilute extract and add 1 mL of anthrone reagent in a 10 mL glass tube. Vortex.
Boil for 15 min in a fume hood and cool to room temperature. Vortex.
Carefully, pour all the contents of the tube into a 1.5 mL cuvette.
Measure the absorbance at 620 nm.
Dispose of the sample in a container (do not pour it down the drain, as it must be discarded as hazardous waste), rinse the cuvette with distilled water, and clean the cuvette by removing the residual water through capillary action using vacuum before reading the next sample.
To prepare the standard curve, prepare samples containing from 50 to 400 nM Glc by adding 0, 5, 10, 20, and 40 μL of 10 mM Glc, and completing with dH2O to a final volume of 200 μL.
Note: Values for the total carbohydrates, starch, and structural carbohydrates fractions are expressed as polysaccharides, using a factor of 0.9 to convert the measured glucose value to polysaccharide [15].
Carbohydrates determination:
Calculate slope (α) and intercept (β) coefficients of the linear regression model (y = αx + β) from the calibration data as seen in Figure 2, and use these coefficients to calculate carbohydrate concentration in every sample and replicate according to the equation, where the y-axis represents the observed absorbance and the x-axis represents nanomoles of Glc.
Figure 2. Representative calibration of D-glucose (dissolved in distilled water) mixed with anthrone reagent, measured at 620 nm according to this protocol. Each calibration point represents the average of four technical replicates; the error bars represent standard deviations. The line represents the linear fit of the measured points calculated by the least squares method. The linear regression equation (y = αx + β) represents calibration equation slope (α) and intercept (β). The R2 coefficient was calculated by the least squares method.
Table 1 provides a generic example for performing calculations of the obtained carbohydrate quantities and extraction efficiency.
Table 1. Generic example for performing calculations of the obtained carbohydrate quantities and extraction efficiency of three methodological replicates
Sample Sample dilution A620 nm Mean abs 620 nm nmoles Glc μg Glc μg/μL Glc mg/mL * sample dilution *0.9(only in polysaccharides) Total substrate in the sample Carbohydrate percentage
Replicate 1 10 (for example) y1 y1´ A6201= (y1+y1´)/2 z1= (A6201- β)/α X1=((z1*180)/1000) Value1=X1/200 r1=Value1*10 R1=r1*0.9
10 (for example)
%1=((R1*100)/10)
Replicate 2 y2 y2´ A6202= (y2+y2´)/2 z2= (A6202- β)/α X2=((z2*180)/1000) Value2=X2/200 r2=Value2*10 R2=r2*0.9 %2=((R2*100)/10)
Replicate 3 y3 y3´ A6203= (y3+y3´)/2 z3= (A6203- β)/α X3=((z3*180)/1000) Value3=X3/200 r3=Value3*10 R3=r3*0.9 %3=((R3*100)/10)
Validation of protocol
This protocol, or parts of it, has been used and validated in the following research articles:
Bader et al. [12]; Caló et al. [16]; and De Marco et al. [17].
Protocol validation on purified carbohydrates
The protocol was validated on purified substrates by conducting the complete process on 10 mg of sucrose, starch, or cellulose. Additionally, the complete process was carried out on two samples with different proportions of purified carbohydrates. One sample contained 10 mg of sucrose, 10 mg of starch, and 10 mg of cellulose, while the other contained 3, 5, and 2 mg, respectively. The latter ratio was used to approximate the values of each carbohydrate in biomass. For this validation, sucrose, starch, and cellulose were used to represent the fractions of simple sugars, starch, and structural carbohydrates, respectively. Subsequently, the carbohydrates present in each fraction were quantified using the anthrone method (Figure 3).
Figure 3. Validation of protocol on purified carbohydrates. 10 mg of sucrose, starch, cellulose, or 1:1:1 (total = 30 mg) or 3:5:2 (total = 10 mg) mixtures were subjected to the sequential fractionation of carbohydrates. Each fraction (1 mL) was used for sugar determination by the anthrone method. Each data represents the mean and SD of three technical replicates. In each case, the SD remained below 10% of the mean. Recovery was approximately 100%.
Protocol validation on biomass
In addition to validation on mixtures of purified carbohydrates, the protocol was validated by conducting the complete process on 10 mg of three different batches of biomass from the microalgae C. reinhardtii (strains CC125, cw15, and sta6), C. sorokiniana, and S. obliquus containing on average 40%, 39%, 6%, 53% and 50% of total carbohydrates, respectively. Additionally, the extraction process was carried out on 10 mg of A. thaliana biomass under both control growth conditions and saline stress containing 21% and 27% of total carbohydrates in the biomass, respectively. The total carbohydrates recovered after fractionated extraction (the sum of the three fractions) was 95% ± 10%; 92% ± 6%; 99% ± 27%; 94% ± 1%, and 89% ± 5% for C. reinhardtii strains CC125, cw15 and sta6, C. sorokiniana, and S. obliquus, respectively, in comparison to whole biomass carbohydrates determination by the anthrone method. For A. thaliana, the total carbohydrates recovered were 98% ± 3% and 93% ± 2% for the control treatment and the saline stress treatment, respectively (Figure 4).
Figure 4. Validation of protocol on 10 mg of different microalgae and A. thaliana. A. Quantification of the different carbohydrates present in microalgal biomass and in A. thaliana. Samples of different microalgae and A. thaliana were subjected to the sequential extraction procedure. Strain sta6 of C. reinhardtii, which does not accumulate starch, was used as a negative control. Salt treatment is known to trigger sugars and starch accumulation in A. thaliana [18]. B. Recalcitrant carbohydrates present in the strains of C. reinhardtii. Each data represents the mean and SD of three biological replicates; statistical analysis was performed through one-way ANOVA (*, p ≤ 0.05). Recovery was typically 89%–99%.
For the validation of this protocol, three technical replicates of each sample were used. When quantifying carbohydrates with anthrone, each sample was measured in duplicate.
For the analysis of structural carbohydrates present in C. reinhardtii (Figure 4B), experimental data were statistically analyzed by one-way analysis of variance (ANOVA). Means and standard deviations were calculated and statistically examined using an analysis of variance and Dunnett’s multiple comparison test at p < 0.05.
C. reinhardtii mutant strains were particularly useful for validating this simple protocol. As expected, the most significant difference corresponded to a large decrease in the starch fraction in strain sta6 (Figure 4A). Additionally, although the cell wall of C. reinhardtii only contains a minor fraction of carbohydrates, clear differences were observed in this fraction. While the cell wall–deficient strain presented a level of carbohydrates approximately four-fold lower than the WT strain CC125, the sta6 mutant, which is impaired in starch biosynthesis, showed a level of carbohydrates approximately two-fold higher in this fraction (Figure 4B).
Fernandes et al. (2012) found very similar results for starch determination in C. vulgaris biomass by using extraction/hydrolysis by perchloric acid or amylolytic enzymes [19]. Another study conducted a more complex carbohydrate fractionation on C. vulgaris biomass. Sequential extraction with ethanol, di-ammonium oxalate (AmOx), NaOH (at two concentrations), and H2SO4 was followed by hydrolysis and sugars quantitation by HPLC. While starch was mainly recovered in the AmOx and diluted NaOH extractions, cell wall polysaccharides (likely chitin-like polymers and cellulose) were extracted/hydrolyzed by H2SO4 [20]. This protocol is recommended for an in-depth analysis of cell wall polysaccharides in microalgae. Despite differences in strains and culture conditions, carbohydrates partitioning, as determined by the method proposed here, aligns very well with these two reference studies, showing the relative abundance of carbohydrates in microalgae in the following order: starch, recalcitrant cell wall polysaccharides, and sugars.
Similarly, our protocol revealed similar induction levels of sugars and starch (approximately 50%) after salt stress in Thellungiella halophila, a plant closely related to A. thaliana. In that study, sugars were extracted in boiling water, and starch was extracted from the soluble fraction of 0.7 M perchloric acid, followed by precipitation with potassium perchlorate, and Glc determination using α-amylase and amyloglucosidase [21].
In summary, we propose a straightforward and reliable method that could be adapted for high-throughput screening to analyze carbohydrate partitioning in microalgae and plants.
Main advantages of the method
Efficiency and speed: Existing methods for carbohydrate partitioning can be time-consuming and labor-intensive. Thus, simplicity and the possibility of a high-throughput setting could significantly reduce the time and effort required for analysis. The method provides quicker results, especially when dealing with large sample sets.
Cost-effectiveness: Complex methods often involve expensive reagents, specialized equipment, and skilled personnel. Thus, this method could make carbohydrate analysis more accessible and cost-effective. Affordability could encourage broader adoption in research, quality control, and process optimization.
Wide applicability: The method was specifically optimized for determination from small samples of microalgae and A. thaliana seedlings, which are prime laboratory models in scientific and technological research that are frequently at the base of scientific and technological advances and breakthroughs.
General notes and troubleshooting
The viscosity of the acids makes it very important to vortex to achieve the desired results. In the case of cold starch extraction, it is important to adhere to incubation times and temperature to avoid extracting other carbohydrates belonging to the structural carbohydrate fraction.
Acknowledgments
Besides our own previous work, this protocol was modified from Waghmare et al. [2] and Updegraff [1].
This work was supported by grants from Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT, PICT2018-3382, PICT2019-2118) to L.C., and Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET PIP-11220200101701CO) and Universidad Nacional de Mar del Plata (UNMdP, EXA1051/21) to G.M.N. G.M.N, V.F.C. and L.C. are researchers at CONICET. A.N.B., L.S.R., A.D.M., and A.P.L. are Doctoral or Postdoctoral fellows at CONICET.
Competing interests
The authors declare no competing interests.
References
Updegraff, D. M. (1969). Semimicro determination of cellulose inbiological materials. Anal Biochem. 32(3): 420–424.
Waghmare, A. G., Salve, M. K., LeBlanc, J. G. and Arya, S. S. (2016). Concentration and characterization of microalgae proteins from Chlorella pyrenoidosa. Bioresour Bioprocess. 3(1): 16.
Chen, C. Y., Zhao, X. Q., Yen, H. W., Ho, S. H., Cheng, C. L., Lee, D. J., Bai, F. W. and Chang, J. S. (2013). Microalgae-based carbohydrates for biofuel production. Biochem Eng J. 78: 1–10.
Smith, A. M. and Zeeman, S. C. (2020). Starch: A Flexible, Adaptable Carbon Store Coupled to Plant Growth. Annu Rev Plant Biol. 71(1): 217–245.
Pontis, H. G. (2016). Methods for analysis of carbohydrate metabolism in photosynthetic organisms. Plants, green algae and cyanobacteria. Academic press, Elsevier.
Davies, D. R. and Plaskitt, A. (1971). Genetical and structural analyses of cell-wall formation in Chlamydomonas reinhardi. Genet Res. 17(1): 33–43.
Zabawinski, C., Van Den Koornhuyse, N., D'Hulst, C., Schlichting, R., Giersch, C., Delrue, B., Lacroix, J. M., Preiss, J. and Ball, S. (2001). Starchless Mutants of Chlamydomonas reinhardtii Lack the Small Subunit of a Heterotetrameric ADP-Glucose Pyrophosphorylase. J Bacteriol. 183(3): 1069–1077.
Baudelet, P. H., Ricochon, G., Linder, M. and Muniglia, L. (2017). A new insight into cell walls of Chlorophyta. Algal Res. 25: 333–371.
Somerville, C., Bauer, S., Brininstool, G., Facette, M., Hamann, T., Milne, J., Osborne, E., Paredez, A., Persson, S., Raab, T., et al. (2004). Toward a Systems Approach to Understanding Plant Cell Walls. Science. 306(5705): 2206–2211.
Ortiz-Marquez, J. C. F., Do Nascimento, M., Dublan, M. d. l. A. and Curatti, L. (2012). Association with an Ammonium-Excreting Bacterium Allows Diazotrophic Culture of Oil-Rich Eukaryotic Microalgae. Appl Environ Microbiol. 78(7): 2345–2352.
Do Nascimento, M., Ortiz-Marquez, J. C. F., Sanchez-Rizza, L., Echarte, M. M. and Curatti, L. (2012). Bioprospecting for fast growing and biomass characterization of oleaginous microalgae from South–Eastern Buenos Aires, Argentina. Bioresour Technol. 125: 283–290.
Bader, A. N., Sanchez Rizza, L., Consolo, V. F. and Curatti, L. (2020). Efficient saccharification of microalgal biomass by Trichoderma harzianum enzymes for the production of ethanol. Algal Res. 48: 101926.
Dreywood, R. (1946). Qualitative Test for Carbohydrate Material. Ind Eng Chem Anal Ed. 18(8): 499–499.
Gerwig, G. J. (2021). Detection of Carbohydrates by Colorimetric Methods. In: The art of carbohydrate analysis. Techniques in Life Science and Biomedicine for the non-expert. Springer, Cham.
Englyst, K. N., Hudson, G. J. and Englyst, H. N. (2000). Starch Analysis in Food. In: Meyers, R. A. (Ed.). Encyclopedia of Analytical Chemistry: ea1029.
Caló, G., De Marco, M. A., Salerno, G. L. and Martínez-Noël, G. M. A. (2022). TOR signaling in the green picoalga Ostreococcus tauri. Plant Sci. 323: 111390.
De Marco, M. A., Curatti, L. and Martínez-Noël, G. M. A. (2024). High auxin disrupts expression of cell-cycle genes, arrests cell division and promotes accumulation of starch in Chlamydomonas reinhardtii. Algal Res. 78: 103419.
Dong, S., Zhang, J. and Beckles, D. M. (2018). A pivotal role for starch in the reconfiguration of 14C-partitioning and allocation in Arabidopsis thaliana under short-term abiotic stress. Sci Rep. 8(1): 9314.
Fernandes, B., Dragone, G., Abreu, A. P., Geada, P., Teixeira, J. and Vicente, A. (2011). Starch determination in Chlorella vulgaris—a comparison between acid and enzymatic methods. J Appl Phycol. 24(5): 1203–1208.
Weber, S., Grande, P. M., Blank, L. M. and Klose, H. (2022). Insights into cell wall disintegration of Chlorella vulgaris. PLoS One. 17(1): e0262500.
Wang, X., Chang, L., Wang, B., Wang, D., Li, P., Wang, L., Yi, X., Huang, Q., Peng, M., Guo, A., et al. (2013). Comparative Proteomics of Thellungiella halophila Leaves from Plants Subjected to Salinity Reveals the Importance of Chloroplastic Starch and Soluble Sugars in Halophyte Salt Tolerance. Mol Cell Proteomics. 12(8): 2174–2195.
Supplementary information
The following supporting information can be downloaded here:
Supplementary information
Article Information
Publication history
Received: Jun 19, 2024
Accepted: Sep 26, 2024
Available online: Oct 16, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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5,122 | https://bio-protocol.org/en/bpdetail?id=5122&type=0 | # Bio-Protocol Content
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Development of a Rapid Epstein–Barr Virus Detection System Based on Recombinase Polymerase Amplification and a Lateral Flow Assay
YS Yidan Sun
DT Danni Tang
NL Nan Li
YW Yudong Wang
MY Meimei Yang
CS Chao Shen
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5122 Views: 221
Reviewed by: Emilia KrypotouChhuttan L MeenaJibin Sadasivan
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Original Research Article:
The authors used this protocol in Viruses Jan 2024
Abstract
The quality of cellular products used in biological research can impact the accuracy of results. Epstein–Barr virus (EBV) is a latent virus that spreads extensively worldwide, and cell lines used in experiments may carry EBV and pose an infection risk. The presence of EBV in a single cell line can contaminate other cell lines used in the same laboratory, affecting experimental results. Existing tests to detect EBV can be divided into three categories: nucleic acid assays, serological assays, and in situ hybridization assays. However, most methods are time-consuming, expensive, and not conducive to high-volume clinical screening. Therefore, a simple system that allows for the rapid detection of EBV in multiple contexts, including both cell culture and tissue samples, remains necessary. In our research, we developed EBV detection systems: (1) a polymerase chain reaction (PCR)-based detection system, (2) a recombinase polymerase amplification (RPA)-based detection system, and (3) a combined RPA-lateral flow assay (LFA) detection system. The minimum EBV detection limits were 1 × 103 copy numbers for the RPA-based and RPA-LFA systems and 1 × 104 copy numbers for the PCR-based system. Both the PCR and RPA detection systems were applied to 192 cell lines, and the results were consistent with those of the assays specified in industry standards. A total of 10 EBV-positive cell lines were identified. The combined RPA-LFA system is simple to operate, allowing for rapid result visualization. This system can be implemented in laboratories and cell banks as part of a daily quality control strategy to ensure cell quality and experimental safety and may represent a potential new technique for the rapid detection of EBV in clinical samples.
Key features
• Establishes RPA and RPA-LFA detection systems for EBV.
• The RPA-LFA detection system can visualize the results in as short as 15 min.
• The specificity of the RPA and RPA-LFA assay systems for the detection of EBV is validated.
• The minimum EBV detection limit of the RPA and RPA-LFA systems is 1 × 103 copies.
Keywords: RPA LFA EBV Cell quality control Rapid detection
Graphical overview
Pattern of Epstein–Barr virus (EBV) detection by the PCR, recombinase polymerase amplification (RPA), and combined RPA-lateral flow assay (LFA) systems. Reaction pellets are small particles that contain key reagents. These pellets usually contain the core components needed for the RPA reaction (recombinase, recombinase loading factor, and single-stranded binding protein). At the start of the reaction, reaction pellets dissolve these components to initiate the amplification reaction.
Background
As biological research continues to advance, cells are increasingly at risk of being misidentified or contaminated with other cell types or exogenous factors [1,2] including viruses; such contamination can be difficult to detect and remediate.
Epstein–Barr virus (EBV) is a member of the human herpesvirus family. EBV displays prolonged latency in lymphocytes, interfering with immune functions and potentially inducing cell proliferation and transformation. EBV infection involves many organ systems and is often misdiagnosed or underdiagnosed. Multiple reports in the literature have described the detection of EBV infection in various cell types maintained in cell culture banks [3,4]. These infections can be attributed to the presence of an existing EBV infection during the initial process of cell line establishment, EBV contamination of culturing materials, or improper manipulation by the experimental staff.
The most widely used of the existing EBV assays is PCR, which is a well-established and reliable molecular method. Quantitative PCR (qPCR) can be used to monitor disease progression or therapeutic efficacy by detecting changes in EBV load in the blood [5]. Iso-thermal amplification techniques, such as recombinant enzyme-mediated isothermal amplification and loop-mediated isothermal amplification (LAMP), are easy to perform, have limited equipment requirements, return rapid responses, and display high sensitivity. However, designing primers for use in the LAMP technique is complicated and can be difficult to replicate. In situ hybridization assay to detect Epstein–Barr early RNA (EBER) is the current gold standard for detecting EBV infection in clinic. However, this technique can only be applied to tissue samples and is generally limited to the clinical diagnosis of EBV-related diseases, such as cancer and lymphoma.
Recombinase polymerase amplification (RPA) is a highly sensitive and selective isothermal amplification technique that can be performed at 39 °C and can be used to amplify a large number of samples in a short period of time. RPA, first introduced in 2006 by Niall Armes of ASM Scientific Ltd., UK [6], relies on modified homologous recombination mechanisms. RPA reagents are commercially available, and the basic RPA reaction kit can be augmented by additional commercially available kits that use different probes that can be cleaved by different enzymes: exo (exonuclease III), fpg (formamidopyrimidine DNA Glycosylase), and nfo (endonuclease IV) [7]. The exo and fpg probes are typically used for real-time detection, whereas the nfo probe is typically used for detection systems that use lateral flow dipsticks.
In this study, we combined RPA technology with a lateral flow assay (LFA) to develop an RPA-LFA detection system for application to the rapid and bulk screening of EBV contamination in cell lines stored by cell banks and to conduct regular and daily inspection of cell lines used in biological experiments, ensuring the quality of cell lines and the safety of experimental personnel. This system also demonstrates high potential for clinical adaptation to improve EBV detection in blood and tissue samples.
Materials and reagents
Biological materials
B95-8 cell line (China Center for Type Culture Collection, GDC0015)
HeLa cell line (China Center for Type Culture Collection, GDC0009)
Epstein–Barr virus (China Center for Type Culture Collection, GDV132)
Reagents
Trypsin (AMEKO, catalog number: RC01145)
Ethanol (National Pharmaceutical Group Chemical Reagent Co., Ltd., catalog number: 10009228)
PBS (Gibco, catalog number: 70013032)
TIANamp Genomic DNA Kit (TIANGEN, catalog number: DP304)
E.Z.N.A Viral DNA Kit (OMEGA, catalog number: D3892)
Takara 10× loading buffer (Takara, catalog number: AA0151)
DL2,000 DNA marker (Takara, catalog number: 3427A)
TS-GelRed nucleic acid dye (TSINGKE, catalog number: TSJ003)
Agarose (TSINGKE, catalog number: TSJ001)
RNase-free water (Takara, catalog number: takara.9012)
Twist Amp Basic kit (TwistDX, catalog number: 111-035-003)
Thermo Scientific endonuclease IV(Nfo) and its buffer (FastDigest, catalog number: EN0591)
Test strip dilution (Aoke Botai Biotech)
Primer and probe:
FP: 5'-CTTGGAGACAGGCTTAACCAGACTCA-3'
RP: 5'-CCATGGCTGCACCGATGAAAGTTAT-3'
LFA-RP: 5'-[BIOTIN]CCATGGCTGCACCGATGAAAGTTAT-3'
LFA-Probe: 5'-[FITC]TGCCGGCCCCTCGAGATTCTGACCGGGGACC[THF]CTGGTTGCTCTGTTG[C3-Spacer]-3'
Laboratory supplies
Cell culture flask (NEST, catalog number: 707001)
Serological pipettes (NEST, catalog number: 327001)
Pipette tips (Biosharp, catalog numbers: BS-10-T, BS-200-T, BS-1000-T)
Micro-centrifuge tube (Biosharp, catalog number: BS-15-M)
Axygen PCR tubes (AXYGEN, catalog number: Axygen- PCR-05-A)
Lateral flow stick (Suzhou Xianda Gene Technology, catalog number: TS101)
Equipment
Biological safety cabinet (Esco, catalog number: A2AC2-S)
Refrigerator [CHANGHONG MEILING, catalog number: MCF(L)-398LDWEP]
Thermostatic water bath (Dichbio, catalog number: JB Academy)
PCR automatic serialization analyzer (SensoQuest, catalog number: Labcycler 48)
Low-speed centrifuge (DLAB, catalog number: D1008E, D1008)
Small high-speed freezer-type centrifuge (Eppendorf, model: 5420)
CO2 Incubator (Memmert, catalog number: INC)
Ultra-micro UV spectrophotometer (Thermo Fisher Scientific, model: NanoDrop2000)
Micropipettes 0.1–2.5 µL, 2–20 µL, 20–200 µL (Eppendorf, catalog number: 3123000217, 3123000233, 3123000250)
Electrophoresis apparatus (Beijing Liuyi Biotechnology Co., Ltd., catalog number: 112-0630)
Gel imaging system (Bio-Rad, catalog number: ChemiDoc XRS+)
Vortex mixer (Kylin-bell, catalog number: VORTEX-6)
Software and datasets
DNAMAN (version 8.0)
National Center for Biotechnology Information (https://www.ncbi.nlm.nih.gov/)
Procedure
Primer & probe & lateral flow strip design
General primer design
Download the nucleic acid sequence of EBV from NCBI (Genebank: V01555).
Design PCR and RPA primers for conserved regions using DNAMAN 8.0 software.
Considering that the length of RPA primers is longer than that of normal PCR primers, after primer testing, we finally designed primers that can participate in both PCR and RPA as follows:
Forward primer: 5'-CTTGGAGACAGGCTTAACCAGACTCA-3'
Reverse primer: 5'-CCATGGCTGCACCGATGAAAGTTAT-3'
The position of the Epstein–Barr virus (EBV) fragment detected by primers is illustrated in Figure 1A.
Figure 1. Schematic diagram of primer screening and lateral flow strip design. (A) Location of the Epstein–Barr virus (EBV) fragment detected by primers on the BamHI restriction map of the prototype EBV B95-8 genome. (B) Lateral flow strip design.
RPA-LFA primer & probe design
The same amplified fragments as in PCR were selected but with BIOTIN modification at the 5' end of the reverse primer and a probe introduced to enhance its amplification specificity.
The probe was designed to have a FITC modification at the 5' end, a modifying group (C3-Spacer) at the 3' end, and tetrahydrofuran (THF) labeled in the middle of the 5' and 3' ends to serve as the recognition site for NFO. The final primers and probes were as follows:
Forward primer: 5'-CTTGGAGACAGGCTTAACCAGACTCA-3'
LFA-Reverse primer: 5'-[BIOTIN]CCATGGCTGCACCGATGAAAGTTAT-3'
LFA-Probe: 5'-[FITC]TGCCGGCCCCTCGAGATTCTGACCGGGGACC[THF]CTGGTTGCTCTGTTG[C3-Spacer]-3'
Lateral flow strip design
Gold particles of BIOTIN antibody and FITC antibody are immobilized on a lateral flow strip (test-line), and the target fragment will be captured to appear as a positive band. (Analysis of results: If only one control line can be observed, the result is regarded as negative; if both the control line and the test line can be observed at the same time, the result is regarded as positive; if the control line is not observed, it indicates that the test strip is invalid.) The structure of the test strip is shown in Figure 1B.
Sample preparation
DNA extraction
Collect cells in good growth condition, centrifuge with trypsin digestion, and aspirate the cell suspension. If cells are freeze-stored, directly thaw and transfer to microcentrifuge tubes and aspirate the cell suspension. The latter steps are operated according to the blood/cell/tissue genomic DNA extraction kit of Tiangen Biochemical Technology (Beijing) Co. as follows:
Place the pre-treated cell suspension (containing a cell number approximately equal to the number of cells that can be cultured with one t25) into a 1.5 mL microcentrifuge tube, centrifuge at 9,306× g for 1 min, remove the supernatant, and add 1 mL of PBS reagent to keep the cells in suspension. Resuspend to rewash the cell precipitate, centrifuge at 9,306× g for 1 min, remove the supernatant, add 200 μL of buffer solution GA, and shake the mixture until it is fully suspended.
Add 20 μL of proteinase K solution to the bottom of the micro-centrifuge tube and mix well; then, add 200 μL of buffer GB to it and mix well. Place the micro-centrifuge tube in a thermostat at 70 °C for 10 min and centrifuge briefly to remove the water droplets on the inner wall of the cap.
Add 200 μL of anhydrous ethanol to it, shake it well for 15, and centrifuge it briefly to remove the water droplets on the inner wall of the cap.
Add the liquid from this step to the adsorbent column and centrifuge at 13,400× g for 30 s. Discard the flowthrough liquid.
Add 500 μL of buffer GD with anhydrous ethanol to the column and centrifuge at 13,400× g for 30 s. Discard the waste liquid.
Add 600 μL of rinse solution PW to the column, centrifuge at 13,400× g for 30 s, and discard the waste solution. Repeat step B1g with another addition of rinse solution PW.
Place the column back into the collection tube, centrifuge at 13,400× g for 2 min, discard the waste liquid, and let dry thoroughly. Prepare a clean micro-centrifuge tube and place the column in it. Add 50 μL of ddH2O dropwise to the column and centrifuge at 13,400× g for 2 min. The resultant liquid obtained after centrifugation is the cellular DNA.
DNA dilution
Measure the DNA concentration using a NanoDrop microspectrophotometer. Calculate concentration by diluting 50 ng/μL with ddH2O and then store in the refrigerator at -20 °C.
Nucleic acid amplification
RPA-specific assay
Cellular DNA diluted with ddH2O at a concentration of 50 ng/μL is selected as the template in the system. In the reaction tube containing the RPA lyophilized powder, add 29.5 μL of buffer, 2 μL each of unlabeled 10 μM upstream and downstream primers, 1 μL of cellular extracted DNA, and 13 μL of ddH2O. Finally, add a 280 mmol/L solution of magnesium acetate in a volume of 2.5 μL to initiate the reaction. Uniformly distribute the above reaction solution and place it into the PCR instrument at 39 °C for 20 min. After the amplification reaction is completed, add 6 μL of 10× loading buffer and thoroughly mix. Then, load 10 μL of the sample into the wells of a 2% agarose gel and operate on agarose gel electrophoresis (120 V, 200 A). The amplification of the target bands is observed using a gel-image system (Figure 2A).
Figure 2. Establishment of recombinase polymerase amplification (RPA) testing system. (A) Validation of the RPA system. 1: ddH2O; 2: HeLa cell DNA (negative control); 3: B95-8 cell DNA (positive control); M: DL2000 marker. (B) Determination of the optimal RPA time for B95-8 cell DNA. 1: 5 min; 2: 10 min; 3: 15 min; 4: 20 min; M: DL2000 marker. (C) Determination of the optimal RPA temperature for B95-8 cell DNA. 1: 43 °C; 2: 41 °C; 3: 39 °C; 4: 37 °C; 5: 35 °C; M: DL2000 marker. (D) Sensitivity of Epstein–Barr virus (EBV) detection by the RPA method. M: DL2000 marker; 1: 1 × 106 copies/μL; 2: 1 × 105 copies/μL; 3: 1 × 104 copies/μL; 4: 1 × 103 copies/μL; 5: 1 × 102 copies/μL; 6: 1 × 101 copies/μL; 7: 1 × 100 copies/μL; M: DL2000 marker.
Optimization of RPA reaction system
B95-8 cell DNA at 50 ng/μL is used as a template and optimized in terms of time and temperature, respectively. Thoroughly mix the resulting product with 6 μL of 10× loading buffer and confirm the amplification efficiency by agarose gel electrophoresis (120 V, 200 A).
Time optimization: Maintain reaction temperature at 39 while the reaction time is varied at 5, 10, 15, and 20 min to observe the amplification of different reaction times (Figure 2B).
Temperature optimization: Maintain reaction time at 15 min, and vary the reaction temperatures at 35, 37, 39, 41, and 43 °C to observe the amplification of RPA reaction at different temperatures (Figure 2C).
RPA sensitivity examination
Dilute the previously prepared EBV DNA with a ddH2O gradient to 1 × 106, 1 × 105, 1 × 104, 1 × 103, 1 × 102, 1 × 101, 1 × 100 copies/μL as templates. Subsequently, use 1 μL of the diluted DNA for ordinary RPA amplification at 39 °C for 15 min. Following this amplification process, add 6 μL of 10× loading buffer to the reaction mixture. After thorough mixing, take a sample volume of 10 μL for agarose gel electrophoresis (120 V, 200 A) to verify the amplification effect (Figure 2D).
Lateral flow assay
RPA-LFA specific assay
Set up the RPA reaction with a common forward primer, a reverse primer with Biotin labeling, buffer, DNA template, ddH2O, and magnesium acetate. The reaction takes place at 39 °C for 20 min.
Add the FITC-labeled probe, nfo enzyme, and nfo enzyme buffer to the reaction product from the previous step. The specific reaction system is as follows:
Buffer 29.5 μL
10 μM FP 2 μL
10 μM LFA-RP 2 μL
10 μM probe 0.6 μL
280 mmol/L magnesium 2.5 μL
ddH2O 13 μL
RPA amplification products 50 μL
Template DNA 1 μL
NFO enzyme 1.5 μL
NFO enzyme buffer 5 μL
Dilute the final amplification product with diluent at a ratio of 1:100. Then, take 50 μL and add it to the lateral flow strip. Wait for 5 min to observe the results (Figure 3A).
Optimization of RPA-LFA reaction system
Optimize the reaction conditions of RPA-LFA, such as reaction time and temperature, using 50 ng/μL B95-8 cellular DNA as the reaction template. Dilute the amplification products at a ratio of 1:100 and then add dropwise to the test strips to wait for the detection results.
Time optimization: 1) Optimization of the first part of the reaction. The reaction time of the first step for common RPA is set to 5, 10, 15, and 20 min. After adding the probe, nfo enzyme, and its buffer, set the amplification and digestion time to 15 min. Then, test the resulting amplification product using a test strip (Figure 3B). 2) Optimization of the reaction time for the second step. The time of the first step is set at 5 min, and the time of the second step of enzymatic amplification is set at 5, 10, and 15 min. Add the amplification products obtained dropwise to the test strip to observe the results (Figure 3C).
Reaction temperature: Maintain the reaction time of both steps at 15 min and vary the reaction temperatures of the samples in both steps at 35, 37, 39, 41, and 43 °C, respectively (Figure 3D).
RPA-LFA sensitivity examination
Dilute the EBV DNA to 1 × 106, 1 × 105, 1 × 104, 1 × 103, 1 × 102, 1 × 101, 1 × 100 copies/μL in a gradient, and take 1 μL of each concentration as the template for amplification. Set the temperature of the reaction to 39 °C and the amplification time for both steps to 15 min. After amplification, proportionally dilute 50 μL and add to the test strip for 5 min to observe the results (Figure 3E).
Figure 3. Establishment of combined recombinase polymerase amplification–lateral flow assay (RPA-LFA) testing system. (A) Validation of the RPA-LFA system. a: ddH2O; b: HeLa cell DNA (negative control); c: B95-8 cell DNA (positive control). (B) RPA-LFA was used to assess assays with different reaction times (first step). a: 5 min; b: 10 min; c: 15 min; d: 20 min; (C) RPA-LFA was used to assess results for assays with different reaction times (second step). a: 5 min; b: 10 min; c: 15 min. (D) Determination of the optimal RPA-LFA temperature for B95-8 cell DNA. a: 35 °C; b: 37 °C; c: 39 °C; d: 41 °C; e: 43 °C. (E) Sensitivity of Epstein–Barr virus (EBV) detection by the RPA method. M: DL2000 marker; a: 1 × 106 copies/μL; b: 1 × 105 copies/μL; c: 1 × 104 copies/μL; d: 1 × 103 copies/μL; e: 1 × 102 copies/μL; f: 1 × 101 copies/μL; g: 1 × 100 copies/μL.
Data analysis
Determination of primer and probe specificity.
The primers and probes were tested for specificity by RPA and RPA-LFA assay and the results showed good specificity of the primers and probes. To further validate their specificity, 192 cells from different species were validated using RPA. The results are shown in the Supplementary Information. In order to ensure the accuracy of the experiment, we also chose two other EBV detection methods, PCR and qPCR. The detection results of these three methods are shown in Table 1 below:
Table 1. Evaluation of EBV test results using different methods of detection
Result Method
PCR qPCR RPA
Positive 9 9 9
Negative 183 183 183
Total 192 192 192
To verify the specificity of the RPA-LFA system, we screened several cell lines containing endogenous viruses among the 192 tested cell lines: HeLa (containing human papillomavirus type 18), SiHa (containing human papillo-mavirus type 16), Hep-G2/2.2.15 (containing hepatitis B virus), and RK13 (containing bovine viral diarrhea virus). The RPA-LFA method was validated using these cell lines, and the results were negative for EBV (Figure 4A), indicating that the system has good specificity. In addition, we randomly selected cell lines for further validation, including three EBV-negative cell lines (A-204, H9, and HTR-8) and the remaining nine cell lines (NK-92, Daudi, ARH-77, Raji, JVM-2, A-431, Farage, MC/CAR, and CCRF-SB) that showed EBV positivity in previous tests. All cell detection results were consistent with expectations.
Figure 4. Combined recombinase polymerase amplification–lateral flow assay (RPA-LFA) detection of Epstein–Barr virus (EBV) system in different cells. (A) Detection of cells containing other endogenous viruses using the RPA-LFA system. a: HeLa; b: SiHa; c: Hep-G2/2.2.15; d: RK13. (B) RPA-LFA detection of EBV-negative cell lines. a: A-204; b: H9; c: HTR-8. (C) RPA-LFA detection of EBV-positive cell lines. a: NK-92; b: Daudi; c: ARH-77; d: Raji; e: JVM-2; f: A-431; g: Farage; h: MC/CAR; i: CCRF-SB.
Reaction condition optimization.
Through the optimization experiments of RPA reaction conditions, it was found that the optimal reaction time is 15 min, and the optimal reaction temperature is 39 °C. In the optimization experiment of RPA-LFA reaction conditions, it was observed that positive results could still be obtained when both the first and the second steps were reacted for 5 min at a temperature of 39 °C.
Determination of detection limit.
We constructed a plasmid containing the EBV gene and used this for qPCR to plot an EBV standard curve according to the CT value (Figure 5). The CT value of 100 copies/μL was discarded because the deviation of this point was large, and a standard curve was plotted. The CT value was inversely proportional to the gene copy number, and the R2 value of the curve was 0.99. Based on this standard curve, the concentration of EBV DNA extracted from B95-8 cells was calculated, and the DNA was diluted with ddH2O to 1 × 106 copies/μL. The EBV DNA was further diluted with ddH2O to produce a gradient of 1 × 106, 1 × 105, 1 × 104, 1 × 103, 1 × 102, 1 × 101, and 1 × 100 copies/μL.
The limit of detection is determined by the presence or absence of the test line. Based on the results of experiments in which viruses of different concentration gradients were used as templates, the following could be concluded: In the RPA and RPA-LFA experiments, bands were still observed on the test line when the copy number of template EBV was 1×103copies, so the limit of detection for the RPA and RPA-LFA assays were 1 × 103 copies (Figure 2D; Figure 3E).
Figure 5. Epstein–Barr virus (EBV) standard curve
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Sun et al. [8] Development of a Rapid Epstein–Barr Virus Detection System Based on Recombinase Polymerase Amplification and a Lateral Flow Assay.
General notes and troubleshooting
General notes
Please pay attention to cross-contamination and aerosol contamination during operation.
The length of RPA primer should be around 30 bp (a primer too short will affect the amplification speed and detection sensitivity).
When adding reaction reagents to RPA, it is recommended to add magnesium ions at the end to activate the reaction.
Troubleshooting
Problem: The drag band is obvious after agarose gel electrophoresis with RPA.
Possible cause: The reaction system is not mixed well; the agarose gel concentration is too low.
Solution: Ensure thorough mixing of the reaction system during preparation and add Mg ions to the lid at the end; centrifuge and shake down to activate the reaction; prepare an agarose gel with a concentration of 2% or higher.
Acknowledgments
This work was supported by grants from the National Science and Technology Infrastructure (NSTI-BMCR20, NSTI-BMCR21, and NSTI-BMCR22) and Science and Technology Innovation Grants of Hubei Province (2021CFB357) to Chao Shen.
The protocols described here are adapted from our previous work (Sun et al. [8]).
Competing interests
The authors declare no competing interests.
References
Stacey, G. N. (2000). Cell contamination leads to inaccurate data: we must take action now. Nature. 403(6768): 356–356.
Hughes, P., Marshall, D., Reid, Y., Parkes, H. and Gelber, C. (2007). The Costs of using Unauthenticated, Over-Passaged Cell Lines: How Much more Data do we Need? Biotechniques. 43(5): 575–586.
Uphoff, C. C., Denkmann, S. A., Steube, K. G. and Drexler, H. G. (2010). Detection of EBV, HBV, HCV, HIV-1, HTLV-I and -II, and SMRV in Human and Other Primate Cell Lines. J Biomed Biotechnol. 2010: 1–23.
Shioda, S., Kasai, F., Watanabe, K., Kawakami, K., Ohtani, A., Iemura, M., Ozawa, M., Arakawa, A., Hirayama, N., Kawaguchi, E., et al. (2018). Screening for 15 pathogenic viruses in human cell lines registered at the JCRB Cell Bank: characterization of in vitro human cells by viral infection. R Soc Open Sci. 5(5): 172472.
Hübner, M., Bozic, M., Konrad, P. M., Grohs, K., Santner, B. I. and Kessler, H. H. (2015). Analytical and clinical performance of a new molecular assay for Epstein-Barr virus DNA quantitation. J Virol Methods. 212: 39–43.
Piepenburg, O., Williams, C. H., Stemple, D. L. and Armes, N. A. (2006). DNA Detection Using Recombination Proteins. PLoS Biol. 4(7): e204.
Lobato, I. M. and O'Sullivan, C. K. (2018). Recombinase polymerase amplification: Basics, applications and recent advances. TrAC, Trends Anal Chem. 98: 19–35.
Sun, Y., Tang, D., Li, N., Wang, Y., Yang, M. and Shen, C. (2024). Development of a Rapid Epstein–Barr Virus Detection System Based on Recombinase Polymerase Amplification and a Lateral Flow Assay. Viruses. 16(1): 106.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. RPA detection of EBV system in different cells.
Article Information
Publication history
Received: Aug 1, 2024
Accepted: Oct 4, 2024
Available online: Oct 16, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
How to cite
Category
Microbiology > Pathogen detection > PCR
Molecular Biology > DNA > PCR
Medicine
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Compartment-Resolved Proteomics with Deep Extracellular Matrix Coverage
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Original Research Article:
The authors used this protocol in Journal of Proteome Research Feb 2023
Abstract
The extracellular matrix (ECM) is a complex network of proteins that provides structural support and biochemical cues to cells within tissues. Characterizing ECM composition is critical for understanding this tissue component’s roles in development, homeostasis, and disease processes. This protocol describes an integrated pipeline for profiling both cellular and ECM proteins across varied tissue types using mass spectrometry–based proteomics. The workflow covers stepwise extraction of cellular and extracellular proteins, enzymatic digestion into peptides, peptide cleanup, mass spectrometry analysis, and bioinformatic data processing. The key advantages include unbiased coverage of cellular, ECM-associated, and core-ECM proteins, including the fraction of ECM that cannot be solubilized using strong chaotropic agents such as urea or guanidine hydrochloride. Additionally, the method has been optimized for reproducible ECM enrichment and quantification across diverse tissue samples. This protocol enables systematic mapping of the ECM at a proteome-wide scale.
Key features
• Improved profiling of core extracellular matrix and matrisome-associated proteins through multi-step decellularization and chemical extraction of insoluble ECM
• Extraction buffers optimized for effectiveness across a broad range of tissue types and compatibility with varied MS platforms
• Measurement of protein solubility via resistance to detergent and chaotrope extraction
• Integrated LC-MS/MS analysis and data processing pipeline for ECM-focused analysis
Keywords: proteomics mass spectrometry extracellular matrix matrisome collagen protein extraction
Graphical overview
ECM-focused protein extraction and analysis workflow
Background
The extracellular matrix (ECM) is a complex biomolecular network composed of collagens, glycoproteins, proteoglycans, and other components that provide structural support and regulate the function of the ECM [1]. The ECM provides essential physical scaffolding for cellular constituents, as well as critical biochemical and biomechanical cues that regulate tissue development, homeostasis, and regeneration [2]. Given its fundamental roles, dysregulation of the ECM is implicated in numerous pathologies including fibrosis [3,4], cardiovascular diseases [5,6], and cancer progression [7]. Comprehensive mapping of ECM composition is crucial for understanding the diverse functions of this intricate network across different tissue microenvironments and disease states. Early efforts to catalog the ECM focused on profiling a subset of known ECM proteins [8] or employed antibody-based approaches [9], which are inherently biased and limited in coverage. More recently, mass spectrometry (MS)-based proteomics has enabled unbiased and system-wide profiling of the ECM at a proteome scale. However, technical challenges in extracting and enriching the ECM while minimizing contamination from cellular proteins have impeded comprehensive ECM characterization across a wide range of tissue types and organs. Previous analyses have shown that more than 80% of the total fibrillar collagen in many tissues resides in a chaotrope-resistant insoluble ECM fraction and is not solubilized by traditional proteomic extraction methods [10], highlighting the need for additional extraction steps when comprehensive characterization of ECM composition is a priority (Figure 1).
Figure 1. Proportion of total collagen I identified in the cellular, soluble extracellular matrix (sECM) and insoluble ECM (iECM, requiring chemical digestion) fractions of a 3-step ECM extraction via QconCAT-based absolute quantification. Adapted from Goddard et al. [10]. This protocol outlines an integrated experimental and computational pipeline tailored for deep proteomic profiling of the ECM across multiple mouse tissues and organs. The methods here have been refined across a broad range of tissue types, including 25 different mouse organs [11], and have been comprehensively benchmarked against other prominent methods in the field to validate their performance [12]. The workflow covers 1) enhanced step-wise extraction of cellular and ECM proteins facilitated by optimized extraction buffers effective across varied tissue types; 2) enzymatic digestion of proteins into peptides; 3) subsequent sample quantification and cleanup for MS analysis; 4) tandem mass spectrometry (LC-MS/MS) analysis of cellular and ECM protein fractions; and 5) bioinformatic identification of peptide sequences and computational matrisome categorization to classify ECM and associated proteins.
Materials and reagents
Reagents
Tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl) (Roche, catalog number: 10812846001)
3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonate (CHAPS) (Sigma-Aldrich, catalog number: C3023)
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: E4884)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
Halt protease inhibitor cocktail (Thermo Fisher, catalog number: 78429)
Guanidine hydrochloride (Gnd-HCl) (Sigma-Aldrich, catalog number: G4505)
Hydroxylamine-HCl (NH2OH–HCl) (Sigma-Aldrich, catalog number: 159417)
Ammonium bicarbonate (ABC) (Sigma-Aldrich, catalog number: A6141)
Double-distilled water (ddH2O)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D9779)
Iodoacetamide (IAM) (Sigma-Aldrich, catalog number: I1149)
Potassium carbonate (K2CO3) (Sigma-Aldrich, catalog number: 209619)
Sodium hydroxide (NaOH) (Sigma-Aldrich, catalog number: S5881)
Urea (Sigma-Aldrich, catalog number: U5128)
Buffer A: 0.1% formic acid (FA) in H2O, Optima (Thermo Fisher, catalog number: LS118)
Buffer B: 0.1% FA in 80% ACN, Optima (Thermo Fisher, catalog number: LS122500)
Sequencing-grade modified trypsin (Promega, catalog number: V5111)
Lysyl Endopeptidase®, mass spectrometry grade (Lys-C) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 125-05061)
Optional (if necessary based on protocol notes):
Gel-loading pipette tips (Corning, catalog number: CLS4853)
OptimaTM acetone (Thermo Fisher, catalog number: A929)
PierceTM 660 nm protein quantification assay (Thermo Fisher, catalog number: 22660)
Solutions
High salt buffer (see Recipes)
Gnd-HCl solution (see Recipes)
DTT solution (see Recipes)
IAM solution (see Recipes)
Hydroxylamine (HA) buffer preparation (pH 9.0) (see Recipes)
Urea solution (see Recipes)
Digest master mix (per sample) (see Recipes)
Recipes
High salt buffer
Reagent Final concentration Amount
Tris-HCl (1M, pH 7.4) 50 mM 5 mL
CHAPS 0.25% 25 mg
EDTA (500 mM) 25 mM 5 mL
NaCl 3 M 17.53 g
Halt protease inhibitor (100×) 1× 1 mL
Total n/a 100 mL
Combine buffer components and bring to 100 mL with ddH2O. Buffer without protease inhibitors can be prepared in advance and stored at 4 °C for up to one month. Protease inhibitors must be added fresh before use.
Gnd-HCl solution
Reagent Final concentration Amount
Gnd-HCl 6 M 57.32 g
ABC (1 M, pH 8.0) 100 mM 10 mL
Total n/a 100 mL
Dissolve Gnd-HCl in ddH2O before adding 10 mL of 1 M ABC. Adjust pH to 9.0 using NaOH. Add ddH2O to 100 mL. Prepare fresh before use.
DTT solution
Reagent Final concentration Amount
DTT 100 mM 15.4 mg
ABC (1 M) 100 mM 1 mL
Total n/a 10 mL
Combine components and add ddH2O to 10 mL. Prepare fresh before use.
IAM solution
Reagent Final concentration Amount
IAM 500 mM 92.5 mg
ABC (1 M) 100 mM 1 mL
Total n/a 10 mL
Combine components and add ddH2O to 10 mL. Prepare fresh before use and store in the dark.
Hydroxylamine (HA) buffer preparation (pH 9.0)
Reagent Final concentration Amount
Gnd-HCl solution (Recipe 2) 70% v/v 70 mL
NH2OH–HCl 1 M 7 g
NaOH (50% w/v) 4% w/v 8 mL
K2CO3 (1 M) 0.2 M 20 mL
Total n/a 100 mL
Prepare Gnd-HCl solution according to Recipe 2. Dissolve NH2OH–HCl in 50 mL of Gnd-HCl solution. Add NaOH, mix, then add K2CO3. Add Gnd-HCl solution to 90 mL before adjusting pH to 9.0 using HCl. Add Gnd-HCl solution to 100 mL. Prepare fresh before use.
Urea solution
Reagent Final concentration Amount
Urea 8 M 24.02 g
ABC (1 M) 100 mM 5 mL
Total n/a 50 mL
Combine components and add ddH2O to 50 mL. Prepare fresh before use.
Digest master mix (per sample)
Reagent Final Concentration Amount
ABC (1 M) 25 mM 2.5 μL
Lys-C 1:400 enzyme:protein ratio 0.075 μg (for 30 μg protein)
Total n/a 100 μL
Laboratory supplies
MS-compatible pipette tips (any brand)
Safe-Lock 1.75 mL tubes (Eppendorf, catalog number: 003012361)
2 mL snap cap tubes (any brand)
Glass beads, 1 mm diameter (Next Advance, catalog number: GB10)
10 kDa molecular weight cutoff (MWCO) filters (Sartorius, catalog number: VN01H02)
PierceTM quantitative colorimetric peptide assay (Thermo Fisher, catalog number: 23275)
Flat-bottom, clear 96-well plate (Grenier, catalog number: 655101)
Empore sealing tape for 96-well plate (Millipore Sigma, catalog number: 66881-U)
PierceTM C18 spin tips (Thermo Fisher, catalog number: 84850)
Verex 300 μL vials with pre-slit cap (Phenomenex, catalog number: AR0-39S0-13-C/AR0-8972-13-C)
Equipment
Ceramic mortar and pestle (any brand)
Measuring pipettes, p1000, p200, p10 (any brand)
FreeZone 4.5L benchtop freeze dryer (Labconco, catalog number: 7750020)
Clear complete fast-freeze flask, any size (Labconco)
Scroll vacuum pump (Labconco, catalog number: 7587000)
Bullet blender (Next Advance, model: BBX24)
Centrifuge (Eppendorf, model: 5430R)
Vortex mixer with 1.5 mL tube attachment (Labnet, model: VX-200)
Orbital shaker, Orbit M60 (Labnet, catalog number: LS-2020-M60)
Spark® microplate reader (TECAN, catalog number: 30086376)
CentriVap vacuum concentrator (Labconco, catalog number: 7810016)
Fusion Lumos Tribrid mass spectrometer (Thermo Fisher)
EASY-nLC 1200 System (Thermo Fisher, catalog number: LC140)
Analytical column made in-house: 100 μm i.d. × 150 mm fused silica capillary packed with 2.7 μm CORTECS C18 resin (Waters). Commercial alternative: EASY-Spray C18 column, 75 μm i.d. × 150 mm fused silica capillary packed with 3 μm resin (Thermo Fisher, catalog number: ES900)
Software and datasets
XCalibur Version 4.5 (Thermo Fisher)
Mascot Server Version 2.5 (Matrix Science)
Proteome Discoverer Version 2.5 (Thermo Fisher)
MSFragger (Nesvizhskii Lab) [13]
UniProtKB (https://www.uniprot.org/help/uniprotkb)
CRAPome Contaminant Database [14]
MatrisomeDB [15]
Procedure
Tissue preparation
Mill flash-frozen tissue samples in liquid nitrogen using a mortar and pestle until a fine powder is produced.
Lyophilize milled, frozen tissue samples in a clean fast-freeze flask (Labconco) of appropriate size for your sample set, ensuring that vessel pressure remains below 400 μBar during the entire lyophilization process.
Lyophilization is strongly recommended to increase the ease of sample handling and to avoid sample thawing (and subsequent enzymatic activity) during processing. However, it is required only when tissue dry weights are desired. If performing this extraction method on fresh tissue, proceed to step B1.
Once samples have been completely dried (5–16 h depending on tissue type and size), remove samples from the lyophilizer.
After lyophilization, store milled and dried samples in 1.75 mL safe-lock tubes at -80 °C until further processing. After removing samples from the -80 °C freezer, allow samples to come fully to room temperature before opening the tube lids to prevent condensation from forming on the sample and the inner tube walls.
Decellularization
Weigh approximately 5 mg (dry weight) of milled, lyophilized tissue into a 1.5 mL safe-lock tube. Record exact weights for each sample.
If performing extraction on fresh tissue, weigh approximately 20 mg of tissue to account for water content. During weighing of fresh tissue, work quickly and keep all tubes and spatulas on dry ice unless when actively being used to avoid sample thawing. Record weights quickly to avoid accumulation of condensation on the frozen sample.
Add 100 mg of 1 mm glass beads to each sample tube.
Add ice-cold high salt buffer to each sample tube at 200 μL/mg starting dry weight. Maintain cold buffer on ice during decellularization.
If using fresh tissue, add this buffer and all subsequent buffers at 50 μL/mg starting wet weight.
Homogenize samples in buffer using the bead beater (Bullet Blender) for 3 min on power 8.
Vortex samples at maximum power for 20 min at 4 °C.
Centrifuge samples at 18,000× g for 20 min in a 4 °C refrigerated centrifuge.
Remove supernatant, being careful to avoid disturbing the pellet, and reserve as fraction 1.
Keep collected supernatant on ice while completing subsequent decellularization washes.
Repeat steps B3–B7 two additional times, producing fractions 2 and 3. Combine fractions 1–3 to produce the cellular fraction.
During removal of the final supernatant, ensure that all solution is carefully removed from beneath the beads to avoid protein carryover into the next fraction. If the pellet is disturbed during collection, centrifuge again and remove all supernatant without disturbing pellet.
If the solution is difficult to completely remove without disturbing the pellet here or during subsequent fraction collection steps, use a gel-loading pipette tip to remove the last 100–200 μL of solution with greater precision.
Freeze the produced cellular fractions at -80 °C immediately after collecting the third wash and combining it with previous fractions. Store at -80 °C until digestion.
Soluble ECM extraction
Homogenize ECM-enriched pellets in Gnd-HCl solution at 200 μL/mg of the initial tissue dry weight using the Bullet Blender at power 8 for 1 min.
Vortex the homogenate at room temperature (25 °C) overnight at power 5.
After vortexing, centrifuge the homogenate at 18,000× g for 20 min at 4 °C.
Collect the supernatant and reserve it as the soluble ECM (sECM) fraction. Ensure that all solution is carefully removed from below the beads to prevent carryover into the next fraction.
Freeze sECM fractions at -80 °C and store at -80 °C until digestion.
HA buffer treatment
Treat remaining pellets with freshly prepared HA buffer at 200 μL/mg of the initial tissue dry weight.
Homogenize using the Bullet Blender at power 8 for 1 min.
Incubate at 45 °C with shaking on an orbital shaker (1,000 rpm) for 4 h.
Centrifuge at 18,000× g for 20 min at 4 °C.
Collect supernatant and reserve as the insoluble ECM (iECM) fraction.
Freeze iECM immediately alongside previous fractions to prevent further HA cleavage and store fractions at -80 °C until digestion.
Proteolytic digestion using filter-aided sample preparation (FASP)
Aliquot a volume of each extract, which corresponds to roughly 30 μg of protein for each sample, into individual 1.5 mL tubes. Protein quantification is challenging at this point due to high extraction buffer solute concentrations. Aliquot samples according to the following protein concentration estimates (based on typical tissue content) and perform peptide quantification after digestion to accurately assess protein content. Volume estimates for 30 μg of protein based on typical tissue sample:
Cellular fraction: 200 μL
sECM fraction: 100 μL
iECM fraction: 100 μL
If quantification is desired at this step, accurate results can sometimes be obtained using the PierceTM 660 nm protein quantification assay. To use this assay, dilute all buffers at a 1:3 ratio with ddH2O to ensure assay compatibility and then perform the assay according to the manufacturer’s protocol.
Add urea buffer to bring each sample to 250 μL.
Add DTT solution to a final concentration of 10 mM (28 μL of DTT solution for 250 μL of sample).
Incubate samples at 37 °C for 30 min.
While samples are reducing, prepare one 10 kDa MWCO filter for each sample by adding 100 μL of 0.1% FA to each filter.
Centrifuge filters at 14,000× g for 15 min.
Add IAM solution to a final concentration of 22 mM (12.3 μL for 250 μL of sample).
Incubate in the dark at room temperature for 15 min.
Add 100 μL of urea solution to each filter and centrifuge filters at 14,000× g for 15 min.
Load entire reduced and alkylated samples onto equilibrated 10 kDa MWCO filters.
Centrifuge filters at 14,000× g for 15 min.
Discard flowthrough from the collection tube.
Add 200 μL of urea solution to the filter unit and centrifuge at 14,000× g for 20 min.
Repeat wash with 200 μL of urea solution for a second time.
Discard flowthrough from the collection tube.
Add 100 μL of 50 mM ABC to each filter and centrifuge at 14,000× g for 15 min.
Repeat wash with 50 mM ABC two additional times for a total of three washes.
Transfer filters to clean collection tubes.
Prepare 100 μL of digest master mix using Lys-C at a 1:400 enzyme:protein ratio for your number of samples plus one to ensure sufficient digest solution for all samples.
Add 100 μL of digest master mix to each filter.
Incubate filters at 37 °C for 2 h. Ensure high humidity within the incubator by adding a tray of water to evaporate during digestion.
Remove samples from the incubator and add trypsin directly from the commercial stock solution to each sample at a 1:100 enzyme:protein ratio.
Incubate filters at 37 °C overnight for a maximum of 16 h. Ensure high humidity within the incubator by adding a tray of water to evaporate during digestion.
After overnight incubation, centrifuge samples at 14,000× g for 15 min.
Add 75 μL of 0.2% FA to each filter and centrifuge at 14,000× g for 15 min.
Repeat elution with an additional 75 μL of 0.2% FA.
Begin desalting immediately after elution or freeze samples at -80 °C for later processing.
Sample quantification
Prepare an 8-point standard curve (ranging from 1,000 to 0 μg/mL) by diluting the peptide standard included with the PierceTM quantitative colorimetric peptide assay into 0.2% FA. The last point of the standard curve should be a 0.2% FA blank.
Determine the amount of working reagent necessary for your samples (180 μL/sample, including the standard curve) and prepare working reagent by combining the included reagents A, B, and C in a 50:48:2 ratio.
Load 20 μL of each standard curve point and each sample into individual wells of a clear, flat-bottom 96-well plate.
Add 180 μL of working reagent to each sample in the 96-well plate.
Seal the plate with adhesive sealing mat and incubate at 37 °C for 15 min.
Remove sealing mat and load the 96-well plate into Spark microplate reader.
Read the absorbance of the plate at 480 nm. Record values and use the standard curve to determine the concentration of each sample.
Desalting
Load PierceTM C18 spin tips into 2 mL tubes using the provided adapters.
Wash resin using 70 μL of 0.1% FA in 80% ACN. Centrifuge at 1,000× g for 1 min.
Equilibrate resin using 70 μL of 0.1% FA in H2O. Centrifuge at 1,000× g for 1 min.
Discard flowthrough from the 2 mL tube.
Load sample (up to 70 μL) onto C18 spin tips. Use quantification values to determine the volume that corresponds to 8 μg of protein. If more than 70 μL of solution is required to reach 8 µg, load the sample in two successive rounds. Centrifuge at 1,000× g for 1 min.
Wash sample 3× with 70 μL of 0.1% FA. After each buffer addition, centrifuge at 1,000× g for 1 min.
Transfer C18 spin tips to clean 1.5 mL collection tubes using the provided adapters.
Elute peptides with two consecutive washes of 70 μL of 0.1% FA in 80% ACN. After each wash, centrifuge at 1,000× g for 1 min.
Transfer eluted samples to autosampler vials and freeze for 15 min at -80 °C.
Samples can be stored in 0.1% FA, 80% ACN at -80 °C until MS analysis. When ready for analysis, proceed with step G10 using samples directly from the freezer.
Dry samples without autosampler lids in a SpeedVac until 1–3 μL of volume remains.
Rehydrate samples in 9 μL of 0.1% FA for MS analysis. Samples in this state can be stored at 4 °C for up to one week.
LC−MS/MS analysis
Inject 8 μL of each prepared sample at a maximum pressure of 700 bar into a 20 μL sample loop on the EASY-nLC 1200 system, coupled to the Fusion Lumos Tribrid mass spectrometer through a nanoelectrospray interface.
Elute samples from the analytical column at a flow rate of 400 nL/min over a 120-min gradient consisting of 6% buffer B for the first 3 min. For cellular fractions, utilize a subsequent linear gradient from 6% to 42% buffer B over 102 min. For sECM fractions, utilize a linear gradient from 6% to 36% buffer B over 102 min. For iECM fractions, utilize a linear gradient from 6% to 24% buffer B over 102 min. After the linear elution gradient, wash the column by increasing to 95% buffer B and holding for 10 min. Re-equilibrate the column with 6% buffer B for the final 5 min. Gradient schematics and resulting elution profiles for each fraction can be found in Figure 2.
Figure 2. Schematic of elution gradients used for LC-MS/MS analysis of cellular (A), soluble extracellular matrix (sECM) (B), and insoluble extracellular matrix (iECM) (C) fractions from Mus musculus heart tissue. Elution profiles for cellular (D), sECM (E), and iECM (F) fractions using the corresponding gradients.
The Fusion Lumos mass spectrometer is operated using the vendor-provided XCalibur software. Analyze the injected samples using data-dependent acquisition according to the following parameters. Perform full MS1 scans from 375–1600 m/z using the orbitrap at a resolution of 120,000. Select peptides for HCD MS/MS fragmentation (30% collision energy) using the default automatic gain control target and a 35 ms maximum ion injection time. Perform MS/MS detection in the orbitrap with an isolation window of 1/6 m/z. Enable dynamic exclusion after fragmenting a precursor 1 time for a duration of 45 s.
Data processing
Analyze data using Mascot integrated with Proteome Discoverer. Within your processing workflow, first, use Proteome Discoverer to convert the raw files to Mascot generic format (mgf) peaks lists. Search the data against the appropriate UniProt database for your organism using the following parameters: precursor tolerance of ±10 ppm, fragment tolerance of ±20 ppm, and 2 allowed missed cleavages. For cellular and sECM fractions, use fully specific trypsin as the enzyme definition. For the iECM fraction, use a fully-specific HA/Trypsin cleavage definition, defined as cleaving C-terminal of K and R but not before P, as well as C-terminal of N and before G, A, or R. Set fixed modifications as carbamidomethyl (C) and variable modifications as oxidation (M), oxidation (P) (hydroxyproline), Gln->pyro-Glu (N-term Q), deamidation (NQ), and acetyl (protein N-term). Additional modifications commonly identified on fibrillar collagen include hydroxylation (K), galactosylation (K), and galactosyl-glucosylation (K)[16]. Filter results to 1% false discovery rate (FDR) at the peptide level using the Percolator node and at the protein level using the Protein FDR Validator node. Perform label-free quantification (LFQ) using the Precursor Ions Quantifier node, including unique and razor peptides. Example data displays the percentage of total label-free quantification intensity assigned ECM and cellular proteins and summary statistics of protein identifications for each analyzed fraction (Figure 3).
Data can alternatively be processed using the open-source database search software MSFragger through the FragPipe interface [13]. Apply the settings described above to MSFragger database searches with the exception of enzyme cleavage, which should be set to semi-specific trypsin cleavage for ECM fractions. Set maximum variable modifications per peptide to 5. Apply default settings for all fields not described here.
Figure 3. Composition of identified proteins across cell (A), soluble extracellular matrix (sECM) (B), and insoluble extracellular matrix (iECM) (C) fractions from 25 mouse tissues. Values are calculated as a percentage of total LFQ signal assigned to each category across all 25 analyzed tissues from McCabe et al. [11]. Total protein and unique peptide sequence identifications are displayed below the corresponding pie chart for each fraction.
For many datasets, normalization to total LFQ intensity by fraction, so that the total signal in each fraction is equal across samples before fractions are aggregated for analysis, can improve intra-group variability by accounting for inaccuracies in protein load between samples.
Normalization to total LFQ intensity by fraction is not appropriate in cases when there is a significant difference in protein levels between groups.
After normalization (if applying), sum the intensity from the cellular, sECM, and iECM fractions for each sample to generate the total identified intensity across all analyzed fractions. Use this summed data for analysis of total protein composition and the data separated by fraction for analysis of protein solubility/resistance to extraction.
Protein solubility can be measured by calculating the percentage of the total identified signal detected in the iECM fraction for each protein in each sample. Adjust solubility measures based on sample load as a percentage of the total sample by dividing the intensity values for each fraction by the percentage of total extract volume that was loaded onto the mass spectrometer.
For ECM-focused analysis, annotate ECM and matrisome-associated proteins within the Excel output files using MatrisomeDB [15].
Validation of protocol
This protocol has been used and validated in the following research article:
McCabe et al. [11]. Mass Spectrometry-Based Atlas of Extracellular Matrix Proteins across 25 Mouse Organs. Journal of Proteome Research.
In the article above, the present protocol was employed for ECM-focused proteomics analysis of 25 mouse organs. All data within the article is derived from this protocol. For 12 of the analyzed tissues (reported within the article), three replicate mice (male, female, and pregnant female) were analyzed, and statistical testing was performed using multiple comparison–adjusted t-tests and ANOVA. For the remaining 13 tissues, samples were derived from a single pregnant female mouse harvested when the embryo was in the E12−16 embryonic stage.
General notes and troubleshooting
Troubleshooting
Issue Cause Solution
Layer forms at the top of decellularization extract and will not pellet. Tissue sample contains lipids that do not pellet or become solubilized by extraction buffer. Perform acetone precipitation on samples before extraction to remove lipids according to the protocol from Simpson and Beynon [17].
Pellet is collected during the removal of supernatant during any extraction step. Pellet was not fully compacted by centrifugation or was disturbed during pipetting. Return extract to tube with pellet, repeat centrifugation, and repeat supernatant removal, taking care to avoid the pellet. A gel-loading pipette tip can be used for additional precision during supernatant removal if necessary.
FASP filter clogs during sample loading, sample will not flow through. Too much protein or particulate from extract fraction loaded onto filter. Centrifuge samples at 18,000× g for 10 min before loading and pipette from the top of the solution to avoid any pelleted particulate. If this does not fix the problem, repeat with reduced extract volume.
Post-digest quantification does not provide reliable values or indicates that digested protein is insufficient for analysis.
1. Tissue sample had less protein than expected, resulting in insufficient digested protein.
2. Digestion was unsuccessful/incomplete and undigested proteins remained on the FASP filter after elution.
3. Protein is distributed differently across the three extracted fractions than the estimates provided for a typical tissue, causing unsuccessful quantification for one fraction due to low protein content. For example, bone without marrow will have greater protein content in the iECM fraction and less in the cellular fraction than a typical tissue.
1. Utilize post-digest quantification to revise amounts of extract loaded for digestion. Increase loaded volume to contain approximately 30 µg of protein.
2. Repeat digestion, ensuring that enzymes are active and that all wash buffer is fully eluted from the FASP filter before adding digest buffer. If desired, SDS-PAGE can be utilized to assess digestion efficacy.
3. Repeat FASP digest of fraction with low protein content using 2–3× the previous digest volume. Adjust content of other fractions for C18 cleanup based on quantification values.
Lower identification than expected after database searching.
1. Incomplete digestion—determined by high content (>5%) of trypsin missed-cleavage identifications.
2. Carbamylation of proteins from urea adducts.
1. Repeat digestion of extracted fractions, increasing enzyme:protein ratios to 1:200 and 1:75 for Lys-C and trypsin, respectively. If necessary, increase digestion time for Lys-C to 4 h before adding trypsin for overnight incubation.
2. Make sure that urea solution is freshly made and is not heated. Additionally, urea solutions can be passed over mixed mode resins for removal of impurities. Alternatively, switch urea solution to Gnd-HCl solution during FASP digestion. If carbamylation is a suspected issue but sample re-preparation is not possible, carbamylation (+43.005814, Unimod, catalog number: 5) can be added as a variable modification to database searches to identify carbamylated peptides.
Acknowledgments
This work was supported by the NIH (Grant Nos. R33CA183685, RM1GM131968, P01HL152961, and R01HL146519) and the University of Colorado Cancer Center Support Grant (P30CA046934). This protocol was previously described and validated in McCabe et al., 2023 (Journal of Proteome Research)[11].
Competing interests
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
Ethical considerations
Mouse tissues utilized in McCabe et al. 2023 [11], where this protocol was initially described and validated, were acquired from the Jackson Laboratory and approved by the Institutional Animal Care and Use Committee (IACUC) and Association for Assessment and Accreditation of Laboratory Animal Care, International (AAALAC) approvals for mouse research.
References
Mecham, R. P. (2012). Overview of Extracellular Matrix. Curr Protoc Cell Biol. 57(1): ecb1001s57. https://doi.org/10.1002/0471143030.cb1001s57
Hynes, R. O. (2009). The Extracellular Matrix: Not Just Pretty Fibrils. Science (1979). 326(5957): 1216–1219. https://doi.org/10.1126/science.1176009
Harvey, A., Montezano, A. C., Lopes, R. A., Rios, F. and Touyz, R. M. (2016). Vascular Fibrosis in Aging and Hypertension: Molecular Mechanisms and Clinical Implications. Can J Cardiol. 32(5): 659–668. https://doi.org/10.1016/j.cjca.2016.02.070
Wynn, T. (2007). Cellular and molecular mechanisms of fibrosis. J Pathol. 214(2): 199–210. https://doi.org/10.1002/path.2277
Barallobre-Barreiro, J., Didangelos, A., Schoendube, F. A., Drozdov, I., Yin, X., Fernández-Caggiano, M., Willeit, P., Puntmann, V. O., Aldama-López, G., Shah, A. M., et al. (2012). Proteomics Analysis of Cardiac Extracellular Matrix Remodeling in a Porcine Model of Ischemia/Reperfusion Injury. Circulation. 125(6): 789–802. https://doi.org/10.1161/circulationaha.111.056952
Biernacka, A., Frangogiannis, N. G., Cardiology, D., Einstein, A. and Ny, B. (2012). Aging and Cardiac Fibrosis. Aging Dis. 2(2): 158–173.
Barrett, A. S., Maller, O., Pickup, M. W., Weaver, V. M. and Hansen, K. C. (2018). Compartment resolved proteomics reveals a dynamic matrisome in a biomechanically driven model of pancreatic ductal adenocarcinoma. J Immunol Regener Med. 1: 67–75. https://doi.org/10.1016/j.regen.2018.03.002
Stoilov, I., Starcher, B. C., Mecham, R. P. and Broekelmann, T. J. (2018). Measurement of elastin, collagen, and total protein levels in tissues. Methods Cell Biol. 143: 133–146. https://doi.org/10.1016/bs.mcb.2017.08.008
Rickelt, S. and Hynes, R. O. (2018). Antibodies and methods for immunohistochemistry of extracellular matrix proteins. Matrix Biol. 71–72: 10–27. https://doi.org/10.1016/j.matbio.2018.04.011
Goddard, E. T., Hill, R. C., Barrett, A., Betts, C., Guo, Q., Maller, O., Borges, V. F., Hansen, K. C. and Schedin, P. (2016). Quantitative extracellular matrix proteomics to study mammary and liver tissue microenvironments. Int J Biochem Cell Biol. 81: 223–232. https://doi.org/10.1016/j.biocel.2016.10.014
McCabe, M. C., Saviola, A. J. and Hansen, K. C. (2023). Mass Spectrometry-Based Atlas of Extracellular Matrix Proteins across 25 Mouse Organs. J Proteome Res. 22(3): 790–801. https://doi.org/10.1021/acs.jproteome.2c00526
McCabe, M. C., Schmitt, L. R., Hill, R. C., Dzieciatkowska, M., Maslanka, M., Daamen, W. F., van Kuppevelt, T. H., Hof, D. J. and Hansen, K. C. (2021). Evaluation and Refinement of Sample Preparation Methods for Extracellular Matrix Proteome Coverage. Mol Cell Proteomics. 20: 100079. https://doi.org/10.1016/j.mcpro.2021.100079
Kong, A. T., Leprevost, F. V., Avtonomov, D. M., Mellacheruvu, D. and Nesvizhskii, A. I. (2017). MSFragger: ultrafast and comprehensive peptide identification in mass spectrometry–based proteomics. Nat Methods. 14(5): 513–520. https://doi.org/10.1038/nmeth.4256
Mellacheruvu, D., Wright, Z., Couzens, A. L., Lambert, J. P., St-Denis, N. A., Li, T., Miteva, Y. V., Hauri, S., Sardiu, M. E., Low, T. Y., et al. (2013). The CRAPome: a contaminant repository for affinity purification–mass spectrometry data. Nat Methods. 10(8): 730–736. https://doi.org/10.1038/nmeth.2557
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Article Information
Publication history
Received: May 1, 2024
Accepted: Sep 23, 2024
Available online: Oct 22, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Quantitative Measurement of the Kinase Activity of Wildtype ALPK1 and Disease-Causing ALPK1 Mutants Using Cell-Free Radiometric Phosphorylation Assays
TS Tom Snelling
Published: Vol 14, Iss 22, Nov 20, 2024
DOI: 10.21769/BioProtoc.5124 Views: 258
Reviewed by: Ralph Thomas BoettcherMasashi AsaiMartin V Kolev
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Original Research Article:
The authors used this protocol in bioRxiv Sep 2024
Abstract
ALPK1 is an atypical protein kinase that is activated during bacterial infection by ADP-heptose and phosphorylates TIFA to activate a cell signaling pathway. In contrast, specific mutations in ALPK1 allow it to also be activated by endogenous human nucleotide sugars such as UDP-mannose, leading to the phosphorylation of TIFA in the absence of infection. This protocol describes a quantitative, cell-free phosphorylation assay that can directly measure the catalytic activity of wildtype and disease-causing ALPK1 in the presence of different nucleotide sugars. In this method, overexpressed ALPK1 is first immunoprecipitated from the extracts of ALPK1 knockout HEK-Blue cells transfected with plasmids encoding either FLAG-tagged wildtype or mutant ALPK1, and then subjected to a radioactive phosphorylation assay in which the phosphorylation of purified GST-tagged TIFA by ALPK1 is quantified by measuring the incorporation of radioactivity derived from radiolabeled ATP.
Key features
• Quantitative measurement of protein kinase activity of wildtype and mutant ALPK1 in the presence or absence of different nucleotide sugars such as ADP-heptose and UDP-mannose.
• Cell-free experimental setup overcoming the challenge of distinguishing constitutive activity and activation by endogenous mammalian metabolites in cell-based assays.
• Requires approximately 50 µg of cell extract protein/reaction, allowing up to 150 assays to be performed from an extract prepared from a single 15 cm dish of transfected cells.
Keywords: ALPK1 Nucleotide sugar ADP-heptose ROSAH Spiradenoma Phosphorylation Protein kinase
Background
Alpha-protein kinase 1 (ALPK1) is an atypical protein kinase that is activated by the binding of the bacterial metabolite ADP-heptose to its N-terminal domain [1]. This allows ALPK1 to phosphorylate TIFA (TRAF-interacting protein with forkhead-associated domain), which in turn triggers its polymerization into TIFAsomes that initiate signaling events that lead to the activation of transcription factors such as NF-κB and AP-1 [1,2] (see [3] for a schematic overview of the ADP-heptose signaling pathway).
Mutations in ALPK1 are the cause of at least two human diseases. The Thr237Met, Tyr254Cys, and Ser277Phe mutations cause ROSAH syndrome (retinal dystrophy, optic nerve edema, splenomegaly, anhidrosis, and migraine headache), which is an autosomal dominant genetic disorder [4–6], whereas the ALPK1[V1092A] mutation is a driver of spiradenoma, a rare type of hair follicle tumor that can transform into a malignant form called spiradenocarcinoma, which is invariably fatal [7]. These mutations allow ALPK1 to be activated by endogenous mammalian nucleotide sugars, such as UDP-mannose and ADP-ribose, in addition to bacterial ADP-heptose, leading to pathological signaling in the absence of bacterial infection [3,6].
This protocol describes a cell-free phosphorylation assay that can be used to quantify the protein kinase activity of wildtype and mutant forms of ALPK1 in the presence or absence of different nucleotide sugars. Existing methods for measuring the activity of ALPK1 mutants have relied on measuring their activity within cells, but the complexity of the intracellular environment makes it challenging to identify specific activators. Furthermore, these cell-based methods rely on indirect measurements of ALPK1 activity after many hours, such as the activation of transcription factors or the secretion of cytokines or chemokines. In contrast, this phosphorylation assay has the advantage of enabling rapid and quantitative measurements of ALPK1 activity by directly measuring its ability to phosphorylate TIFA in a cell-free system. Additionally, this assay is ideal for validating potential ALPK1 inhibitors by studying their potency and selectivity for targeting wildtype or mutant forms of this protein kinase.
Materials and reagents
Biological materials
Cells should be cultured by incubation at 37 °C with 5% CO2 and tested regularly for mycoplasma using a MycoAlert Mycoplasma Detection Kit (Lonza, catalog number: LT07-318). The cells should be passaged once confluent at a ratio of 1:10 (v/v), and not used beyond 30 passages.
ALPK1 knockout (KO) HEK-Blue cells (InvivoGen, #hkb-koalpk)
General reagents
Storage conditions are given in parentheses unless it is room temperature.
ADP-L-heptose triethylammonium salt (InvivoGen, catalog number: tlrl-adph-l) (-80 °C)
Note: Resuspend 250 µg in 3,470 µL of reaction buffer (see Recipes) to generate a 0.1 mM stock.
UDP-α-D-mannose triethylammonium salt (synthesized in-house, available upon request) (-80 °C)
Note: Resuspend 2 mg in 3,280 µL of reaction buffer to generate a 1 mM stock.
ADP-D-ribose sodium salt (Sigma-Aldrich, catalog number: C7344) (-80 °C)
Note: Resuspend 2 mg in 3,580 µL of reaction buffer to generate a 1 mM stock.
3,000 Ci/mmol [γ32P]ATP (Revvity, catalog number: BLU002A001MC) (-20 °C)
Caution: [γ32P]ATP must be handled according to radioactivity safety regulations. Exposure to radiation must be minimized by working behind a plexiglass shield and storing samples in plexiglass boxes.
Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, catalog number: 11960-085) (4 °C)
200 mM L-Glutamine (Gibco, catalog number: 25030024) (-20 °C)
Penicillin-Streptomycin 100× stock (Gibco, catalog number: 15140122) (-20 °C)
OptiMEM I reduced serum medium (Gibco, catalog number: 31985062) (4 °C)
Lipofectamine 2000 (Thermo Fisher, catalog number: 11668019) (4 °C)
Trypan Blue solution (Gibco, catalog number: 11538886)
Anti-FLAG M2 affinity resin (Millipore, catalog number: A2220)
Phosphate-buffered saline (PBS) (Gibco, catalog number: 10010023) (4 °C)
10 mM ATP (Thermo Fisher, catalog number: PV3227) (-20 °C)
Note: This commercial stock solution does not contain magnesium ions.
5 M NaCl (Sigma-Aldrich, catalog number: S6546)
1 M DTT (Thermo Fisher, catalog number: P2325) (-20 °C)
Note: Prepare 0.1 mL single-use aliquots.
1 M Tris-HCl (pH 7.5) (Thermo Fisher, catalog number: 15567027)
0.5 M EGTA (pH 8.0) (Thermo Fisher, catalog number: J60767.AD)
0.5 M EDTA (pH 8.0) (Sigma-Aldrich, catalog number: 03690)
10% (v/v) Triton X-100 (Sigma-Aldrich, catalog number: 93443) (4 °C)
Fetal bovine serum (FBS) (Sigma-Aldrich, catalog number: F7524) (-20 °C)
Sucrose powder (VWR, catalog number: 27480.360)
InstantBlue protein stain (Abcam, catalog number: ab119211) (4 °C)
MOPS SDS running buffer 20× stock (Formedium, catalog number: SDS5000)
NuPAGE LDS sample buffer 4× stock (Thermo Fisher, catalog number: NP0007)
Precision Plus protein standards (Bio-Rad, catalog number: 1610374) (-20 °C)
2-Mercaptoethanol (Sigma-Aldrich, catalog number: M6250)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: 810533) (4 °C)
Bradford Protein Assay Kit (Thermo Fisher, catalog number: 23200) (4 °C)
1 M Magnesium acetate (MgAc) (Sigma-Aldrich, catalog number: 63052)
Trypsin-EDTA solution (Gibco, catalog number: 25200056) (4 °C)
Complete EDTA-free protease inhibitor cocktail (Roche, catalog number: 11873580001) (4 °C)
Purified proteins
TIFA was purified from bacteria as a GST fusion and is available upon request via the MRC PPU Reagents and Services website (https://mrcppureagents.dundee.ac.uk).
GST-TIFA[2-184] (MRC PPU Reagents and Services, catalog number: DU4241) (-80 °C)
Note: Dialyzed against reaction buffer and stored at 1 mg/mL in single-use aliquots.
Plasmids
These plasmids encode FLAG-ALPK1 or FLAG alone (empty vector control) under a CMV promoter and were purified using NucleoBond Xtra Midi Endotoxin-Free kits (Macherey-Nagel, catalog number: 740420). The yield was ~0.5 mg, resuspended to 0.5 mg/mL in endotoxin-free water. They are stored at -20 °C.
pcDNA5-FRT/TO-FLAG-ALPK1 (MRC PPU Reagents and Services, catalog number: DU65668)
pcDNA5-FRT-TO-FLAG-ALPK1[S277F] (MRC PPU Reagents and Services, catalog number: DU71952)
pcDNA5-FRT-TO-FLAG (MRC PPU Reagents and Services, catalog number: DU41457)
Solutions
Culture media (see Recipes)
Lysis buffer (see Recipes)
Wash buffer (see Recipes)
Salt wash buffer (see Recipes)
Reaction buffer (see Recipes)
Radioactive ATP solution (see Recipes)
Recipes
Culture media (1 bottle)
Reagent Final concentration Amount to add
DMEM Not applicable 500 mL
FBS 10% (v/v) 50 mL
20 mM L-glutamine 2 mM 5.6 mL
Penicillin-Streptomycin 100× 1× 5.6 mL
Store at 4 °C.
Lysis buffer (50 mL)
Reagent Final concentration Amount to add
1M Tris-HCl (pH 7.5) 50 mM 2.5 mL
Sucrose powder 270 mM 4.6 g
10% (v/v) Triton X-100 1% (v/v) 5 mL
0.5 M EDTA (pH 8.0) 1 mM 100 µL
0.5 M EGTA (pH 8.0) 1 mM 100 µL
1 M DTT 2 mM 100 µL
Protease inhibitor cocktail 1× 1 tablet
Water Not applicable 42.2 mL
Use immediately.
Wash buffer (1 L)
Reagent Final concentration Amount to add
1 M Tris-HCl (pH 7.5) 50 mM 50 mL
10% (v/v) Triton X-100 0.1% (v/v) 10 mL
1 M DTT 2 mM 2 mL
Water Not applicable 938 mL
Store at 4 °C. Use within 1 month.
Salt wash buffer (1 L)
Reagent Final concentration Amount to add
1 M Tris-HCl (pH 7.5) 50 mM 50 mL
10% (v/v) Triton X-100 0.1% (v/v) 10 mL
5 M NaCl 0.5 M 100 mL
1 M DTT 2 mM 2 mL
Water Not applicable 838 mL
Store at 4 °C. Use within 1 month.
Reaction buffer (1 L)
Reagent Final concentration Amount to add
1 M Tris-HCl (pH 7.5) 50 mM 50 mL
0.5 M EGTA (pH 8.0) 1 mM 2 mL
1 M DTT 2 mM 2 mL
1 M MgAc 10 mM 10 mL
Water Not applicable 937.6 mL
Store at 4 °C. Use within 1 month.
Radioactive ATP solution (150 µL) (sufficient for 60 kinase reactions)
Reagent Final concentration Amount to add
Undiluted [γ32P]ATP Not applicable 5–30 µL*
10 mM ATP 1 mM 15 µL
Reaction buffer (recipe 5) Not applicable 105–130 µL
Use immediately.
*The amount of undiluted [γ32P]ATP added to the radioactive ATP solution should be calculated based on the activity reference date to achieve a specific activity of approximately 500 cpm per pmol of ATP for each experiment.
Laboratory supplies
15 cm Nunc cell culture dishes (Thermo Fisher, catalog number: 168381)
15 and 50 mL conical centrifuge tubes (Greiner, catalog numbers: 188271 and 227261)
250 mL and 1 L Duran bottles (Thermo Fisher, catalog numbers: FB-800-250 and FB-800-1000)
Serological pipettes (Thermo Fisher, catalog number: 10710810)
Safe-lock 1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 30123611)
Note: Safe-lock tubes are highly recommended to avoid losing the sample during end-to-end rotation.
Cellometer counting chamber (Nexcelom, catalog number: 11522186)
Spin-X 0.22 μm columns (Costar, catalog number: 8161)
NuPAGE Bis-Tris 4%–12% 20-well gels (Thermo Fisher, catalog number: WG1402BOX)
80-well cooling chamber for 1.5 mL tubes (Diversified Biotech, catalog number: CHAM-8000)
Swann-Morton number 22 disposable scalpels (Scientific Laboratory Supplies, catalog number: INS4767)
Equipment
The equipment mentioned below are standard laboratory items, and alternatives can be used in all cases.
Cellometer Auto 2000 (Nexcelom Bioscience)
Pipetman 4-Pipette Kit (Gilson, catalog number: F167360)
Stripettor Ultra Pipet Controller (Corning, catalog number: 4099)
Thermomixer (Eppendorf, model: Comfort 5355)
Cell culture incubator (Binder, model: CB150)
Cell culture hood (Conditioned Air Solutions, model: BioMAT 2-SF)
Liquid scintillation counter (Revvity, model: Tri-Carb 4910 TR)
Geiger counter
Plexiglas benchtop shield
Allegra X-12 benchtop centrifuge (Beckman Coulter, catalog number: 392474)
Benchtop microcentrifuge (Fisherbrand, model: Micro STAR 17)
Refrigerated benchtop microcentrifuge (Fisherbrand, model: Accuspin Micro 17)
Dry block heater (Grant, model: QBT2)
NuPAGE XCell SureLock Midi-Cell system (Invitrogen)
Plate reader (BioTek, model: Epoch)
Software and datasets
Image Lab (BioRad, Version 6.0.1)
Procedure
Before starting this procedure, two confluent 15 cm dishes of ALPK1 KO HEK-Blue cells, no higher than passage 30, are required. This protocol describes how to compare wildtype ALPK1 and the ALPK1[S277F] mutant, but the experiment can also be scaled up to include additional conditions as required.
Transient expression of FLAG-tagged ALPK1 constructs in ALPK1 KO HEK-Blue cells (Days 1–2)
All steps in section A should be performed in a cell culture hood and the cells should be cultured at 37 °C with 5% CO2.
Plate 3 × 15 cm dishes with 15 million ALPK1 KO HEK-Blue cells each, which will be transfected with empty vector (dish 1) or plasmid encoding FLAG-ALPK1 (dish 2) or FLAG-ALPK1[S277F] (dish 3):
In the afternoon, aspirate the culture media from two confluent 15 cm dishes of cells and replace with 10 mL of PBS using a serological pipette.
Aspirate the PBS and add 3 mL of trypsin-EDTA solution to each dish. Return the dishes to the incubator until the cells have detached, which should take 2–3 min for this cell line.
Add 15 mL of culture media to each plate and pipette up and down using a serological pipette until a single-cell suspension has been produced. Combine the cell suspensions into a 50 mL canonical centrifuge tube.
Remove 20 µL of the cell suspension and dilute with 80 µL of culture media. Mix 20 µL of the diluted cell suspension with 20 µL of trypan blue solution and count the number of cells using standard methods, ensuring that the cell viability is at least 90%.
Plate 15 million ALPK1 KO HEK-Blue cells into each of the 3 × 15 cm dishes and add media up to a total volume of 20 mL. Ensure that cells are evenly distributed by moving plates in a figure-of-8 motion prior to returning them to the incubator for 18 h.
Note: Do not change to antibiotic-free culture media for transfection in this protocol.
Transfection of ALPK1 KO HEK-Blue cells with FLAG-tagged ALPK1 constructs:
After 18 h, confirm that the confluency of the ALPK1 KO HEK-Blue cells is approximately 90%.
For each plate to be transfected, add 150 µL of lipofectamine 2000 to 600 µL of OptiMEM in a 1.5 mL microcentrifuge tube (i.e., prepare 3 tubes in this example). Invert five times to mix. In this example, the plates will be transfected with either empty vector or plasmid encoding either FLAG-ALPK1 or FLAG-ALPK1[S277F] (i.e., 3 dishes).
Dilute 60 μg of each plasmid in 600 µL of OptiMEM in 1.5 microcentrifuge tubes. Invert five times to mix.
Add the diluted lipofectamine 2000 to each diluted plasmid. Invert five times to mix and incubate for 10 min at room temperature.
Add each solution dropwise to the relevant dish of cells and return them to the incubator.
After 4 h, carefully aspirate the culture media and add 15 mL of fresh culture media by pipetting slowly against the side of the plate to minimize cell detachment. This step removes the DNA-lipid complexes, which is observed to minimize toxicity. Return the plates to the incubator for 20 h.
Preparation and normalization of cell extracts from transfected ALPK1 KO HEK-Blue cells (Day 3)
Preparation of cell extracts from HEK-Blue cells transfected with different plasmids:
Note: Cell extracts must not be snap-frozen until the end of Section B, since additional freeze-thaw cycles have been observed to lead to a significant reduction in the activity of ALPK1 from cell extracts.
Twenty-four hours post-transfection, use a 15 mL serological pipette to pipette the culture media up and down until the cells are detached (trypsinization is not required). Transfer the cell suspensions into 15 mL canonical centrifuge tubes.
Centrifuge the tubes at 800× g for 5 min at room temperature to pellet the cells. Aspirate the supernatant, add 15 mL of room-temperature PBS to the pellet (do not resuspend), and repeat the centrifugation and aspiration steps for a total of two PBS washes.
After the final PBS wash, ensure that all residual PBS is aspirated and place the cell pellets on ice. All subsequent steps are performed on ice, outside of a sterile cell culture hood, ensure that all residual PBS is aspirated and place the cell pellets on ice.
Add 1 mL of ice-cold lysis buffer to each cell pellet, pipetting up and down until a homogenous suspension is formed.
Transfer each cell lysate to a pre-chilled 1.5 microcentrifuge tube on ice, centrifuge at 18,000× g for 20 min at 4 °C, and transfer each supernatant (cell extract) to a new 1.5 mL microcentrifuge tube on ice.
Use the Bradford protein assay kit to determine the protein concentration in each cell extract. Transfer 5 µL of each cell extract to a 1.5 mL microcentrifuge tube and dilute 1:5 (v/v) by addition of 20 µL of water. Transfer 5 µL of each diluted extract in triplicate to a 96-well plate and 5 µL in triplicate of protein standards containing 2.0, 1.0, 0.75, 0.5, 0.25, 0.125, and 0.0625 mg/mL of BSA in water. Add 195 µL of Bradford reagent and measure the absorbance at 595 nm using a microplate reader. Calculate the protein concentration for each cell extract by interpolation of the BSA standard curve, considering the dilution factor.
Dilute the cell extracts to a final concentration of 2 mg/mL protein using ice-cold lysis buffer in a 15 mL conical centrifuge tube on ice. Leave the samples at 4 °C while performing the steps described below.
Normalization of cell extracts based on relative ALPK1 expression levels:
An aliquot of each cell extract (0.2 mg of protein) will be immunoprecipitated using 15 µL of resin to determine the relative levels of each ALPK1 protein. Prepare 55 µL of resin as detailed below, sufficient for three samples plus 20% extra.
Add 110 µL of anti-FLAG M2 affinity resin slurry (50% resin by volume, i.e., 55 µL of resin) to a 1.5 mL microcentrifuge tube on ice using a pipette tip where the narrow end has been trimmed using a scalpel.
Centrifuge the slurry at 2,000× g for 1 min at 4 °C to pellet the resin. Aspirate the supernatant and add 1 mL of ice-cold lysis buffer. Ensure that the resin is thoroughly resuspended by inverting five times. Repeat the centrifugation and wash steps twice.
After the final wash step, resuspend the beads in 945 µL of ice-cold lysis buffer and invert five times to ensure that the resin is thoroughly resuspended. Use a trimmed pipette tip to add 270 µL of this slurry to 3 × 1.5 mL microcentrifuge tubes (i.e., 15 µL of resin per tube).
Add 100 µL of ice-cold cell extract to each of the tubes (i.e., 0.2 mg). Incubate at 4 °C for 1 h on a rotating wheel to immunoprecipitate the FLAG-tagged ALPK1 from each cell extract.
Centrifuge the samples at 2,000× g for 1 min at 4 °C to pellet the resin, carefully aspirate the supernatant, and resuspend the resin in 1 mL of ice-cold salt wash buffer.
Repeat the centrifugation and wash steps a further two times with ice-cold salt wash buffer (for a total of three washes with this buffer), followed twice with ice-cold wash buffer to remove NaCl prior to SDS-PAGE.
After the final wash, carefully aspirate all residual wash buffer, such that the resin is dry. All following steps in this section are performed at room temperature.
Resuspend the resin in 20 µL of LDS sample buffer 1× with 2.5% (v/v) 2-mercapthetanol and heat for 5 min at 75 °C. This solution is prepared by combining 20 µL of LDS sample buffer 4×, 58 µL of water, and 2 µL of 2-Mercaptoethanol.
Centrifuge the samples at 13,000× g for 30 s and transfer supernatants to Spin-X columns to remove resin.
Centrifuge the Spin-X columns at 13,000× g for 30 s and analyze 10 µL of the eluent (i.e., 50%) by SDS-PAGE alongside 5 µL of the protein ladder (Precision Plus Protein Standards) (see Figure 1A).
Stain the SDS-PAGE gel for 30 min with InstantBlue and destain for 1 h in water with regular changes.
Image the SDS-PAGE gel and quantify the intensity of each FLAG-ALPK1 mutant relative to the WT, for example using Image Lab software.
Calculate the dilution required to normalize cell extracts based on the expression of each ALPK1 in each lysate. In other words, dilute each cell extract to match the concentration of ALPK1 present in the sample containing the lowest amount of ALPK1 (i.e., if the concentration of ALPK1 in a given sample is twice as high as that of the lowest, dilute it in an equal volume). Perform the dilutions using lysate prepared from cells transfected with empty vector plasmid, such that the final protein concentration of each lysate remains unchanged.
In this example, FLAG-ALPK1 and FLAG-ALPK1[S277F] were expressed successfully, leading to a band at approximately 140 kDa that was absent from the empty vector control (Figure 1A). These two forms of ALPK1 were expressed equally well, and normalization was therefore not needed.
Figure 1. Phosphorylation of GST-TIFA by FLAG-ALPK1 and its mutants. (A) SDS-PAGE analysis showing the relative levels of ALPK1 in FLAG immunoprecipitations of the extracts prepared from ALPK1 KO HEK-Blue cells transfected with empty vector or plasmid encoding either FLAG-tagged WT ALPK1 or ALPK1[S277F]. The band intensities were quantified using ImageLab and were found to be identical in this experiment. (B) Stained SDS-PAGE gel from a time course phosphorylation experiment in the presence and absence of ADP-heptose (ADPH). The dashed box indicates the region corresponding to GST-TIFA, from which bands were excised for scintillation counting. (C) Quantification of the incorporation of radioactivity into GST-TIFA from the time course experiment shown in (B). (D) Phosphorylation assays were performed for 20 min with either WT ALPK1 or the ALPK1[S277F] mutant in the presence or absence of 10 µM ADP-heptose (ADPH), 100 µM UDP-mannose (UDPM), or 100 µM ADP-ribose (ADPR). For additional details, please see the main text. (C, D) The points in (C) and the bar heights in (D) represent the mean values, and the error bars indicate plus and minus one standard error of the mean.
Prepare 0.5 mL aliquots of each cell extract on ice, which corresponds to 0.5 mg of WT FLAG-ALPK1 lysate and normalized amounts of lysate containing equivalent amounts of the respective mutant.
There will typically be 15 aliquots of WT FLAG-ALPK1 from a 15 cm dish of transfected cells, which is sufficient for 150 kinase reactions. Snap-freeze these single-use aliquots and store them at -80 °C until use.
Note: It is recommended to use these aliquots within 1 month and not re-freeze them once thawed, since the ALPK1 activity in the cell extract might be reduced or lost.
Determination of linear rate conditions for WT ALPK1 (Days 4–5)
Kinase reactions must be performed under linear rate conditions, and it is therefore essential to perform a time course experiment with WT FLAG-ALPK1 for 5, 10, 20, and 30 min in both the presence and absence of ADP-heptose each time that cell extracts are prepared. This allows the optimal time at which to terminate the reaction to be determined for the experiments in Section D.
Perform time course phosphorylation assays for WT FLAG-ALPK1:
The steps below describe how to assay WT FLAG-ALPK1 with and without ADP-heptose for 5, 10, 20, and 30 min, each time point performed in duplicate. A 0-min point is also included, performed without ADP-heptose (in singlet), plus an extra condition (in singlet) where GST-TIFA is not added. This is a total of 18 reactions (see Figure 1B for a summary of the 18 recommended conditions).
Thaw two aliquots of WT FLAG-ALPK1 lysate on ice and combine them, sufficient for 20 reactions.
Add 600 µL of anti-FLAG M2 affinity resin slurry (50% resin by volume) to a 1.5 mL microcentrifuge tube on ice, which is sufficient for 18 samples plus an excess of 10% (15 µL of resin per reaction).
Centrifuge the slurry at 2,000× g for 1 min at 4 °C to pellet the resin. Aspirate the supernatant and resuspend the resin in 1 mL of ice-cold lysis buffer. Repeat the centrifugation and wash steps twice. After the final wash step, resuspend the dry resin in the lysate (1 mg) expressing WT FLAG-ALPK1. Incubate at 4 °C for 1 h on a rotating wheel.
Wash the sample three times with ice-cold salt wash buffer, twice with ice-cold wash buffer, and once with ice-cold reaction buffer.
Resuspend the 300 µL of dry resin in a total of 350 µL of ice-cold reaction buffer and aliquot 32.5 µL of this slurry into 18 individual 1.5 mL microcentrifuge tubes on ice, ensuring that the pipette tip is pushed to the bottom of the tubes to prevent the resin from drying out (i.e., do not pipette the resin onto the sides of the tubes). Each tube should now contain 15 µL of resin resuspended in 17.5 µL of reaction buffer, on ice.
Add 2.5 µL of 1 mg/mL GST-TIFA (i.e., 2.5 µg, which is 2.1 µM) to each tube on ice (except the no GST-TIFA sample, where reaction buffer should be added instead), followed by either 2.5 µL of reaction buffer or 2.5 µL of 100 µM ADP-heptose in reaction buffer, on ice.
Add 2.5 µL of the radioactive ATP solution (see Recipes) to the samples at 20 s intervals and put them onto a prewarmed thermomixer at 30 °C at 1,300 rpm, except for the 0 min timepoints, which should be terminated as described below, prior to the addition of the radioactive ATP solution.
Note: The radioactive solution must be handled according to safety regulations.
To terminate reactions after the specified lengths of time have passed (i.e., 5, 10, 20, and 30 min), add 8.3 µL of LDS sample buffer 4× containing 10% (v/v) 2-mercaptethanol, heat for 5 min at 75 °C, and remove the resin using Spin-X columns.
Note: Check the centrifuge and heat block for radioactive contamination.
Analyze half of the supernatant (i.e., 16.7 µL) by SDS-PAGE.
Note: SDS-PAGE should be stopped before the dye front enters the running buffer, as this will minimize the generation of aqueous radioactive waste. The dye front should be excised from the gel and discarded as solid radioactive waste.
Stain with InstantBlue for 1 h and destain in water for 24 h with frequent changes.
Note: Discard the destaining water as aqueous radioactive waste.
Figure 1B shows the appearance of the stained gel but has been cropped above the 75 kDa marker, since the amount of FLAG-ALPK1 in the reaction is below the detection limit of the stain and therefore not visible.
Quantify the incorporation of radioactivity into GST-TIFA:
Wash the SDS-PAGE gel five times in water for 5 min each to minimize background radiation.
Note: Discard the destaining water as aqueous radioactive waste.
Transfer the SDS-PAGE gel to an A4 plastic pocket and dab with filter paper to remove excess water.
Cut out individual bands corresponding to GST-TIFA (Figure 1B, dashed box) using a scalpel and transfer to 1.5 mL microcentrifuge tubes. Centrifuge the tubes containing the excised pieces of SDS-PAGE gel samples at 13,000× g for 1 min to bring the gel pieces to the bottom of each tube.
Count each sample using a scintillation counter for 2 min per sample and plot the resulting data (Figure 1C).
Count triplicate 1 µL aliquots of the radioactive ATP solution. Since the radioactive ATP solution contains 1 mM ATP (i.e., 1 nmol ATP per µL), the radioactivity of a 1 µL aliquot is the specific radioactivity in cpm per nmol of ATP. Knowing the specific radioactivity is necessary for comparing the results of different experiments and can be used to calculate the stoichiometry of phosphorylation (not shown).
In this example, the 20 min timepoint was chosen for subsequent experiments because it was the longest timepoint that was still within the linear range of the assay (see Figure 1C).
Measurement of the activity of ALPK1 mutants in the presence of nucleotide sugars (Days 6–7)
Perform endpoint phosphorylation assays with different nucleotide sugars:
Thaw an aliquot of each cell extract on ice, sufficient for 10 reactions. Each lysate will be used for assays in the presence of buffer, ADP-heptose, UDP-mannose or ADP-ribose, each in duplicate, requiring eight reactions.
Since there are two aliquots in total (WT and S277F), add 0.6 mL of anti-FLAG M2 affinity resin slurry (10% excess) to a 1.5 mL microcentrifuge tube on ice and centrifuge the slurry at 2,000× g for 1 min at 4 °C.
Aspirate the supernatant and resuspend the resin in 1 mL of ice-cold lysis buffer. Repeat the centrifugation and wash steps twice.
After the final wash step, resuspend the 300 µL of resin in 300 µL of lysis buffer and add 300 µL of this slurry to each of the aliquots. Incubate at 4 °C for 1 h on a rotating wheel.
Wash each sample three times with ice-cold salt wash buffer, twice with ice-cold wash buffer, and once with ice-cold reaction buffer. After the final wash, resuspend the 150 µL of packed resin in each sample in a total volume of 175 µL of ice-cold reaction buffer.
Aliquot 32.5 µL of each slurry into eight individual 1.5 mL microcentrifuge tubes on ice. Each tube should now contain 15 µL of resin resuspended in 17.5 µL of reaction buffer, on ice.
Add 2.5 µL of 1 mg/mL GST-TIFA to each tube on ice, followed by either 2.5 µL of reaction buffer or 2.5 µL of 100 µM ADP-heptose, 1 mM UDP-mannose, or 1 mM ADP-ribose in reaction buffer on ice.
Add 2.5 µL of the radioactive ATP solution (see Recipes) to the samples at 20 s intervals and put onto a prewarmed thermomixer at 30 °C at 1,300 rpm.
Terminate reactions at the optimal timepoint determined in the preceding section by the addition of 8.3 µL of LDS sample buffer 4× containing 10% (v/v) 2-mercaptethanol and heat for 5 min at 75 °C.
Remove the resin from the samples using Spin-X columns and analyze half of the supernatant (i.e., 16.7 µL) by SDS-PAGE followed by staining with InstantBlue for 1 h and destaining for 24 h in water with frequent changes.
Scintillation counting to determine the incorporation of radioactivity into GST-TIFA:
Analyze the incorporation of radioactivity into GST-TIFA by scintillation counting using the same procedure described in Section C, steps 12–16. An example of the resulting data is shown (Figure 1D).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Snelling et al. [6]. Discovery and Functional analysis of a novel ALPK1 variant in ROSAH syndrome. bioRxiv (Figure 3, panels A and B; Figure 4, panel D)
Snelling et al. [3]. ALPK1 mutants causing ROSAH syndrome or Spiradenoma are activated by human nucleotide sugars. Proc Natl Acad Sci USA (Figure 4, panels A–D)
General notes and troubleshooting
Troubleshooting
Low or undetectable expression of ALPK1 constructs
This is most likely caused by a poor transfection efficiency, the study of a mutation that reduces the expression of ALPK1 (such as Tyr254Cys), or an issue with the immunoprecipitation procedure. To rule out the former, check the percentage of GFP-positive cells by flow cytometry or microscopy 24 h after transfection with a GFP plasmid such as pcDNA5-FRT/TO-GFP-ALPK1 (MRC PPU Reagents and Services, #DU78380). Below are general suggestions for troubleshooting.
Ensure that cells are not overly confluent prior to plating into 15 cm dishes, as this can reduce the transfection efficiency and expression of ALPK1 constructs.
Confirm that cells are not contaminated with mycoplasma or other infectious agents by routine testing.
Check DNA purity by measuring the ratio of absorbance at 260 and 280 nm using a spectrophotometer, which should be between 1.8 and 2.0. Check DNA integrity by agarose gel electrophoresis and re-sequence.
Analyze the expression of ALPK1 constructs in cell extracts by anti-FLAG immunoblotting, which will distinguish between failure to express and an issue with the immunoprecipitation procedure.
Establish the efficiency of immunoprecipitation by comparing the level of ALPK1 in cell extracts before and after immunoprecipitation (i.e., input and supernatant) by immunoblotting.
Cell death during the transfection procedure
Ensure that cells are plated to be approximately 90% confluent at the time of transfection, as it is observed that cell toxicity from transfection increases as the confluency decreases below this value.
Ensure that the culture media is replaced 4 h post-transfection, as this is observed to minimize cell toxicity.
High background phosphorylation signal
High background signal in immunoprecipitation-coupled phosphorylation assays can occur for numerous reasons, such as reactions not being within the linear range, contamination of buffers with microorganisms, inefficient washing following immunoprecipitation, or not rinsing the SDS-PAGE gel prior to excising bands. Below are general suggestions for troubleshooting a high background signal.
Ensure that reactions are within the linear range of the assay by performing time course phosphorylation experiments in both the presence and absence of ADP-heptose, as described within the protocol.
If high-level background radiation is observed in the absence of ADP-heptose, prepare fresh buffers to rule out contamination and confirm that ALPK1 KO HEK-Blue cells are not contaminated by carrying out routine testing (these may be sources of ADP-heptose in the reactions).
Perform time course phosphorylation experiments from extracts expressing FLAG-ALPK1[K1067M] (MRC PPU Reagents and Services, #DU65680), which will establish whether the background signal is arising from the immunoprecipitation or ALPK1 itself, since this is a kinase-inactive form of ALPK1.
Increase the number of washing steps with salt wash buffer to ensure that contaminating kinases are absent prior to performing phosphorylation reactions.
No or little phosphorylation of GST-TIFA in the assay
Confirm the integrity of GST-TIFA by SDS-PAGE and confirm that the concentration is correct.
Ensure that the nucleotide sugars and GST-TIFA are in suitable buffers such as those described in the protocol. For example, EDTA present in buffers may inhibit phosphorylation reactions by chelating magnesium ions.
Confirm whether ALPK1 is active by measuring the incorporation of radioactivity into ALPK1 itself (i.e., an autophosphorylation event). To do this, increase the amount of cell extract per reaction to 1 mg, as otherwise the amount of ALPK1 in the reaction will not be sufficient. As an example, see Figure 6C in [2].
Ensure that protease inhibitors and DTT are included in the lysis buffer as described in the protocol and that all steps from cell lysis onward are performed on ice with ice-cold buffers unless stated otherwise in the protocol until the LDS sample buffer is added.
Some ALPK1 mutants may be inherently unstable, which we observed in the case of Tyr254Cys [3]. It is recommended to perform experiments as quickly as possible after cell lysis and to minimize freeze-thaw.
Acknowledgments
This work was supported by a PhD studentship (#2087974) from the MRC.
Competing interests
The author declares no competing interests associated with this manuscript.
References
Zhou, P., She, Y., Dong, N., Li, P., He, H., Borio, A., Wu, Q., Lu, S., Ding, X., Cao, Y., et al. (2018). Alpha-kinase 1 is a cytosolic innate immune receptor for bacterial ADP-heptose. Nature. 561(7721): 122–126.
Snelling, T., Shpiro, N., Gourlay, R., Lamoliatte, F. and Cohen, P. (2022). Co-ordinated control of the ADP-heptose/ALPK1 signalling network by the E3 ligases TRAF6, TRAF2/c-IAP1 and LUBAC. Biochem J. 479(20): 2195–2216.
Snelling, T., Saalfrank, A., Wood, N. T. and Cohen, P. (2023). ALPK1 mutants causing ROSAH syndrome or Spiradenoma are activated by human nucleotide sugars. Proc Natl Acad Sci USA. 120(50): e2313148120.
Williams, L. B., Javed, A., Sabri, A., Morgan, D. J., Huff, C. D., Grigg, J. R., Heng, X. T., Khng, A. J., Hollink, I. H., Morrison, M. A., et al. (2019). ALPK1 missense pathogenic variant in five families leads to ROSAH syndrome, an ocular multisystem autosomal dominant disorder. Genet Med. 21(9): 2103–2115.
Kozycki, C. T., Kodati, S., Huryn, L., Wang, H., Warner, B. M., Jani, P., Hammoud, D., Abu-Asab, M. S., Jittayasothorn, Y., Mattapallil, M. J., et al. (2022). Gain-of-function mutations in ALPK1 cause an NF-κB-mediated autoinflammatory disease: functional assessment, clinical phenotyping and disease course of patients with ROSAH syndrome. Ann Rheum Dis. 81(10): 1453–1464.
Snelling, T., Garnotel, L. O., Jeru, I., Tusseau, M., Cuisset, L., Perlat, A., Minard, G., Benquey, T., Maucourant, Y., Wood, N. T., et al. (2024). Discovery and Functional analysis of a novel ALPK1 variant in ROSAH syndrome. bioRxiv. doi.org/10.1101/2024.09.13.612837.
Rashid, M., van der Horst, M., Mentzel, T., Butera, F., Ferreira, I., Pance, A., Rütten, A., Luzar, B., Marusic, Z., de Saint Aubain, N., et al. (2019). ALPK1 hotspot mutation as a driver of human spiradenoma and spiradenocarcinoma. Nat Commun. 10(1): 2213.
Article Information
Publication history
Received: Jul 25, 2024
Accepted: Sep 27, 2024
Available online: Oct 15, 2024
Published: Nov 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Biochemistry > Protein > Activity
Cell Biology > Cell signaling > Phosphorylation
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PF Phillipp Fink
JK Jong-Hee Kwon
KF Karl Forchhammer
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5125 Views: 221
Reviewed by: Dennis J NürnbergPatrick Jung Anonymous reviewer(s)
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Abstract
In modern science, the exchange of scientific material between different institutions and collaborating working groups constitutes an indispensable endeavor. For this purpose, bacterial strains are frequently shipped to collaborators to advance joint research projects. Bacterial strains are usually safely shipped as cultures on solid medium, whereas the shipment of liquid cultures requires specific safety measures due to the risk of leakage. Cyanobacterial cultures are frequently maintained as liquid stock cultures, and this problem typically arises. This protocol describes a new method for the shipment of liquid cyanobacterial stock cultures by agarose gel embedding (SCAGE). More specifically, a cyanobacterial culture is mixed with low-melting agarose and cast into sterile plastic bags, resulting in a thin, solid cyanobacterial agarose gel (cyanogel) that can be easily shipped. After delivery, subsequent regeneration of the cyanogel material in liquid media results in full recovery of the examined bacterial strains. Thus, the packaging method devised in the present study comprises an innovative technique to facilitate the shipment of bacterial strains, whilst eliminating previously encountered issues like cell culture leakage.
Key features
• New packaging procedure to reduce culture leakage.
• Novel technique facilitating improved shipment conditions.
• Validated method leading to recovery of tested bacterial strains after 14 days.
Keywords: Synechocystis sp. PCC6803 GT Anabaena (Nostoc) sp. PCC7120 Agarose gel embedding Shipment of cyanobacteria Cyanobacterial culture collection
Graphical overview
Schematic representation of steps for gel embedding and recovery of cyanobacteria
Background
Cyanobacteria are photoautotrophic microorganisms that constitute one of the biggest, most diverse genera of prokaryotes [1]. Among their many characteristics, their ability to produce oxygen and fixate carbon dioxide (CO2) has piqued the interest of scientists worldwide. In recent years, a critical need for a sustainable, CO2-neutral production platform of biofuels and chemicals has emerged and, hence, the investigation of cyanobacterial cell factories has exponentially gained significance [2–4]. Further research on these promising applications of cyanobacteria is necessary and will efficiently accelerate through collaborations between the scientific community.
The exchange of cyanobacterial strains between laboratories is a common procedure to advance joint research projects. Traditionally, bacterial strains are shipped on solid agar plates, in liquid cultures, or cryopreserved and shipped on dry ice. Solid agar plates are effective at preventing leakage but are prone to desiccation, particularly during prolonged shipment. Liquid cultures are suitable for short-term transport but require more complex packaging to prevent leakage. Prolonged shipment under the abovementioned conditions increases the risk of loss of viability of cyanobacterial cells. Cryopreservation is widely used for the long-term preservation of eukaryotic cell lines [5], bacteria, and cyanobacteria [6]; however, shipping cryopreserved samples requires specialized equipment and incurs high costs. Additionally, the shipment of samples on dry ice is prohibited in more than 50% of countries worldwide [7]. Moreover, depending on the cyanobacterial strain, the “reawakening” process can take up to 30 days [6].
Herein, we describe a shipment method for cyanobacteria that increases shipment safety and maintains cells for prolonged times in a viable state by embedding the bacterial strain in an agarose matrix (shipment of cyanobacteria by agarose gel embedding, SCAGE). This agarose gel (cyanogel) is inherently unlikely to display leakage due to the solid state of the material. Moreover, the risk of bacterial desiccation or damage to bacterial vials is greatly reduced due to the embedment in a sterile sealed bag. The casting of thin cyanogels allows for low-cost, efficient shipment in padded envelopes. Furthermore, cyanobacterial cells remain viable and can be successfully recovered after 14 days. A recent study also demonstrated the effective transportation of eukaryotic cell lines using agarose gel-based preservation [8]. This protocol highlights the casting of cyanogels and the subsequent recovery procedure for cyanobacteria.
Materials and reagents
Biological materials
Synechocystis sp. PCC6803, glucose-tolerant (GT) [9]
Anabaena (Nostoc) sp. PCC7120 [10]
Reagents
NaNO3 (Sigma-Aldrich, CAS: 7631-99-4)
K2HPO4 (water-free) (Sigma-Aldrich, CAS: 7758-11-4)
MgSO4·7H2O (AppliChem, CAS: 10034-99-8)
CaCl2·2H2O (Merck, CAS: 10035-04-8)
Na2-EDTA·2H2O (Merck, CAS: 6381-92-6)
Na2CO3 (Merck, CAS: 497-19-8)
Fe(III)-citrate (Sigma-Aldrich, CAS: 3522-50-7)
Citric acid (Roth, CAS: 77-92-9)
H3BO3 (Merck, CAS: 10043-35-3)
MnCl2·4H2O (Roth, CAS: 13446-34-9)
ZnSO4·7H2O (Fluka, CAS: 7758-99-8)
Na2MoO4·2H2O (Sigma-Aldrich, CAS: 10102-40-6)
CuSO4·5H2O (Fluka, CAS: 7758-99-8)
Co(NO3)2·6H2O (Sigma-Aldrich, CAS: 10026-22-9)
NaHCO3 (Fisher Chemica, CAS: 144-55-8)
Agarose low melt (Roth, CAS: 39346-81-1)
Solutions
BG11 medium (see Recipes)
5% (w/v) agarose low melting (ALM) solution (see Recipes)
Recipes
BG11 medium (modified after Rippka et al. [11]):
Prepare 200 mL of each stock solution (1–7) from Table 1, 100 mL of trace element solution (Table 2), and 50 mL of 1 M NaHCO3 solution (Table 3). Use Milli-Q water for each solution.
Stock solutions 1–7 and the trace element solution are sterilized by autoclaving (121 °C, 15 psi, 20 min), while the 1 M NaHCO3 solution is sterilized by filter sterilization (0.22 μm).
Note: Store solutions 1–6 at room temperature and solution 7 [Fe(III)-citrate & citric acid] protected from light at room temperature. Store trace solution at 4 °C and 1 M NaHCO3 at -20 °C.
To prepare 1 L of BG11 (1×), add 5 mL of each stock solution 1–7 (200×), 1 mL of the trace element solution (1,000×), and up to 995 mL of Milli-Q water. Sterilize by autoclaving (121 °C, 15 psi, 20 min). Add 5 mL of 1 M NaHCO3 solution before use.
Note: For the modified BG11 medium, conduct the following adjustments:
Na2CO3: 0.04 g/L (originally 0.02 g/L).
Ferric ammonium citrate was replaced by Fe(III)-citrate.
Table 1. Stock solutions (200×) for BG11 medium
Solution Chemical Molar mass M (g/mol) Molarity c (mM) in 200× stock solution Mass concentration β (g/L) in 200× stock solution Molarity c (mM) 1× medium Mass concentration β (g/L) in 1× medium Mass m (g) in 200 mL stock solution
1 NaNO3 84.99 35230 300 17.65 1.5 60
2 K2HPO4 (water-free) 174.18 35.9 6.25 0.18 0.03125 1.25
3 MgSO4·7H2O 246.48 60.9 14.8 0.3 0.075 3
4 CaCl2 ·2H2O 147.02 49.0 7.2 0.24 0.036 1.44
5 Na2-EDTA·2H2O 372.24 0.5 0.2 0.003 0.001 0.04
6 Na2CO3 105.99 75.5 8 0.38 0.04 1.6
7 Fe(III)-citrate 244.94 4.9 1.2 0.024 0.006 0.24
8 Citric acid 192.13 6.2 1.2 0.031 0.006 0.24
Table 2. Trace element solution (1,000×) for BG11 medium
Chemical Molar mass M (g/mol) Molarity c (mM) in 1,000× stock solution Mass concentration β (g/L) in 1,000× stock solution Molarity c (μM) 1× medium Mass m (mg) in 100 mL stock solution
H3BO3 61.83 46.26 2.86 46.26 286
MnCl2·4H2O 197.91 9.15 1.81 9.15 181
ZnSO4·7H2O 287.54 0.77 0.222 0.77 22.2
Na2MoO4·2H2O 241.95 1.61 0.39 1.61 39
CuSO4·5H2O 249.68 0.32 0.079 0.32 7.9
Co(NO3)2·6H2O 291.03 0.17 0.0494 0.17 4.9
Table 3. 1 M NaHCO3 stock solution for BG11 medium
Chemical Molar mass M (g/mol) Molarity c (mM) Mass concentration β (g/L) in 1 M stock solution Molarity c (mM) 1× medium Mass concentration β (g/L) in 1× medium Mass m (g) in 100 mL stock solution
NaHCO3 84.01 1,000 84 5 0.42 8.4
5% (w/v) agarose low melting (ALM) solution
To prepare 100 mL of 5% (w/v) ALM solution, add 5 g of ALM in 100 mL of BG11 medium. Sterilize by autoclaving (121 °C, 15 psi, 20 min). Store at room temperature.
Laboratory supplies
Pipette tips with filter (Mettler Toledo, type RAININ)
Serological pipette, 5 mL (Sarstedt, catalog number: 86.1253.001)
Conical centrifuge tubes, 15 mL (Sarstedt, catalog number: 62.554.502)
Disposal bags (Roth, catalog number: E706.1)
Grease-proof paper Profissimo (DM-Drogerie Markt, catalog number: 470799)
Rubber band “Alco 758” 80 × 4 mm (Lyreco, catalog number: 5.112.077)
Steristopper (VWR, catalog number: HERE1013700)
Plastic cuvettes (Sarstedt, catalog number: 67.742)
Erlenmeyer wide-neck flask DIN ISO 24450, 100 mL (VWR, catalog number: 214-1131)
Laboratory bottle, 1 L (VWR, catalog number: 215-1517P)
Beaker, 800 mL (Roth, catalog number: X694.1)
Aluminum foil (Roth, catalog number: 1399.1)
Sterile indicator strip (Roth, catalog number: XC20.1)
Multimark overhead marker permanent (Faber-Castell, catalog number: 151304)
Equipment
Pipettes (Mettler Toledo, model: Pipet-Lite XLS series)
Pipetboy (VWR, catalog number: 613-4438)
Rotary shaker (GFL, catalog number: GFL-3017)
OSRAM LUMILUM DE LUX Daylight lamp (OSRAM, catalog number: L58W/954)
Biosafety cabinet (Bioquell, model: ABS1500CLS2-MK2)
Microwave (OK, model: OMW 330 D-M)
Centrifuge 5804 R (Eppendorf, catalog number: 5804 R)
Centrifuge rotor S-4-72 (Eppendorf, catalog number: S-4-72)
Photometer (Thermo Scientific, model: Spectronic Helios model δ)
Foil welding apparatus (BOSCH, catalog number: FG16-0717901002)
Procedure
Note: All steps should be carried out under sterile conditions.
Cultivation of cyanobacteria
Put the Steristopper (paper plug) on top of clean Erlenmeyer flasks and cover the Erlenmeyer flasks with grease-proof paper. Secure the grease-proof paper with a rubber band.
Sterilize the prepared Erlenmeyer flask by autoclaving (121 °C, 15 psi, 20 min) and let it cool to room temperature.
Open the Erlenmeyer flask under sterile conditions and inoculate Synechocystis sp. PCC6803 GT or Anabaena (Nostoc) sp. PCC7120 in 50 mL of BG11 medium. Close the Erlenmeyer flask after inoculation.
Grow Synechocystis sp. PCC6803 GT or Anabaena (Nostoc) sp. PCC7120 under continuous light illumination (~ 40 μmol m-2·s-1) at 120 rpm and 28 °C until the stationary phase (OD750 > 4) is reached.
Preparation for gel embedding of cyanobacteria
Sterilize disposable bags in a beaker covered with aluminum foil by autoclaving (121 °C, 15 psi, 20 min) and let them cool down to room temperature.
Prepare 5% (w/v) ALM solution (see Recipes).
Gel embedding
Collect 15 mL of liquid bacterial culture in a conical reaction tube and harvest cells by centrifugation at 4000× g for 10 min at room temperature. Discard the supernatant and resuspend cells in 2 mL of BG11 medium.
Simultaneously, heat the 5% (w/v) ALM suspension until a clear, homogenous solution is obtained.
Caution: The flask containing the ALM solution will be hot and even a boiling delay could be observed. Open the cap of the bottle slightly. Use appropriate safety measures to prevent burning yourself.
Let the solution cool down to approximately 30 °C under the sterile hood.
Once the appropriate temperature is reached, add 3 mL of the 5% (w/v) ALM solution to the resuspended cyanobacterial strain (obtained in step C1) and mix through careful pipetting, resulting in the cyanogel.
Note: Re-use the serological pipette to mix the components to save time before the cyanogel is solidified. The concentration of ALM in the resulting cyanogel will be 3% (w/v).
Open the disposable bag under sterile conditions and transfer the cyanogel into the bag by slow and careful pipetting.
Situate the disposable bag containing the gel on a flat surface and apply a slight amount of pressure with your hands on the upper side of the bag to evenly distribute the cyanogel within the bag. A smooth plane should be obtained (8 cm × 20 cm × ≤1 mm), which can then be sealed with the foil welding machine (section D).
Sealing the cyanogel
Place the bag between the contacts of a foil welding machine and close the machine for 3 s.
Caution: Be cautious when pressing the machine down since the contacts will get hot. Take measures to prevent burning yourself.
Open the apparatus again and wait for 5 s.
Note: Removing the cyanogel before letting it cool may result in deformation or breakage of the bag. Usually, 5 s is enough to let the sealed bag cool down.
Label the bag appropriately, e.g., with a permanent marker, for later identification of the cyanobacterial strain contained within.
Packaging of embedded cyanobacteria
Package the embedded cyanobacteria in an appropriate container (e.g., an envelope) and send it to the location of your choice.
Note: Depending on the destination of your cyanobacterial samples, different regulations regarding labeling may apply.
Recovering of cyanobacteria out of the cyanogel
Prepare 50 mL of BG11 medium in an Erlenmeyer flask under sterile conditions.
Open the received cyanogel and squeeze the gel out of the disposal bag into the BG11 medium.
Note: Use a sterile pair of scissors for cutting an edge of the cyanogel for easier and more precise gel squeezing.
Let the cyanobacteria grow under dimmed light conditions (flask wrapped with a paper towel) of ~20 µmol m-2·s-1 at 120 rpm for 24 h at 28 °C.
After 24 h of incubation, remove the paper towel and cultivate the cyanobacteria under standard cultivation conditions.
Validation of protocol
For validation of the protocol presented here, the procedure was performed with two different cyanobacterial strains [Synechocystis sp. PCC6803 and Anabaena (Nostoc) sp. PCC7120]. Synechocystis and Anabaena (Nostoc) were cultivated under standard conditions until an OD750 of 9.3 and 5.2 was reached, respectively. 25 mL of culture was centrifuged and resuspended in 8 mL of BG11 medium; 2 mL of resuspended cyanobacteria were mixed with 3 mL of BG11 medium and used as a control group. Triplicates of 2 mL culture were mixed with 3 mL of 5% (w/v) ALM and cast into disposable bags. Bags were sealed with a foil welding machine and were stored for one week in a non-transparent, padded envelope at room temperature in the dark to simulate packaging shipment (Figure 1).
Figure 1. Disposal bags filled with cyanobacteria. Left: Synechocystis sp. PCC6803 GT; right: Anabaena (Nostoc) sp. PCC7120 embedded in 5% (w/v) ALM [cyanogel, end concentration 3% (w/v) ALM] before and after shipment simulation.
After one week of shipment simulation, regeneration of cyanobacterial strains was performed according to the described method. Full recovery of both strains was reached after 14 days of cultivation (Figure 2).
Figure 2. Recovery of cyanobacteria over 14 days. Left: Synechocystis sp. PCC6803 GT; right: Anabaena (Nostoc) sp. PCC7120.
Acknowledgments
This project was conducted with the support of the Korea Evaluation Institute of Industrial Technology (KEIT) grant funded by the Korean government (MOTIE) (No. RS-2022-00155902) and Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (2021R1I1A3055799).
Competing interests
The authors have no conflicts of interest to declare.
References
Stanier, R. Y. and Cohen-Bazire, G. (1977). PHOTOTROPHIC PROKARYOTES: THE CYANOBACTERIA. Annu Rev Microbiol. 31(1): 225–274. https://doi.org/10.1146/annurev.mi.31.100177.001301
Clark, J. H., Luque, R. and Matharu, A. S. (2012). Green Chemistry, Biofuels, and Biorefinery. Annu Rev Chem Biomol Eng. 3(1): 183–207. https://doi.org/10.1146/annurev-chembioeng-062011-081014
Gao, X., Sun, T., Pei, G., Chen, L. and Zhang, W. (2016). Cyanobacterial chassis engineering for enhancing production of biofuels and chemicals. Appl Microbiol Biotechnol. 100(8): 3401–3413. https://doi.org/10.1007/s00253-016-7374-2
Wirth, T. E., Gray, C. B. and Podesta, J. D. (2003). The Future of Energy Policy. Foreign Affairs. 82. http://dx.doi.org/10.2307/20033654
Heydarzadeh, S., Kheradmand Kia, S., Boroomand, S. and Hedayati, M. (2022). Recent developments in cell shipping methods. Biotechnol Bioeng. 119(11): 2985–3006. https://doi.org/10.1002/bit.28197
Esteves-Ferreira, A. A., Corrêa, D. M., Carneiro, A. P. S., Rosa, R. M., Loterio, R. and Araújo, W. L. (2012). Comparative evaluation of different preservation methods for cyanobacterial strains. J Appl Phycol. 25(4): 919–929. https://doi.org/10.1007/s10811-012-9927-9
FedEx. (2024). Dangerous Goods - Countries and Territories Served. FedEx. Retrieved August 28, 2024, from https://www.fedex.com/en-us/service-guide/dangerous-goods/international-locations.html
Yang, L., Li, C., Chen, L. and Li, Z. (2009). An Agarose-Gel Based Method for Transporting Cell Lines. Curr Chem Genomics. 3: 50–53. https://doi.org/10.2174/1875397300903010050
Kaneko, T., Sato, S., Kotani, H., Tanaka, A., Asamizu, E., Nakamura, Y., Miyajima, N., Hirosawa, M., Sugiura, M., Sasamoto, S., et al. (1996). Sequence Analysis of the Genome of the Unicellular Cyanobacterium Synechocystis sp. Strain PCC6803. II. Sequence Determination of the Entire Genome and Assignment of Potential Protein-coding Regions. DNA Res. 3(3): 109–136. https://doi.org/10.1093/dnares/3.3.109
Kaneko, T. (2001). Complete Genomic Sequence of the Filamentous Nitrogen-fixing Cyanobacterium Anabaena sp. Strain PCC 7120. DNA Res. 8(5): 205–213. https://doi.org/10.1093/dnares/8.5.205
Rippka, R., Stanier, R. Y., Deruelles, J., Herdman, M. and Waterbury, J. B. (1979). Generic Assignments, Strain Histories and Properties of Pure Cultures of Cyanobacteria. Microbiology (N Y). 111(1): 1–61. https://doi.org/10.1099/00221287-111-1-1
Article Information
Publication history
Received: Jul 8, 2024
Accepted: Sep 17, 2024
Available online: Oct 22, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Microbiology > Microbial cell biology > Cell viability
Cell Biology > Cell viability > Cell survival
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In-Gel Activity Assay of Mammalian Mitochondrial and Cytosolic Aconitases, Surrogate Markers of Compartment-Specific Oxidative Stress and Iron Status
WT Wing-Hang Tong
TR Tracey A. Rouault
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5126 Views: 292
Reviewed by: Elizabeth Calzada Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Proceedings of the National Academy of Sciences of the United States of America Aug 2008
Abstract
Two aconitase isoforms are present in mammalian cells: the mitochondrial aconitase (ACO2) that catalyzes the reversible isomerization of citrate to isocitrate in the citric acid cycle, and the bifunctional cytosolic enzyme (ACO1), which also plays a role as an RNA-binding protein in the regulation of intracellular iron metabolism. Aconitase activities in the different subcellular compartments can be selectively inactivated by different genetic defects, iron depletion, and oxidative or nitrative stress. Aconitase contains a [4Fe-4S]2+ cluster that is essential for substrate coordination and catalysis. Many Fe-S clusters are sensitive to oxidative stress, nitrative stress, and reduced iron availability, which forms the basis of redox- and iron-mediated regulation of intermediary metabolism via aconitase and other Fe-S cluster-containing metabolic enzymes, such as succinate dehydrogenase. As such, ACO1 and ACO2 activities can serve as compartment-specific surrogate markers of oxygen levels, reactive oxygen species (ROS), reactive nitrogen species (RNS), iron bioavailability, and the status of intermediary and iron metabolism. Here, we provide a protocol describing a non-denaturing polyacrylamide gel electrophoresis (PAGE)-based procedure that has been successfully used to monitor ACO1 and ACO2 aconitase activities simultaneously in human and mouse cells and tissues.
Key features
• Monitoring aconitase activity changes in the mitochondria and cytosol simultaneously in response to oxidative or nitrative stress, iron depletion, and various pathophysiological conditions.
• Optimized for human and mouse cell lines and tissue samples.
• Semi-quantitative detection of aconitase isoforms with different states of phosphorylation and/or post-translational modification.
Keywords: Aconitase ACO1 ACO2 Iron response element binding protein 1 IRE-BP1 Iron regulatory protein 1 IRP1 Iron metabolism Oxidative stress Nitrative stress
Background
Aconitases play a central role in the crosstalk between citrate and iron metabolism [1]. Aconitase is best known for its metabolic function in the mitochondrial citric acid cycle, the primary catabolic pathway that reduces NADP+ to generate NADPH that feeds into the oxidative phosphorylation pathway for ATP production. In addition to the mitochondrial isoform ACO2, mammalian cells express various levels of a bifunctional cytosolic ACO1 that can register iron and oxidative stress through its labile Fe–S cluster. Upon the loss of its Fe-S cluster, ACO1 converts to an iron response element (IRE) binding protein IRE-BP1 (also known as iron regulatory protein 1, IRP1), which binds to IRE-containing RNAs and thereby regulates the expression of various proteins that are important in iron metabolism, including iron storage protein ferritin, iron transport proteins (transferrin receptor TfR1 and ferroportin), and erythrocyte-specific heme biosynthesis protein aminolevulinate synthase, among several targets [1]. Thus, the spatial separation of ACO1/IRP1 and ACO2 allows them to serve as indexes of compartment-specific Fe-S cluster biogenesis/repair pathways, oxidative stress, nitrative stress, and iron status.
The UV/vis spectrophotometric coupled enzyme assay that couples the reactions of aconitase and NADP-dependent (IDH) has offered a very sensitive method for detecting and measuring aconitase activities in purified protein or in whole-cell lysates [2]. However, although subcellular fractions can be used for assaying ACO1 and ACO2 separately, the time and laborious procedure needed for obtaining relatively pure subcellular fractions often results in the loss of the Fe-S cluster and diminished enzyme activities. Mitochondrial and cytosolic aconitase activities in whole-Drosophila extracts have also been assayed jointly after electrophoretic separation on cellulose acetate membranes [3]; however, empirical testing showed that the method was not suitable for quantitative analysis of mammalian aconitase activities, given the different pKa values of aconitases in different species. In addition, it is not possible to know the exact amount of protein loaded onto the cellulose acetate membrane, and the signals from mammalian cell line extracts are much weaker compared to those from the whole fly extracts.
The current protocol describes the non-denaturing polyacrylamide gels and buffers for in-gel assay for human or mouse aconitase activities (adapted from previous work [4]). Aconitase activity was detected chromogenically by incubating the gel after electrophoresis in a coupled enzyme assay mix containing cis-aconitate, NADP-dependent isocitrate dehydrogenase, NADP, thiazolyl blue tetrazolium bromide and phenazine methosulfate. Quantification of the activity of each aconitase isoform is carried out by densitometric analysis.
Materials and reagents
Reagents
Dry ice
Triton X-100 (Sigma-Aldrich, catalog number: T-8787)
RIPA extraction buffer (e.g., RIPA Lysis and Extraction Buffer, Thermo Scientific, catalog number: 89900, ts: 25 mM Tris·HCl pH 7.6, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS)
Potassium chloride (KCl) (Sigma, catalog number: P3911)
Protein assay reagents (e.g., Protein Assay Kit I, Bio-Rad, catalog number: 5000001EDU)
Ethanol (generic)
30% (w/v) acrylamide/methylene bisacrylamide solution (37.5:1 ratio) (e.g., ProtoGel 30%, National diagnostics, catalog number: EC-890)
TEMED (National Diagnostics, catalog number: EC-503)
Ammonium persulfate (National Diagnostics, catalog number: EC-504)
Tris base (e.g., Sigma, catalog number: T1503)
Boric acid (Sigma-Aldrich, catalog number: B0394)
Glycine (Sigma-Aldrich, catalog number: G7126)
Sodium citrate (Sigma-Aldrich, catalog number: S4641)
NADP-dependent IDH (Sigma-Aldrich, catalog number: I-2002)
Cis-aconitic acid (Sigma-Aldrich, catalog number: A-3412)
β-NADP (Sigma-Aldrich, catalog number: N-0505)
Thiazolyl blue tetrazolium bromide (also known as MTT) (Sigma-Aldrich, catalog number: M2128)
Phenazine methosulfate (PMS) (Sigma-Aldrich, catalog number: P-9625)
Magnesium chloride (MgCl2) (Sigma-Aldrich, catalog number: M4880)
Protease inhibitor cocktail (e.g., Roche Complete mini protease inhibitor cocktail tablet, Millipore Sigma, catalog number: 11836153001)
Dithiothreitol (DTT) (e.g., Pierce DTT, No-WeightTM Format Thermo Fisher Scientific, catalog number: A39255)
Glycerol (generic)
Manganese chloride (MnCl2) (Sigma-Aldrich, catalog number: M1787)
Bromophenol blue (Bio-Rad, catalog number: 1610404)
Pre-stained protein standard (e.g., Thermo Fisher SeeBlue Plus2, catalog number: LC5925)
Optional: ACO2 antibody (Proteintech, catalog number: 11134-1-AP)
Optional: Tubulin antibody (Sigma-Aldrich, catalog number: T9026)
Phosphate buffered saline (PBS) (e.g., Corning, catalog number: 46-013-CM)
Isopropyl alcohol (e.g., Sigma-Aldrich, catalog number: I9516)
Stock reagents:
1 M Tris-HCl (adjusted to pH 7.5 and 8.0 with HCl), store at room temperature (RT)
40 mM KCl, 25 mM Tris-Cl, pH 7.5, store at RT
25% Triton, store at RT
1 M sodium citrate (500×), store at -20 °C
20 mM MnCl2, store at -20 °C
20 mM β-NADP (20×), store at -20 °C
50 mM cis-aconitate (20×), store at -20 °C
1 M MgCl2, store at -20 °C
0.5 mg/20 μL PMS, store at -20 °C
10% ammonium persulfate (APS) (prepare fresh)
0.1 M DTT (prepare fresh)
Solutions
1% Triton-citrate extraction buffer (see Recipes)
RIPA-citrate extraction buffer (see Recipes)
Electrophoresis buffers for the human aconitase in-gel activity assay (see Recipes)
10× Tris-glycine-8.3 stock for running buffer
Running buffer: Tris-glycine-citrate-8.3
Gel buffer: Tris-borate-8.3
4× loading buffer
Gel recipes for the human aconitase in-gel activity assay (see Recipes)
Separating gel for human aconitases
Stacking gel for human aconitases
Electrophoresis buffers for the mouse aconitase in-gel activity assay (see Recipes)
10× Tris-glycine-8.7 stock for running buffer
Running buffer: Tris-glycine-citrate-8.7
Gel buffer: Tris-borate-8.7
4× loading buffer
Gel recipes for the mouse aconitase in-gel activity assay (see Recipes)
Separating gel for mouse aconitases
Stacking gel for mouse aconitases
Aconitase activity coupled enzyme assay mix (see Recipes)
Recipes
1% Triton-citrate extraction buffer
Reagent Final concentration Quantity or Volume
Buffer (40 mM KCl, 25 mM Tris-Cl, pH 7.5) 9.18 mL
Triton 25% 1% 0.40 mL
Roche Complete mini protease inhibitor cocktail tablet 1
DTT 0.1 M (made fresh) 1.0 mM 0.10 mL
Sodium citrate, 1 M, 500× 2.0 mM 0.020 mL
MnCl2, 20 mM 0.60 mM 0.30 mL
Total volume 10 mL
Notes:
Store the Triton-citrate extraction buffer in 0.5–1 mL aliquots at -20 °C for 3–6 months.
Citrate is added to stabilize the Fe-S cluster in aconitase.
RIPA-citrate extraction buffer
Reagent Final concentration Quantity or Volume
RIPA Lysis and Extraction Buffer 9.58 mL
Roche Complete mini protease inhibitor cocktail tablet 1
DTT 0.1 M (made fresh) 1.0 mM 0.10 mL
Sodium citrate, 1 M, 500× 2.0 mM 0.020 mL
MnCl2, 20 mM 0.60 mM 0.30 mL
Total volume 10 mL
Notes:
Store the RIPA-citrate extraction buffer in 0.5–1 mL aliquots at -20 °C for 3–6 months.
Citrate is added to stabilize the Fe-S cluster in aconitase.
Electrophoresis buffers for the human aconitase in-gel activity assay
10× Tris-glycine-8.3 stock for running buffer
Reagent Final concentration Quantity or Volume
Tris base 250 mM 30.2 g
Glycine 1.92 M 144 g
H2O Enough to make 1 L buffer
Total volume 1.0 L
Note: Do not adjust the pH of this 10× stock. The pH of this stock will be ~8.3.
Running buffer: Tris-glycine-citrate-8.3
Reagent Final concentration Quantity or Volume
10× Tris-glycine-8.3 stock 1× 80 mL
1 M sodium citrate 3.6 mM 2.88 mL
H2O Enough to make 0.8 L buffer
Total volume 0.80 L
Gel buffer: Tris-borate-8.3
Reagent Final concentration Quantity or Volume
Tris base 890 mM 108 g
Boric acid 890 mM 55 g
H2O Enough to make 1 L buffer
Total volume 1.0 L
Note: Do not adjust the pH of this stock. The pH of this buffer will be ~8.3.
4× loading buffer
Reagent Final concentration Quantity or Volume
1 M Tris-Cl pH 8.0 100 mM 0.10 mL
Glycerol 40% 0.40 mL
Bromophenol blue 1 mg 0.1%
H2O Enough to make 1 mL buffer
Total volume 1.0 mL
Gel recipes for the human aconitase in-gel activity assay
For HeLa cell extract, an 8% acrylamide separating gel at pH 8.3 and electrophoresis at 170 V for 2.5 h at 4 °C gives a good resolution of two bands corresponding to ACO1 and ACO2. For better resolution of samples that contain more than two isoforms (e.g., aconitases with different phosphorylation), a 6% or 7% acrylamide separating gel may be used.
Separating gel for human aconitases
8% acrylamide
Tris-borate-citrate-8.3
mini gel
6% acrylamide
Tris-borate-citrate-8.3
mini gel
8% acrylamide
Tris-borate-citrate-8.3
big gel
ProtoGel 30% 1.68 mL 1.26 mL 6.7 mL
Tris-borate-8.3 gel buffer 0.94 mL 0.94 mL 3.75 mL
H2O 3.64 mL 4.06 mL 14.55 mL
Sodium citrate, 1 M 22.5 μL 22.5 μL 90 μL
APS 10% 31 μL 31 μL 125 μL
TEMED 6.25 μL 6.25 μL 25 μL
Stacking gel for human aconitases
4% acrylamide
Tris-borate-citrate-8.3
mini gel
4% acrylamide
Tris-borate-citrate-8.3
big gel
ProtoGel 30% 0.20 mL 0.80 mL
Tris-borate-8.3 gel buffer 0.11 mL 0.45 mL
H2O 1.16 mL 4.65 mL
Sodium citrate, 1 M 4.2 μL 21 μL
APS 10% 21 μL 84 μL
TEMED 3.75 μL 15 μL
Electrophoresis buffers for the mouse aconitase in-gel activity assay
10× Tris-glycine-8.7 stock for running buffer
Reagent Final concentration Quantity or Volume
Tris base 250 mM 30.2 g
Glycine 960 mM 72 g
H2O Enough to make 1 L buffer
Total volume 1.0 L
Note: Do not adjust the pH of this 10× stock.
Running buffer: Tris-glycine-citrate-8.7
Reagent Final concentration Quantity or Volume
10× Tris-glycine-8.7 stock 1× 80 mL
1 M Sodium citrate 3.6 mM 2.88 mL
H2O Enough to make 0.8 L buffer
Total volume 0.80 L
Gel buffer: Tris-borate-8.7
Reagent Final concentration Quantity or Volume
Tris base 890 mM 54 g
Boric acid 445 mM 13.75 g
H2O Enough to make 0.5 L buffer
Total volume 0.50 L
Note: Do not adjust the pH of this stock.
4× loading buffer
Reagent Final concentration Quantity or Volume
1 M Tris-Cl pH 8.0 100 mM 0.10 mL
Glycerol 40% 0.40 mL
Bromophenol blue 1 mg 0.10%
H2O Enough to make 1 mL buffer
Total volume 1.0 mL
Gel recipes for the mouse aconitase in-gel activity assay
For mouse liver samples, a 5%–6% acrylamide Tris-borate-citrate-8.7 separating gel and 4%–5% stacking gel and electrophoresis at 170 V for 3–4 h at 4 °C gives nice resolution and detection of two ACO1 bands and two ACO2 bands.
Separating gel for mouse aconitases
5% acrylamide
Tris-borate-citrate-8.7
mini gel
6% acrylamide
Tris-borate-citrate-8.7
mini gel
ProtoGel 30% 1.1 mL 1.26 mL
Tris-borate-8.7 gel buffer 0.94 mL 0.94 mL
H2O 4.22 mL 4.06 mL
Sodium citrate, 1 M 22.5 μL 22.5 μL
APS 10% 31 μL 31 μL
TEMED 6.5 μL 6.5 μL
Stacking gel for mouse aconitases
4% acrylamide
Tris-borate-citrate-8.7
mini gel
5% acrylamide
Tris-borate-citrate-8.7
mini gel
ProtoGel 30% 0.20 mL 0.25 mL
Tris-borate-8.7 buffer 0.11 mL 0.11 mL
H2O 1.16 mL 1.11 mL
Sodium citrate, 1 M 4.2 μL 4.2 μL
APS 10% 21 μL 21 μL
TEMED 3.75 μL 3.75 μL
Aconitase activity coupled enzyme assay mix
Prepare fresh immediately before use.
Reagent Final concentration Quantity or Volume
Tris, 1 M, pH 8.0 100 mM 1.0 mL
NADP, 20 mM (20×) 1.0 mM 0.50 mL
cis-aconitate 50 mM (20×) 2.5 mM 0.50 mL
MgCl2 1 M, (200×) 5.0 mM 0.050 mL
MTT, 24 mM (20×) 1.2 mM 0.50 mL
PMS (0.5 mg/20 μL) 40 μL
IDH 5 U/mL Dependent on specific activity
H2O ~7.40 mL
Total 10 mL
Notes:
Use 10–15 mL for small gels and more for big gels.
MTT is light-sensitive. Prewarm H2O at 37 °C in a light-protected container. Add the remaining reagents, IDH last, right before staining.
Laboratory supplies
Snap-cap or screw-cap microcentrifuge tubes that can be closed tightly (e.g., USA Scientific, catalog number: 1615-5500)
Homogenization beads (e.g., Next Advance ZrOB10 Zirconium Oxide Beads 1.0 mm)
Equipment
Refrigerated microcentrifuge
Homogenizer/cell disruptor (e.g., Next Advance Bullet Blender or mortar and pestle)
Vertical gel electrophoresis system (e.g., Bio-Rad Laboratories, model: Mini-PROTEAN® II)
Mini-gel glass plates (e.g., Gel Company, model: GBS07B-10S or GBS07L-10)
Mini-gel 10-lane or 12-lane 1.0 mm thick comb (Bio-Rad or Gel Company)
Mini-gel 10-lane or 12-lane 1.0 mm thick spacers (Bio-Rad or Gel Company)
pH meter
Scanner
Gel dryer
Software and datasets
Densitometric analysis software, e.g., ImageJ or similar software
Procedure
Prepare cell culture extracts
Notes:
Although protein extracts stored at -80 °C have been used successfully for aconitase in-gel assay, it is recommended to use fresh extracts because it generally results in a stronger signal and better-defined bands of activity. Repeated sample freeze-thaw can also decrease enzymatic activity.
If using fresh protein extracts for the non-denaturing PAGE, prepare the extracts while the gel is polymerizing and cooling (steps C2–4).
Keep reagents and tubes containing cell pellets and extracts at 0–4 °C between steps. The addition of the aconitase substrate citrate (or isocitrate) to the extraction and electrophoresis buffers protects the Fe-S cluster and markedly reduces the loss of aconitase activity.
Try to keep extract protein concentration high, e.g., 20–40 μg/μL, to keep the loading volume to a minimum for better resolution and to minimize the amount of salt and detergent loaded into the gel.
Harvest cell cultures and wash cell pellets 3× with PBS at 4 °C. Lyse the cell pellets for analysis immediately or flash-freeze cell pellets on dry ice. Store cell pellets at -80 °C until use.
Note: One 100 mm plate of near-confluent HeLa S3 cells yields approximately 1 mg of protein.
Lyse cell pellets using Triton-citrate or RIPA-citrate extraction buffer at 4 °C. Add 1–2 v/v of Triton-citrate or RIPA-citrate extraction buffer to each cell pellet. Disperse cells using a pipette. Incubate on ice for 10 min, with intermittent agitation. Centrifuge at >16,000× g for 5 min in a refrigerated microcentrifuge. Collect the supernatant (protein extract). Repeat the centrifugation step. Use the extracts immediately or store at -80 °C.
Notes:
A concentrated protein extract rather than a diluted extract is better for detection and resolution in the aconitase in-gel assay. Protein extract containing 20–40 μg of protein per microliter is a good starting point.
Aliquot samples if necessary. Repeated sample freeze-thaw can decrease enzymatic activity.
Proceed with protein quantification.
Note: We use Bradford Protein Assay from Bio-Rad and follow the manufacturers’ protocol. Other protein quantification methods may be used. For instance, a BCA detection kit can be used for extracts made without DTT.
Prepare tissue extracts
Notes:
Although extracts stored at -80 °C have been used successfully, it is recommended to use fresh extracts because it generally results in stronger signals and better-defined bands of activity. Repeated sample freeze-thaw can decrease enzymatic activity.
If using fresh protein extracts for the non-denaturing PAGE, prepare the extracts while the gel is polymerizing and cooling (steps C2–4).
Keep reagents and tubes containing cell pellets and extracts at 4 °C or on ice between steps. Set up the Bullet Blender in the cold room.
Oxygen levels vary between 0% and 19% in healthy mammalian tissues. Aconitase activity levels vary significantly in different mammalian tissues because of different expression levels and the tissue-specific Oz levels. Tissue extraction and electrophoresis procedures may also be carried out inside an anaerobic chamber, using reagents that were prepared with degassed H2O and/or equilibrated inside the anaerobic chamber. The addition of the aconitase substrate citrate (or isocitrate) to the extraction and electrophoresis buffers protects the Fe-S cluster and markedly reduces the loss of aconitase activity.
Try to keep extract protein concentration high, e.g., 20–40 μg/μL, to keep loading volume to a minimum and to minimize the amount of salt and detergent in the sample loaded.
Harvest tissues, cut into small pieces, and flash-freeze on dry ice. Store at -80 °C until use.
For homogenization using the Bullet Blender, chill microcentrifuge tubes containing ~100 μL of homogenization beads on dry ice.
Note: For homogenization using the Bullet Blender, microcentrifuge tubes that can be closed tightly are strongly recommended (e.g., high-quality snap-cap or screw-cap tubes).
Weight out 20–40 mg of tissue pieces in the microcentrifuge tubes containing the homogenization beads.
Add 5 μL of cold RIPA-citrate extract buffer per milligram of tissue to each tube, e.g., 100 μL of extraction buffer for 20 mg of tissue.
For mouse liver and heart samples, run the Bullet Blender for 2 min at speed 8 at 4 °C. Wait for 2 min. Repeat once.
Note: Homogenization times, speeds, and beads may need to be adjusted if working with different tissues. Follow the manufacturer’s instructions.
Centrifuge at 2,000× g for 2 min in a refrigerated microcentrifuge. Transfer the supernatant into chilled microcentrifuge tubes. Centrifuge at maximum speed (21,000× g) for 10 min at 4 °C. Transfer the supernatant (tissue RIPA extract) to chilled microcentrifuge tubes. Use immediately or store at -80 °C until use.
Notes:
Repeated sample freeze-thaw can decrease enzymatic activity. Aliquot extracts if needed.
Some tissue extracts, e.g., liver, have high lipid content, which can cause band distortion in the in-gel assay. Carefully transfer the lower layer of the supernatant (the red aqueous phase containing the protein extract) into a clean, chilled microcentrifuge tube using a 200 μL pipette tip. Avoid pipetting the top white lipid layer. A gel loading tip can be substituted. Repeat centrifugation to remove more lipids. Carefully transfer the lower layer of the supernatant into a clean chilled microcentrifuge tube.
Proceed with protein quantification.
Note: We use Bradford Protein Assay from Bio-Rad and follow the manufacturers’ protocol. Other protein quantification methods may be used. For instance, a BCA detection kit can be used for extracts made without DTT.
Non-denaturing polyacrylamide gel electrophoresis
Notes:
We use a mini-Protean II PAGE system from Bio-Rad and gel spacers of 1.0 mm thickness.
Keep samples at 0–4 °C to minimize loss of enzymatic activity.
The electrophoresis should be run at a low temperature (0–4 °C) to minimize loss of enzymatic activities. Chill 1 L of H2O for making 1× running buffer more than 12 h before the experiment. Use running buffer at 0–4 °C. Chill down the electrophoresis cell before and during the electrophoresis. Run the electrophoresis with the electrophoresis cell in an ice bath or in a cold room.
Prepare 1× running buffer (see Recipes) and keep it cold (0–4 °C).
Cast the separating gel (see Recipes). Overlay the separating gel with 0.5 mL of isopropyl alcohol to exclude oxygen from the gel surface during polymerization and ensure an even interface between the separating and stacking gels. Allow polymerization to occur for 15 min at RT. After 15 min, rinse out the alcohol and remove residual H2O with a blotting paper.
Notes:
Prepare fresh 10% APS shortly prior to use.
Although visible gelation occurs in 15–20 min, polymerization continues for much longer (https://www.bio-rad.com/webroot/web/pdf/lsr/literature/Bulletin_1156.pdf). Allow >90 min for total separating gel polymerization time before running electrophoresis for best results.
For diluted samples, cast a shorter separating gel such that the height of the stacking gel is at least 2× the height of the sample in the well, which ensures band sharpness for large loading volumes (https://www.bio-rad.com/webroot/web/pdf/lsr/literature/Bulletin_6201.pdf).
Cast the stacking gel (see Recipes) and insert the well-forming comb without trapping air under the teeth. Allow polymerization to occur for 15 min at RT.
Assemble the electrophoresis cell.
Remove the comb from the gel, rinse wells with 1× running buffer, and assemble the electrophoresis cell.
Fill the inner and outer buffer chambers with cold running buffer. Allow gel polymerization and cooling to continue in an ice bath or in a cold room at 4 °C for 1 h.
Prepare loading samples: Mix the protein extract with the appropriate amount of the loading buffer and deionized H2O to reach 12–20 μL of final volume for a 12-well 1.0 mm thick gel. For example, to load 80 μg of protein per lane, add 4 μL of 20 µg protein/μL extract to 3 μL of 4× loading buffer and 5 μL of H2O.
Notes:
Do not boil the samples. Boiling results in irreversible loss of enzymatic activity.
The amount of total protein extract needed per lane could be variable depending on aconitase expression and activity levels. For human muscle biopsies and mouse liver extracts, 5–20 μg of protein per lane is a good starting point. For human or mouse cell lines cultured at 21% O2 atmosphere, 50–100 µg of protein per lane is a good starting point.
For best resolution, load a smaller volume rather than a larger volume. Using 12–16 µL of loading samples per lane in a 10- or 12-well mini-gel is a good starting point. However, if the extracted protein concentration is <20 μg/μL, to avoid high salt concentration in the loading samples, increase the loading volume and use a gel with a taller stacking gel.
Load samples slowly to allow them to settle evenly on the bottom of the wells.
Run electrophoresis at constant 170 V at 4 °C for 2.5–4 h for a mini-gel (8.3 × 10 cm) and 5–6 h for a big gel (16 × 20 cm).
Notes:
The electrophoresis running time may vary with different electrophoresis cells. For pilot experiments, use Invitrogen SeeBlueTM Plus2 Pre-Stained Standard as a guide. Running electrophoresis until the top two blue bands are ~2 cm apart is a good starting point for human and mouse aconitase in-gel assays.
Optional: Run a parallel SDS-PAGE gel with the same samples. After running the gel, perform immunoblot analysis using anti-aconitase and anti-tubulin antibodies or Coomassie staining to check protein loading.
In-gel aconitase activity staining
Notes:
The MTT reagent is sensitive to light. Avoid extended exposure of MTT to direct light.
IDH is sensitive to heat. Add IDH to the reaction mix immediately before staining the gel.
Prewarm the H2O for the aconitase activity assay mix: put the appropriate amount of H2O in a light-protected container (e.g., a tube wrapped in aluminum foil) and put the tube in a 37 °C water bath or incubator for >10 min.
Immediately before removing the gel from the glass plates, add all the assay reagents, except IDH, to the prewarmed H2O.
Remove the gel carefully from between the glass plates and place the gel in a clean tray (e.g., recycled pipette tip box).
Add IDH to the aconitase activity assay mix and add to the tray containing the gel. Cover with foil. Incubate with gentle agitation at RT or in a 37 °C incubator for 5–60 min.
Note: The staining time could be variable depending on aconitase expression level and activity. For human muscle biopsies and mouse liver extracts, strong signals may be visible in 5–15 min. For human or mouse cell cultures, incubate for 15–60 min.
Gel washing: Wash gel 5–10 times (5–10 min each) with H2O, with gentle agitation and protection from direct light.
Note: The total washing time could be variable depending on the staining time needed for a good signal of the aconitase activity. The longer the staining time, the longer the washing time needed to remove background staining. However, a very long washing time may result in weaker or less well-defined aconitase activity bands.
Gel drying: Dry gel between a filter paper and a plastic wrap using a gel vacuum dryer or air-drying methods. Take pictures or scan (>400 dpi) before (in case the gel cracks during drying) and after gel drying.
Comparison and quantification of aconitase activities can be done by densitometric analysis using open-source software following manufacturer’s instruction (e.g., ImageJ, https://imagej.net/ij/docs/guide/user-guide.pdf).
Validation of protocol
This protocol has been shown to be robust and reproducible in detecting changes in mammalian aconitase activities in response to oxidative or nitrative stress, iron depletion, and various pathophysiological conditions in human cell line HeLa S3 [1,4], human muscle biopsies [5], murine macrophage cell line RAW264.7 [6], human RAW264.7-mouse Caco2 co-culture [7], mouse tissues [8], etc. Statistical analyses in these studies were performed by Student's t-test for paired samples. All data are representative of three or more independent experiments. Figures 1 and 2 show some examples of in-gel activity assays for human and mouse aconitases, respectively. The assignment of bands corresponding to ACO1 and ACO2 was confirmed using subcellular fractions and extracts from ACO1 or ACO2 knockdown cells or knockout animals.
Figure 1. In-gel activity assays for human mitochondrial and cytosolic aconitases. Non-denaturing PAGE of HeLa S3 cell Triton extracts and subcellular fractions at constant 170 V at 4 °C for 2–2.5 h followed by gel staining with a coupled enzyme assay mix at 37 °C. (A) Iron deficiency, induced by treatment with the iron chelator Desferal (Dfo), lowers both ACO1 and ACO2 activities. Note: The aconitase bands run between the top two SeeBlue protein markers. (B) Aconitase in-gel assay of HeLa S3 cells transfected with non-targeting (NT) compared to ACO2 siRNA. 80 µg of protein were loaded in each lane in (A) and (B). (C) Aconitase in-gel assay of four separate HeLa S3 cell cultures demonstrated reproducibility of the assay. (D) Aconitase in-gel assay of HeLa S3 Triton extracts containing 128, 64, and 32 μg of protein, compared to a HeLa S3 cytosolic fraction. (E) Quantitative analysis of aconitase activity of the Triton extracts in (D).
Figure 2. In-gel aconitase activity assays for mouse tissue or cell line extracts. Because of the difference in pKa values of the mouse and human aconitases, gels with 5%–6% acrylamide at a higher pH and longer running time (3–4 h) are needed for better resolution of mouse ACO1 and ACO2. (A) Comparison of aconitases in mouse liver extracts and mouse 3T3 L1 cell cytosolic (Cyt) and mitochondrial (Mito) fractions with human aconitases in HeLa cell lysates. Note: In wild-type (WT) mouse liver extracts, two mitochondrial aconitase isoforms and two cytosolic aconitase isoforms were detected, representing various states of phosphorylation and/or post-translational modification. In IRP1-/- mouse liver extracts, only the ACO2 isoforms were detected. (B) Aconitase in-gel assay of mouse liver (L) and heart (H) using a freshly prepared gel according to the current protocol showed a stronger signal and better resolution compared to a commercial pre-cast Tris-Glycine gel. Courtesy of Dr Suh Young Jeong. This panel also shows that different tissues have different relative amounts of ACO1 and ACO2 activities. For example, mouse heart samples have predominantly ACO2 activity.
General notes and troubleshooting
Advantages and limitations
The main advantage of the in-gel PAGE-based aconitase assay over the classical UV/vis spectrophotometric coupled enzyme assay [2] and commercial aconitase assay kits is that the in-gel assay allows simultaneous monitoring of mitochondrial and cytosolic aconitase activities without the need for the fractionation procedure, which is time-consuming and might result in loss of the oxygen-sensitive Fe-S cluster. However, in our experience, the in-gel assay is ~10× less sensitive than the UV/vis spectrophotometric coupled enzyme assay. In general, this is not an issue for our experiments using HeLa cells or mouse tissues, since we routinely obtained protein extracts at 20–40 μg protein/μL from a 100 mm plate of HeLa cells or a 15 mg piece of mouse liver, heart, or brain, which is enough for the in-gel aconitase assays and western blot controls. That said, for some cell lines, such as Hep3B, it might be more difficult to obtain protein extracts at 20 μg protein/μL from a 100 mm plate of cells (personal communication). Users who have a very limited amount of samples may consider using the UV/vis spectrophotometric coupled enzyme assay to measure total (mitochondrial and cytosolic) or subcellular fraction aconitase activities.
Aconitase in-gel assay using a gel freshly prepared according to the current protocol showed a stronger signal and better resolution compared to a commercial pre-cast Tris-Glycine gel (Figure 2B). One likely factor is the absence of citrate in the commercial pre-cast gel. Another factor is the buffer in the gels. We have made gels using Tris-glycine-citrate buffer for comparison and found that gels made with Tris-borate-citrate buffer gave sharper bands.
Troubleshooting
Problem Options
Bands appear smeared or distorted Incomplete gel polymerization. Allow the gel to polymerize for a longer time. Use freshly prepared APS. Use new TEMED.
Detergent or gel residues on glass gel plates. Clean and rinse gel plates thoroughly. Optional: Wipe plates with alcohol.
Gel too warm. Keep the gel and running buffer well-chilled before and during running.
High salt concentration in the loading sample.
• Keep extract protein concentration high, e.g., 20–40 μg/μL, and the volume of lysate less than 40% of the final loading volume.
• For more diluted protein extract (<20 μg/μL), increase the loading volume (e.g., 20 μL for a 12-well 1.0 mm thick gel).
• For higher loading volume, increase the height of the stacking gel so that the height of the stacking gel is 2× the height of the sample in the well.
High lipid content (e.g., in liver samples). Remove the protein extract from under the lipid layer carefully. Repeat centrifugation if necessary.
Poor resolution Reduce the loading volume. Use more concentrated lysates.
Run electrophoresis for a longer time.
Gel ran too fast. Current too high. Decrease voltage by 25%.
Aconitase activity bands weak or not detected Keep samples cold (0–4 °C) and run electrophoresis at a low temperature to minimize loss of enzymatic activities.
Use more IDH or a new batch of IDH and longer staining time.
For tissue samples or cells cultured at low O2 atmosphere, reduce exposure of samples to O2. Degas (e.g., https://assets.thermofisher.com/TFS-Assets/LSG/Application-Notes/TR0029-Degas-buffers.pdf) or deoxygenate (e.g., equilibrating H2O or buffers inside an anaerobic hood or passing N2 gas through H2O or buffers using a gas dispersion tube) buffers. Run electrophoresis in an anaerobic chamber following the manufacturer’s instructions.
Include a positive control, e.g., HeLa cells cultured in iron-rich media or mouse liver samples.
Acknowledgments
We thank Dr. Esther Meyron-Holtz and Dr Suh Young Jeong for their valuable feedback in the testing of this protocol. This work was supported by the intramural program of the National Institute of Child Health and Human Development. The protocols were adapted from the research article of Tong et al. [4].
Competing interests
The author declares no conflicts of interest.
Ethical considerations
All experimental procedures including animal subjects were reviewed and approved by the National Institute of Child Health and Human Development Animal Care and Use Committee and met NIH guidelines for the humane care of animals.
References
Tong, W. H. and Rouault, T. A. (2007). Metabolic regulation of citrate and iron by aconitases: role of iron–sulfur cluster biogenesis. BioMetals. 20549–564.
Rose, I. A. and O'Connell, E. L. (1967). Mechanism of Aconitase Action. J Biol Chem. 242(8): 1870–1879.
Missirlis, F., Hu, J., Kirby, K., Hilliker, A. J., Rouault, T. A. and Phillips, J. P. (2003). Compartment-specific Protection of Iron-Sulfur Proteins by Superoxide Dismutase. J Biol Chem. 278(48): 47365–47369.
Tong, W. H. and Rouault, T. A. (2006). Functions of mitochondrial ISCU and cytosolic ISCU in mammalian iron-sulfur cluster biogenesis and iron homeostasis. Cell Metab. 3(3): 199–210.
Mochel, F., Knight, M. A., Tong, W. H., Hernandez, D., Ayyad, K., Taivassalo, T., Andersen, P. M., Singleton, A., Rouault, T. A. and Fischbeck, K. H. (2008). Splice Mutation in the Iron-Sulfur Cluster Scaffold Protein ISCU Causes Myopathy with Exercise Intolerance. Am J Hum Genet. 82(3): 652–660.
Tong, W. H., Maio, N., Zhang, D. L., Palmieri, E. M., Ollivierre, H., Ghosh, M. C., McVicar, D. W. and Rouault, T. A. (2018). TLR-activated repression of Fe-S cluster biogenesis drives a metabolic shift and alters histone and tubulin acetylation. Blood Adv. 2(10): 1146–1156.
Fahoum, L., Moshe-Belisowski, S., Zaydel, K., Ghatpande, N., Guttmann-Raviv, N., Zhang, W., Li, K., Tong, W. H., Nyska, A. and Waterman, M. (2024). Iron regulatory protein 1 is required for the propagation of inflammation in inflammatory bowel disease. J Biol Chem. 300(9): 107639.
Jeong, S. Y., Hogarth, P., Placzek, A., Gregory, A. M., Fox, R., Zhen, D., Hamada, J., van der Zwaag, M., Lambrechts, R. and Jin, H. (2019). 4'‐Phosphopantetheine corrects CoA, iron, and dopamine metabolic defects in mammalian models of PKAN. EMBO Mol Med. 11(12): e201910489.
Article Information
Publication history
Received: Jun 20, 2024
Accepted: Sep 25, 2024
Available online: Oct 17, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Molecular Biology > Protein > Activity
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The authors used this protocol in Plant Physiology Dec 2023
Abstract
Gene expression analysis is a fundamental technique to elucidate the regulatory mechanisms of genes of interest or to reveal the patterns of plant response to environmental stimuli. Traditionally, gene expression analyses have required RNA extraction, followed by cDNA synthesis and qPCR analyses. However, this conventional method is costly and time-consuming, limiting the amount of data collected. The protocol outlined in this study, which utilizes a chemiluminescence system, offers a cost-effective and rapid method for assessing the expression of Arabidopsis (Arabidopsis thaliana) genes, exemplified by analyzing the nitrate-inducible expression of a major nitrate transporter gene, nitrate transporter 2.1 (NRT2.1). A reporter construct, containing the NRT2.1 promoter fused to the firefly luciferase gene, was introduced into wild-type and mutant Arabidopsis plants. Seeds obtained from the transgenic lines were grown for 3 days in 96-well microplates containing a nitrate-free nutrient solution. After 3 days, the nutrient solution was replaced with a fresh batch, which was supplemented with luciferin potassium. One hour later, nitrate was added at various concentrations, and the temporal expression pattern of NRT2.1 was analyzed by monitoring the chemiluminescence signals. This method allowed for the cost-effective, quantitative, and high-throughput analysis of NRT2.1 expression over time under the effects of various nutrient conditions and genetic backgrounds.
Key features
• Small-scale and immediate assessment of NRT2.1 promoter activity using 3-day-old Arabidopsis seedlings expressing the firefly luciferase gene under the control of the Arabidopsis NRT2.1 promoter.
• Comparison of various Arabidopsis genotypes and nutrient conditions using 96-well microplates.
• Quantitative assessment of the temporal changes in gene expression levels.
Keywords: Arabidopsis thaliana Nitrate Gene expression Luciferase Microplate Seedling Temporal expression pattern
Graphical overview
Graphical summary of the microplate-based NRT2.1 expression monitoring system in planta.
Note: The steps within gray square brackets are part of a general protocol and are not included in this manuscript.
Background
Gene expression analysis is a fundamental approach used to understand the plant response to environmental stimuli and examine the effects of gene manipulation. The current method of gene expression analysis typically involves RNA extraction, cDNA synthesis, and qPCR, which often limits the number of samples that can be handled at one time. In addition, RNA extraction and purification per se typically cost >$3 per sample, potentially hindering the analysis of a large number of samples. Consequently, many laboratories perform gene expression analyses only at a few time points, which may obscure the investigation of important genes that exhibit time-dependent expression patterns owing to the effects of multiple transcriptional regulators. In such cases, analysis of gene expression at several time points is crucial for elucidating the mechanisms employed by plants to fine-tune gene expression and for understanding the role of each transcriptional regulator in this process.
Arabidopsis nitrate transporter 2.1 gene (NRT2.1), which encodes a major high-affinity nitrate transporter required for nitrate uptake by the root [1], is an example of a gene regulated by multiple transcriptional regulators. NRT2.1 expression is under the control of multiple transcription factors with antagonistic functions, including NIN-like protein (NLP) transcription factors, which act as nitrate-activated transcriptional enhancers, and nitrate-inducible GARP-type transcriptional repressor1 (NIGT1) proteins, which act as NLP-inducible transcriptional repressors [2] (Figure 1A). NRT2.1 shows a bell-shaped expression pattern upon nitrate supply [2,3], owing to its initial NLP-induced upregulation [4], followed by NIGT1-mediated repression. To understand the contribution of different transcription factors to NRT2.1 expression, it is crucial to analyze the temporal expression pattern of NRT2.1 in different genetic backgrounds, which necessitates the analysis of NRT2.1 expression in numerous samples.
Reporter proteins, such as green fluorescent protein (GFP) and luciferase (LUC), are versatile biological tools that facilitate the analysis of protein localization or promoter activity without laborious procedures. Firefly-derived LUC gene is often used as a reporter gene to determine promoter activity in plant and animal cells because of its high sensitivity and linear range. To analyze the effects of transcription factors on target promoters, LUC is often expressed under the control of a given promoter in the analysis of plant genes in a protoplast-based assay system [5]. LUC has also been used in planta to analyze the temporal expression patterns of genes, such as those related to the circadian rhythm [6,7]. The abundance of LUC protein increases upon activation of the promoter regulating LUC expression. LUC protein converts its substrate, luciferin, to oxyluciferin with the help of a magnesium ion, which emits light. As long as the substrates (luciferin, ATP, and oxygen) are not depleted, the intensity of emitted light is assumed to be proportional to the amount of LUC protein, which enables the assessment of promoter activity (Figure 1B).
This protocol describes a detailed, step-by-step method optimized for a microplate-based LUC reporter assay system, which was used to evaluate the temporal activity of the NRT2.1 promoter. This rapid, cost-effective, and high-throughput method holds potential for analyzing the expression of various genes.
Figure 1. Regulatory mechanism of Arabidopsis NRT2.1 and the principle of luciferase (LUC) assay. (A) The known regulatory mechanism of NRT2.1 in Arabidopsis. Nitrate directly binds to and activates NLP. Activated NLP (NLP*) directly binds to the promoters of NRT2.1 and NIGT1, enhancing their expression. Subsequently, NIGT1 suppresses the expression of not only NRT2.1 but also NIGT1 genes. (B) Mode of action of LUC. The level of LUC protein increases upon the activation of NRT2.1 promoter (NRT2.1pro). LUC protein catalyzes the conversion of luciferin to an excited state (oxyluciferin) with the aid of Mg2+. Subsequently, light is emitted during the transition of oxyluciferin to the ground state.
Materials and reagents
Biological materials
Transgenic Arabidopsis thaliana seeds harboring the LUC gene under the control of NRT2.1 promoter [2]
Reagents
Sodium hypochlorite (FUJIFILM Wako Pure Chemical, catalog number: 197-02206)
D-luciferin potassium salt (FUJIFILM Wako Pure Chemical, catalog number: 120-05114)
Potassium nitrate (FUJIFILM Wako Pure Chemical, catalog number: 160-04035)
Ammonium succinate (Kanto Chemical, catalog number: 01319-30)
Murashige and Skoog salts without nitrogen, phosphorus, and iron (FUJIFILM Wako Pure Chemical, special order)
2-Morpholinoethanesulfonic acid (MES) (Dojindo, catalog number: 343-01626)
Potassium hydroxide (FUJIFILM Wako Pure Chemical, catalog number: 168-21815)
Potassium dihydrogen phosphate (FUJIFILM Wako Pure Chemical, catalog number: 169-02425)
Iron (II) sulfate heptahydrate (FUJIFILM Wako Pure Chemical, catalog number: 094-01082)
Sucrose (FUJIFILM Wako Pure Chemical, catalog number: 196-00015)
Solutions
MES buffer (see Recipes)
10 mM iron (II) sulfate solution (see Recipes)
1 M potassium dihydrogen phosphate solution (see Recipes)
100 mM ammonium succinate solution (see Recipes)
1× Murashige and Skoog solution (N-, P-, Fe-free) (see Recipes)
0.1× N-free MS medium (see Recipes)
1 M potassium nitrate stock solution (see Recipes)
10 mM D-luciferin potassium stock solution (see Recipes)
Recipes
170 mM MES buffer
Note: Adjust the pH of 170 mM MES buffer to 5.7 using potassium hydroxide solution. Filter-sterilize the buffer and store at room temperature.
Reagent Final concentration Quantity or Volume
2-Morpholinoethanesulfonic acid 170 mM 3.6 g
Potassium hydroxide n/a n/a
Ultrapure water (Milli-Q) n/a Up to 100 mL
Total n/a 100 mL
10 mM iron (II) sulfate stock solution (10 mL)
Note: Filter the solution, aliquot into 1.5 mL tubes, and store at -20 °C to prevent precipitation.
Reagent Final concentration Quantity or Volume
Iron (II) sulfate heptahydrate 10 mM 27.8 mg
Ultrapure water (Milli-Q) n/a Up to 10 mL
Total n/a 10 mL
1 M potassium dihydrogen phosphate stock solution (10 mL)
Note: Autoclave or filter-sterilize the solution and store at room temperature.
Reagent Final concentration Quantity or Volume
Potassium dihydrogen phosphate 1 M 1.36 g
Ultrapure water (Milli-Q) n/a Up to 10 mL
Total n/a 10 mL
100 mM ammonium succinate stock solution (10 mL)
Note: Autoclave or filter-sterilize the solution and store at room temperature.
Reagent Final concentration Quantity or Volume
Ammonium succinate 100 mM 0.15 g
Ultrapure water (Milli-Q) n/a Up to 10 mL
Total n/a 10 mL
1× Murashige and Skoog medium (without N, P, and Fe) (1 L)
Note: To conduct studies involving nutrient-deficiency treatments, make a specialized Murashige and Skoog plant salt mixture lacking KNO3, NH4NO3, KH2PO4, and FeSO4, and store the solution at 4 °C.
Reagent Final concentration Quantity or Volume
Murashige and Skoog plant salt mixture (N-, P-, and Fe-free) 1× 1 bag
Ultrapure water (Milli-Q) n/a Up to 1 L
Total n/a 1 L
0.1× N-free MS medium + 500 µM ammonium succinate (100 mL)
Note: Mix all components, except iron (II) sulfate solution, and autoclave. After autoclaving, add the iron (II) sulfate solution and mix thoroughly. Store the solution at 4 °C.
Reagent Final concentration Quantity or Volume
1× Murashige and Skoog medium (without N, P, and Fe) 0.1× 10 mL
Sucrose 0.5% (w/v) 0.5 g
100 mM ammonium succinate stock solution 500 µM 0.5 mL
1 M potassium dihydrogen phosphate stock solution 125 µM 12.5 µL
170 mM MES stock solution 3.5 mM 2.1 mL
10 mM iron (II) sulfate stock solution (add after autoclave) 10 µM 100 µL
Ultrapure water (Milli-Q) n/a Up to 100 mL
Total n/a 100 mL
1 M potassium nitrate stock solution (10 mL)
Note: Autoclave or filter-sterilize the solution and store at room temperature.
Reagent Final concentration Quantity or Volume
Potassium nitrate 1 M 1.0 g
Ultrapure water (Milli-Q) n/a Up to 10 mL
Total n/a 10 mL
10 mM luciferin potassium stock solution (10 mL)
Note: Aliquot the solution into 1.5 mL tubes and store at -80 °C.
Reagent Final concentration Quantity or Volume
Luciferin potassium 10 mM 31.8 mg
Ultrapure water (Milli-Q) n/a Up to 10 mL
Total n/a 10 mL
Laboratory supplies
Black 96-well microplates (Greiner, F-bottom, catalog number: 655077)
Transparent lids for 96-well plates (e.g., TPP, catalog number: 92696)
1.5 mL tubes (e.g., Watson, catalog number: 131-415C)
200 µL pipette tips (e.g., Watson, catalog number: 110-705C)
Surgical tape (e.g., 3M Company, catalog number: 1530-0)
Beaker
Equipment
Microplate reader (Tecan, model: Infinite M1000)
Electronic balance
Autoclave
Tabletop centrifuge
pH meter
Magnetic stirrer
Clean bench
Growth chamber (equipped with LED lights)
Pipettes (200 µL, single- and 8-channel)
Software and datasets
Magellan software (Tecan)
Microsoft Excel (Microsoft)
Procedure
Seed preparation
Place transgenic Arabidopsis seeds in a 1.5 mL tube.
Sterilize the seeds with 0.7% sodium hypochlorite solution for 5 min at room temperature.
Mix 860 µL of sterile deionized water and 140 µL of 5% sodium hypochlorite in a 1.5 mL tube.
Vigorously mix the contents of the tube using a vortex mixer.
Centrifugate briefly (~5 s) at 2,000× g and remove the supernatant with a pipette.
Rinse the seeds with 1 mL of sterile deionized water, vortex, and remove the supernatant after a brief centrifugation (~5 s at 2,000× g).
Repeat step A2d 4–5 times.
Stratify the seeds by placing them in a refrigerator (4 °C), along with a small amount of water, enough to cover the seeds, for 2–3 days.
Note: The water and pipette tips used above must be autoclaved or purchased sterile to ensure that plants stay free from infection during growth.
Seed sowing and plant growth
Add 200 µL of nutrient solution (0.1× N-free MS medium + 500 µM ammonium succinate) to each well of a 96-well microplate.
Note: Black microplates are recommended as white microplates may produce higher background signals.
Fill the gap between wells with sterile deionized water to prevent evaporation of the nutrient solution.
Place four seeds in each well using a pipette tip.
Note: To ensure that plants stay free from infection during growth, the above steps must be performed on a clean bench.
Cover the microplate with a transparent lid and seal the lid with surgical tape.
Incubate the plate under continuous light (60 µE; around 30 cm from the light source in our case) in a growth chamber maintained at 23 °C for 3 days.
To apply the phosphorus deficiency treatment, remove the nutrient solution from the wells 15 h before measurement of LUC activity; wash the seedlings with 200 µL of 0.1× N- and P-free MS medium (+500 µM ammonium succinate) once; and add 200 µL of fresh 0.1× N-free MS medium (+500 µM ammonium succinate) containing a reduced amount of phosphorus (0 or 5 µM compared with 125 µM in the control solution).
Note: Three-day-old plants were found to be optimal, 2-day-old plants produced relatively lower signal intensity, and 4-day-old plants were too tall and sometimes exceeded the well depth.
Treatment
Uncover the plate 1 h before treatment onset and remove the solution using an 8-channel pipette.
Note: Ensure that the seedlings are not mechanically damaged by the pipette tip.
Add a new nutrient solution (200 µL) containing 100 µM of luciferin potassium to each well using a single-channel or an 8-channel pipette.
In the phosphorus deficiency treatment, the phosphorus concentration in the treatment solution should remain as low as in the growth solution (0 or 5 µM compared with 125 µM in the control solution).
Cover the plate with a lid and incubate under light for 1 h in the same growth chamber set at 23 °C with continuous illumination.
Add potassium nitrate at a final concentration of 0–10 mM to each well and mix the contents of each well by pipetting with an 8-channel pipette (at least 5 times). We typically add 2 µL of potassium nitrate solution concentrated 100-fold (e.g., 2 µL of 500 mM potassium nitrate for a final concentration of 5 mM).
Note: The inclusion of appropriate controls, such as non-transgenic plants or nitrate-free medium, is recommended to account for background signals.
Measurement
Immediately after adding potassium nitrate, transfer the plate to the microplate reader.
Record the chemiluminescence as described below:
Shake the plate for 2 s with the linear and 3 mm settings of the instrument.
Read the chemiluminescence of each well for 1.5 s.
Notes:
i. The reading duration can be adjusted according to signal strength.
ii. No wavelength filter was applied during the luminescence measurement.
After recording the signal from all wells, transfer the plate back to the incubator set at 23 °C with continuous illumination.
Note: This step is necessary to maintain signal intensity over time, likely because the emission of chemiluminescence by LUC requires ATP as a substrate (Figure 1B).
Perform steps D2 and D3 at regular intervals (e.g., at 20-min intervals for a total of 360 min).
Data analysis
Export the raw values (luminescence units) to Microsoft Excel and open the data.
Note: We typically prepare eight wells (n = 8) for each sample group and calculate the mean value. Plot the mean luminescence units against time (Figure 2). If the values are expressed on a relative scale (e.g., 0–1 scale), consider 0 and 1 as the lowest and highest values, respectively, with other values as intermediate.
Figure 2. Summary of data analysis. Raw data files containing luminescence signal values collected at different time points are merged, and a summary table is prepared. A mean value is calculated for each time point and nitrate condition. A graph is subsequently constructed based on the data.
Validation of protocol
This protocol, or parts of it, has been used and validated in the following research article:
Ueda and Yanagisawa [3]. Transcription factor module NLP–NIGT1 fine-tunes NITRATE TRANSPORTER2.1 expression. Plant Physiology (Figure 1, panel B, D; Figure 2; Figure 6, panel A, B).
General notes and troubleshooting
General notes
Effects of different genetic backgrounds and nutrient conditions
This method can be used to analyze the effect of nutrient signaling-related genes and nutrient conditions on gene expression. For example, in our previous study, the NRT2.1pro-LUC transgenic line was crossed with knockout mutants (such as nlp6nlp7, phr1phl1, and nigt1.1/1.2/1.3/1.4) defective in nutrient signaling. Then, plants homozygous for all intended loci were selected and analyzed [3]. The temporal expression pattern of NRT2.1 was compared between the wild-type and mutants, enabling a quantitative evaluation of the contribution of each transcription factor. Additionally, we performed a time-course analysis of the effect of phosphorus deficiency on NRT2.1 expression [3].
Applications in semi-quantitative imaging analysis
The promoter-LUC transgenic lines can also be used for semi-quantitative imaging analysis. Briefly, plants grown on agar plates or in liquid culture under different nutrient conditions are supplied with luciferin potassium, and chemiluminescence is detected with a CCD imager. This analysis has been adopted in our previous studies [2,8].
Potential delay in chemiluminescence detection
We note that the measurement of LUC activity does not account for the abundance of mRNA per se. Since the synthesis of LUC protein and decay of LUC activity do not necessarily coincide with those of NRT2.1 mRNA, there could be a slight discrepancy between LUC activity and NRT2.1 expression. In our previous mathematical modeling study, we explicitly incorporated the process of LUC activity decay (half-life 15.3 min [9]) in the formula, which resulted in a slight difference between the timing of peak LUC activity and that of maximum NRT2.1 expression [3]. Nevertheless, the correlation coefficient calculated using LUC signal intensity and experimentally determined NRT2.1 expression level was >0.95, indicating that the LUC signal could be used as a first-order approximation of NRT2.1 promoter activity under different environmental conditions or at different time points.
Applications in other organisms
A similar microplate-based quantitative LUC assay could likely be established for other plant species, provided the following conditions are met: (1) seeds and seedlings of the plant species fit easily within the microplate wells; and (2) growth conditions, including culture duration and the number of seeds per well, are optimized for the species of interest.
Troubleshooting
Problem 1: Signal intensities are too low.
Possible cause: Weak promoter activity or small plant size.
Solution: Try using a different promoter or larger plants and/or increase the chemiluminescence reading time.
Acknowledgments
This study was partly supported by Japan Society for the Promotion of Science (KAKENHI, grant no. 22H04977). This protocol is based on our recent publication [3].
Competing interests
The authors declare that they have no competing interests.
References
Okamoto, M., Vidmar, J. J. and Glass, A. D. M. (2003). Regulation of NRT1 and NRT2 Gene Families of Arabidopsis thaliana: Responses to Nitrate Provision. Plant Cell Physiol. 44(3): 304–317.
Maeda, Y., Konishi, M., Kiba, T., Sakuraba, Y., Sawaki, N., Kurai, T., Ueda, Y., Sakakibara, H. and Yanagisawa, S. (2018). A NIGT1-centred transcriptional cascade regulates nitrate signalling and incorporates phosphorus starvation signals in Arabidopsis. Nat Commun. 9(1): 1376.
Ueda, Y. and Yanagisawa, S. (2023). Transcription factor module NLP–NIGT1 fine-tunes NITRATE TRANSPORTER2.1 expression. Plant Physiol. 193(4): 2865–2879.
Liu, K. H., Liu, M., Lin, Z., Wang, Z. F., Chen, B., Liu, C., Guo, A., Konishi, M., Yanagisawa, S., Wagner, G., et al. (2022). NIN-like protein 7 transcription factor is a plant nitrate sensor. Science. 377(6613): 1419–1425.
Yoo, S. D., Cho, Y. H. and Sheen, J. (2007). Arabidopsis mesophyll protoplasts: a versatile cell system for transient gene expression analysis. Nat Protoc. 2(7): 1565–1572.
Urquiza-García, U. and Millar, A. J. (2019). Expanding the bioluminescent reporter toolkit for plant science with NanoLUC. Plant Methods. 15(1): 68.
Dixon, L. E., Hodge, S. K., Ooijen, G., Troein, C., Akman, O. E. and Millar, A. J. (2014). Light and circadian regulation of clock components aids flexible responses to environmental signals. New Phytol. 203(2): 568–577.
Ueda, Y., Kiba, T. and Yanagisawa, S. (2020). Nitrate‐inducible NIGT1 proteins modulate phosphate uptake and starvation signalling via transcriptional regulation of SPX genes. Plant J. 102(3): 448–466.
Van Leeuwen, W., Hagendoorn, M. J. M., Ruttink, T., Van Poecke, R., Van Der Plas, L. H. W. and Van Der Krol, A. R. (2000). The use of the luciferase reporter system for in planta gene expression studies. Plant Mol Biol Rep. 18(2): 143–144.
Article Information
Publication history
Received: Jul 11, 2024
Accepted: Oct 7, 2024
Available online: Oct 22, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Plant Science > Plant molecular biology > DNA
Molecular Biology > DNA > Gene expression
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Mouse Model of Lipopolysaccharide (LPS)-Induced Pulpitis
LS Lanting Shao
BC Baian Chen
YZ Ying Zheng
Published: Vol 15, Iss 2, Jan 20, 2025
DOI: 10.21769/BioProtoc.5128 Views: 1591
Reviewed by: Komuraiah MyakalaShivaram SelvamMinal EngavaleRaghavendra Yelahanka Nagaraja
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Abstract
Pulpitis is an important and prevalent disease within the oral cavity. Thus, animal models are necessary tools for basic research focused on pulpitis. Researchers worldwide often use dogs and miniature pigs to construct animal models of pulpitis. However, gene editing in miniature pigs is difficult, the surgical modeling process is complex, and tooth demineralization time is lengthy. Although some researchers have attempted to establish a mouse model of pulpitis, most models have involved direct exposure of dental pulp. However, the causes of pulpitis vary considerably among individuals, hindering effective research. In this study, we established a mouse model of pulpitis by accessing the pulp cavity, exposing the pulp to lipopolysaccharide (LPS), and then filling the tooth. One day after surgery, we observed many necrotic tissues and extensive inflammatory exudate, including neutrophils, around the coronal cavity preparation. Additionally, we noted many more neutrophils and a small amount of chronic inflammatory cell infiltrates at the junction between inflamed and normal tissue. These findings indicated that our model can be used to explore the early stage of pulpitis. Ten days after surgery, we observed vacuolar degeneration in some fibroblasts and proliferation in others at the distal end of the inflamed tissue. We also noted dilation and congestion of the pulp blood vessels. Therefore, our model can also be used to explore the middle and later stages of pulpitis. Thirty days after surgery, we observed necrosis in the coronal pulp cavity and upper half of the root pulp, indicating that our model can also be used to explore the end stage of pulpitis. This model is easy to establish, shows pulpitis progression in the dental pulp, exhibits a clear inflammatory phenotype, and can be readily combined with gene editing techniques. Accordingly, it is suitable for basic research focused on pulpitis and has substantial practical value.
Key features
• Lipopolysaccharide (LPS) can induce pulpitis in mice.
• The mouse model of LPS-induced pulpitis can be used in basic studies of pulpitis.
• After 1 day, the mouse model of LPS-induced pulpitis can demonstrate the main phenotypes of early-stage pulpitis.
• After 10 days, the mouse model of LPS-induced pulpitis can display the main phenotypes of middle and late stage pulpitis.
Keywords: Pulpitis Mouse model Lipopolysaccharide Inflammation Histopathology
Graphical overview
Figure 1. Graphical overview of the C57BL/6 mouse model of lipopolysaccharide (LPS)-induced pulpitis. A. Weigh the mouse. B. Anesthetize the mouse. C. Secure the mouse to the surgical pad and expose its oral cavity. D. Open the pulp chamber of the right maxillary first molar. E. Rinse the medullary foramen with 0.9% NaCl solution. Apply a small cotton ball saturated with 1 mg/mL LPS to the medullary foramen for 5 min, then cover the medullary foramen with Esthet-X flow and irradiate the site. F. Perform tissue decalcification and paraffin embedding (sample collection, decalcification, dehydration, wax embedding, and sectioning), followed by Histopathology staining, microscopy examination, image acquisition, and analysis.
Background
Although pulpitis is a common oral disease caused by anaerobic bacteria, its underlying mechanisms remain poorly understood. The construction of an animal model that simulates the processes involved in human pulpitis can aid in exploring the onset, progression, and outcomes of pulpitis. Thus far, pulpitis models have been constructed in animals such as mice, rats [1], ferrets, cats, dogs, miniature pigs, and monkeys [2]. However, because of their size, larger animals are more expensive and resource-intensive to maintain. Considering that experimental procedures in mice are relatively clear, gene editing is mainly conducted in these organisms, and the cost of mouse feed is low, there is a need to construct a mouse model of pulpitis.
Mouse molars are similar to human molars in many aspects, such as structure and cell types [3]; therefore, mouse models of pulpitis can be used to simulate the progression of human pulpitis. However, mouse molars are small, require skilled experimental techniques, and are difficult to manipulate; as such, mouse models of pulpitis have not been widely used. Most existing mouse models of pulpitis are induced by extended exposure of dental pulp after the pulp chamber has been opened [4–6]. This surgical method does not allow identification of the bacterial species causing pulpitis and is easily influenced by various biological factors, such as stimulant components in the feed and drink, or bacteria introduced into the oral cavity after surgery. In this study, we used lipopolysaccharide (LPS), an inflammatory factor often produced by anaerobic bacteria, as the pulpitis-inducing factor. After model induction, we capped the pulp to exclude interference from oral bacteria.
Materials and reagents
Biological materials
C57BL/6JCnc (B6J) mice (Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing)
Eight-week-old C57BL/6 mice, weighing 25–30 g, were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. and maintained in a pathogen-free environment within Capital Medical University Animal Experiment Center. All animal experiments were approved by the ethics committee of Capital Medical University (AEEI-2024-071). Mice were maintained on a 12:12 h light/dark cycle and given food and water ad libitum. C57BL/6 mice, among the most popular inbred mouse strains in many research laboratories worldwide, were used in the experiments described below.
Reagents
LPS-Pg (InvivoGen, catalog number: tIrl-pglps)
Endotoxin-free water (InvivoGen, catalog number: h2olal-1.5)
Esthet-X flow (DENTSPLY DeTrey GmbH, catalog number: 005-SZ2648021)
75% ethyl alcohol (ANNJET, catalog number: Q/371402AAJ008)
Tribromoethanol (Sigma-Aldrich, catalog number: 75-80-9)
NaCl (Hong Kong JiSiEnBei International Trade Co., Ltd., catalog number: JS0492-1KG)
Tissue specimen fixative (Servicebio, catalog number: G1101-15ML)
Xylene (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 10023418)
Absolute ethanol (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 100092683)
Ethylenediaminetetraacetic acid (EDTA) decalcification solution (Servicebio, catalog number: G1105)
Benzyl alcohol (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 30020618)
Environmentally friendly dewaxing and clearing solution (Servicebio, catalog number: G1128-1L)
10% paraformaldehyde fixative (neutral) (Servicebio, catalog number: G1101)
H&E staining kit (Servicebio, catalog number: G1003)
Neutral gum (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 10004160)
Citric acid antigen repair solution (pH 6.0) (Servicebio, catalog number: G1202)
EDTA antigen repair solution (pH 9.0) (Servicebio, catalog number: G1203)
EDTA antigen repair solution (pH 8.0) (Servicebio, catalog number: G1206)
PBS buffer (Servicebio, catalog number: G0002)
Tissue autofluorescence quenching agent (Servicebio, catalog number: G1221)
Bovine serum albumin (BSA) (Servicebio, catalog number: GC305010)
DAPI staining reagent (Servicebio, catalog number: G1012)
Antifade mounting medium (Servicebio, catalog number: G1401)
MPO primary antibody (Servicebio, catalog number: GB15224)
DSPP primary antibody (Servicebio, catalog number: sc-73632)
Cy3-labeled goat anti-mouse IgG (Servicebio, catalog number: GB21301)
Goldner staining solution suit (Servicebio, catalog number: G1064)
Glacial acetic acid (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 10000218)
Hydrochloric acid (Sinopharm Group Chemical Reagent Co., Ltd., catalog number: 10011018)
Solutions
1.2% tribromoethanol (see Recipes)
0.9% NaCl (see Recipes)
1 mg/mL LPS-Pg (see Recipes)
95% ethyl alcohol (see Recipes)
90% ethyl alcohol (see Recipes)
85% ethyl alcohol (see Recipes)
1% hydrochloric acid alcohol solution (see Recipes)
0.2% glacial acetic acid (see Recipes)
Recipes
1.2% tribromoethanol
Reagent Final concentration Amount
Tribromoethanol (absolute) 1.2% 0.12 g
H2O n/a 10 mL
Total n/a 10 mL
Weigh 0.12 g of tribromoethanol using an electronic scale with 0.001 g accuracy and dissolve it in 10 mL of distilled water; then, mix thoroughly with a vortex mixer.
0.9% NaCl
Reagent Final concentration Amount
NaCl (absolute) 0.9% 0.9 g
H2O n/a 100 mL
Total n/a 100 mL
Weigh 0.9 g of NaCl using an electronic scale with 0.1 g accuracy and dissolve it completely in 100 mL of distilled water.
1 mg/mL LPS-Pg
Reagent Final concentration Amount
LPS-Pg (absolute) 1 mg/mL 1 mg
Endotoxin-free water n/a 1 mL
Total n/a 1 mL
Aspirate 1 mL of endotoxin-free water with a 1 mL injection syringe and inject it into a glass medicine bottle containing 1 mg of LPS-Pg powder; mix these components to obtain a 1 mg/mL LPS-Pg solution.
Aliquot and label the LPS-Pg solution in sterile 0.1 mL tubes and freeze them at -20 °C until use.
Note: LPS-Pg solution at a concentration of 100 mg/mL is stable for >6 months if stored at -20 °C.
95% ethyl alcohol
Reagent Final concentration Amount
Absolute ethanol 95% 95 mL
Distilled water n/a 5 mL
Total n/a 100 mL
Using a 100 mL graduated cylinder, measure 95 mL of absolute ethanol and transfer it to a 100 mL beaker. Using a 5 mL graduated cylinder, measure 5 mL of distilled water and add it to the beaker containing ethanol. Add a magnetic stir bar to the beaker, place the beaker on a magnetic stirrer, and stir for 5 min.
90% ethyl alcohol
Reagent Final concentration Amount
Absolute ethanol 90% 90 mL
Distilled water n/a 10 mL
Total n/a 100 mL
Using a 100 mL graduated cylinder, measure 90 mL of absolute ethanol and transfer it to a 100 mL beaker. Using a 10 mL graduated cylinder, measure 10 mL of distilled water and add it to the beaker containing ethanol. Add a magnetic stir bar to the beaker, place the beaker on a magnetic stirrer, and stir for 5 min.
85% ethyl alcohol
Reagent Final concentration Amount
Absolute ethanol 85% 85 mL
Distilled water n/a 15 mL
Total n/a 100 mL
Using a 100 mL graduated cylinder, measure 85 mL of absolute ethanol and transfer it to a 100 mL beaker. Using a 25 mL graduated cylinder, measure 15 mL of distilled water and add it to the beaker containing ethanol. Add a magnetic stir bar to the beaker, place the beaker on a magnetic stirrer, and stir for 5 min.
1% hydrochloric acid alcohol solution
Reagent Final concentration Amount
Hydrochloric acid 1% 1,000 μL
Absolute alcohol n/a 99 mL
Total n/a 100 mL
Using a 100 mL graduated cylinder, measure 99 mL of absolute ethanol and transfer it to a 100 mL beaker. Using a 1,000 μL pipette, measure 1,000 μL of hydrochloric acid and add it to the beaker containing ethanol. Add a magnetic stir bar to the beaker, place the beaker on a magnetic stirrer, and stir for 30 s.
0.2% glacial acetic acid
Reagent Final concentration Amount
Glacial acetic acid 0.2% 200 μL
Distilled water n/a 99.8 mL
Total n/a 100 mL
Using a 200 μL pipette, measure 200 μL of glacial acetic acid and add it to a 100 mL volumetric flask. Add distilled water to the flask while stirring with a glass rod until the solution reaches a volume of 100 mL.
Laboratory supplies
Diamond bur (MANI, catalog number: TC-SS21F)
Dental file (Velbon, catalog number: K23120)
Operating scissors (Velbon, catalog number: J21030)
Ophthalmic forceps, straight (Velbon, catalog number: JD1050)
Ophthalmic forceps, curved (Velbon, catalog number: JD1060)
Dressing forceps (Velbon, catalog number: J42035)
Icebox (Biosharp, catalog number: BC032)
Icebox, silicone base (Biosharp, catalog number: BC034)
1 mL injection syringe (needle dimensions: 0.45 × 15 mm)
Pipette (10 μL, 200 μL, 1,000 μL) (Servicebio, catalog numbers: SPIP-10, SPIP-200, SPIP-1000)
Bagged tips (10 μL, 200 μL, 1,000 μL) (Servicebio, catalog numbers: P-10, P-200, P-1000)
0.1 mL centrifuge tubes (Thermo Fisher Scientific, catalog number: 4358297)
0.1 mL centrifuge tube rack (Servicebio, catalog number: WGH002)
Nitrile gloves (Servicebio, catalog number: GN1801M)
Stainless-steel lunch box (China Industry Union, catalog number: WGH0001)
Oral surgery kit (HENAN SHENG YUBEI EISAI Co., Ltd., catalog number: 20202171244)
Medical adhesive tape
Medical cotton balls: large and small
Permanent ink markers
Square medical sharps container
Medical waste garbage bags
Beaker
Doctor's scrubs
Embedding frame (Servicebio, catalog number: EF-1)
Ophthalmic scissors (Velbon, catalog number: JC2303)
Immunohistochemical pen (Servicebio, catalog number: G6100)
Graduated cylinder (5 mL, 10 mL, 25 mL, 100 mL)
Magnetic stirrer (Servicebio, catalog number: MS-150)
Magnetic stir bar (Servicebio, catalog number: WGA0023)
Volumetric flask (100 mL)
Equipment
Electronic scale with 0.1 g accuracy (LICHEN, catalog number: YP10001B)
Electronic scale with 0.001 g accuracy (Sartorius, catalog number: BCA224I-1OCN)
-20 °C freezer (MeiLing, catalog number: DW-YL450)
Autoclave (China Industry Union)
Head-mounted dental loupe (3.5×, black, 5 W headlight, China Industry Union)
Vortex mixer (Servicebio, catalog number: SMV-3500)
Portable dental treatment machine (Greeloy, catalog number: GU-P206S)
Timer
Dehydrator (DIAPATH, model: Donatello)
Embedding machine (Wuhan Junjie Electronics Co., Ltd., catalog number: JB-P5)
Freezing platform (Wuhan Junjie Electronics Co., Ltd., catalog number: JB-L5)
Constant temperature shaker (TIANJIN LEIBO TERRY EQUIPMENT Co., Ltd., catalog number: ZHPW-250)
Microtome (Shanghai Leica Instrument Co., Ltd., catalog number: RM2016)
Tissue spreader (Zhejiang Kehua Instrument Co., Ltd., catalog number: KD-P)
Oven (Tianjin Laibo Rui Instrument Equipment Co., Ltd., catalog number: GFL-230)
Adhesive slides (Servicebio, catalog number: G6012)
Cover glass (Citotest Labware Manufacturing Co., Ltd., catalog number: 10212432C)
Upright optical microscope (Nikon, model: NIKON ECLIPSE E100)
Imaging system (Nikon, model: NIKON DS-U3)
Decolorization shaker (Servicebio, catalog number: DS-2S100)
Microwave oven (Galanz, catalog number: P70D20TL-P4)
Scanner (3DHISTECH, model: Pannoramic MIDI)
Software and datasets
CaseViewer (Version: 2.4; Copyright © 2001-2020 3DHISTECH Ltd.; Build: 2.4.0.119028)
ImageJ 1.54f (Wayne Rasband and contributors; National Institutes of Health, USA)
Procedure
Preparation
Prepare Recipes 1–8 following the instructions provided in Recipes section.
Autoclaving of the modeling instruments: Within 1–5 days before modeling, place the modeling instruments in a stainless-steel lunch box and sterilize them using an autoclave.
Modeling surgery
Anesthesia
Weigh the C57BL/6 mouse using a beaker and an electronic scale with 0.1 g accuracy. Use 1.2% tribromoethanol (see Recipes) for anesthesia (intraperitoneal injection), with a dosage of 0.5 mL per 20 g body weight (Figure 1A, B).
Fixing
Pinch the mouse’s toe to check its reaction. If the mouse does not respond when its toe is pinched, the mouse has reached a deep level of anesthesia. Then, secure the mouse to the surgical pad with medical adhesive tape; this will ensure that the mouse cannot interrupt the surgery (Figure 1C).
Opening the pulp chamber
Open the mouse’s oral cavity with curved ophthalmic forceps. Use dressing forceps to expand the oral cavity and protect the tongue, thus preventing injury by the diamond bur; concurrently, completely expose the upper portion of the mouth.
Use the diamond bur to access the maxillary first molar in the mouse, then expand the pulp exposure with the dental file; the final diameter of exposure should be approximately 1 mm. From the exposure, white and slightly red dental pulp tissue should be visible (Figure 1D).
Rinsing
Aspirate 0.1 mL of 0.9% NaCl solution (see Recipes) with a 1 mL injection syringe. Place the tip of the needle over the medullary foramen and quickly empty the syringe to flush out any dentin debris that may have entered the medullary foramen.
LPS induction
Extract 10 μL of 1 mg/mL LPS-Pg (see Recipes) into a 0.1 mL centrifuge tube. Use straight ophthalmic forceps to completely saturate a small cotton ball in the LPS-Pg solution. Keep the saturated small cotton ball on the exposed pulp for 5 min.
Pulp capping and tooth filling
Remove the cotton ball and restore the tooth with Esthet-X flow (Figure 1E).
After surgery, loosen the mouse’s restraints; return the mouse to the cage after it has awakened.
Real surgical images can be found in Figure S1.
Tissue decalcification and paraffin embedding (Figure 2C)
Sample collection
After day 1 and 10 of model establishment, sacrifice the mouse by cervical dislocation; carefully harvest maxillary tissue with straight ophthalmic forceps tweezers and ophthalmic scissors. Place the maxillary tissues into a 15 mL centrifuge tube with 10% paraformaldehyde fixative (neutral) and incubate at room temperature for 24 h.
Decalcification
Place printed labels and tissues in sequential order in an embedding frame. Then, place them in a basin, fill it with fresh decalcification solution, seal the container, and store all components in a constant temperature shaker at 25 °C with a speed of 110 RPM. This process results in decalcification. The decalcification solution should be replaced every 2–3 days. The degree of decalcification should be observed by pricking the tissue with a needle every 2 days. If the needle can be moved in the tissue, cut a plane with a Lycra blade along the edge of the maxillary tissue to remove excess tissue and ensure that the embedding surface is flat, which can accelerate the softening speed.
Figure 2. Operation process of tissue decalcification and paraffin embedding, HE staining, immunofluorescent staining, and immunofluorescent staining
Dehydration and wax infiltration
Remove the softened tissue from the decalcification solution, rinse with tap water for 5 h, place it into a dehydrating basket, and dehydrate using an alcohol gradient in the dehydrator. Prepare reagents for dehydration and wax infiltration: alcohol (75%, 85%, 90%, 95%), anhydrous ethanol (divide the liquid into three separate bottles labeled I, II, and III), benzyl alcohol, xylene (divide the liquid into two separate bottles labeled I and II), and melted paraffin (divide the liquid into three separate bottles labeled I, II, and III). The dehydration and wax infiltration steps are as follows: 75% alcohol for 2 h, 85% alcohol for 2 h, 90% alcohol for 1.5 h, 95% alcohol for 2 h, anhydrous ethanol I for 2 h, anhydrous ethanol II for 2 h, benzyl alcohol for 40 min, xylene I for 40 min, xylene II for 40 min, melted paraffin I for 0.5 h at 65 °C, melted paraffin II for 1 h at 65 °C, and melted paraffin III for 2 h 45 min at 65 °C.
Embedding
Transfer the wax-infiltrated tissue to the embedding machine. First, place the melted wax into the embedding frame. Then, place the tissue into the embedding frame according to the requirements of the embedding surface, and attach appropriate labels before the wax solidifies. Cool the wax on a -20 °C freezing platform. After solidification, remove the wax block from the embedding frame and trim it.
Sectioning
Use the trimmed wax block to prepare 4 μm thick paraffin sections. Float sections in warm water at 40 °C to flatten the tissue. Collect the sections and bake them in an oven at 60 °C. After the water has evaporated and the wax has melted, store the sections at room temperature.
HE staining (Figure 2D)
Dewaxing and rehydration
Prepare reagents: environmentally friendly dewaxing and clearing solution (divide the liquid into two separate bottles labeled I and II), anhydrous ethanol (divide the liquid into two separate bottles labeled I and II), and 75% ethyl alcohol. Immerse paraffin sections in the following sequence: environmentally friendly dewaxing and clearing solution I for 20 min, environmentally friendly dewaxing and clearing solution II for 20 min, anhydrous ethanol I for 5 min, anhydrous ethanol II for 5 min, and 75% ethyl alcohol for 5 min. Subsequently, rinse with tap water.
Hematoxylin staining
Stain the sections with hematoxylin solution for 3–5 min and rinse with tap water. Next, immerse the sections in hematoxylin differentiation solution and rinse with tap water. Finally, immerse the sections in hematoxylin bluing solution and rinse with tap water.
Eosin staining
Immerse sections in the following sequence of solutions: 85% ethanol for 5 min, 95% ethanol for 5 min, and eosin dye for 5 min.
Dehydration and sealing
Prepare reagents for dehydration and wax infiltration: absolute ethanol (divide the liquid into three separate bottles labeled I, II, and III) and xylene (divide the liquid into two separate bottles labeled I and II). Immerse sections in the following sequence of solutions: absolute ethanol I for 5 min, absolute ethanol II for 5 min, absolute ethanol III for 5 min, xylene for 5 min, and xylene for 5 min. Then, seal the sections with neutral gum.
Immunofluorescence staining (Figure 2E)
Dewaxing and rehydration
Prepare reagents: environmentally friendly dewaxing and clearing solution (divide the liquid into three separate bottles labeled I, II, and III) and anhydrous ethanol (divide the liquid into three separate bottles labeled I, II, and III). Sequentially immerse the sections in the following sequence: environmentally friendly dewaxing and clearing solution I for 10 min, environmentally friendly dewaxing and clearing solution II for 10 min, environmentally friendly dewaxing and clearing solution III for 10 min, absolute ethanol I for 5 min, absolute ethanol II for 5 min, and absolute ethanol III for 5 min. Subsequently, rinse with tap water.
Antigen retrieval
Place the sections in a water bath for 30 min. After retrieval is complete, allow sections to cool naturally. Wash the sections three times in PBS (pH 7.4) on a decolorization shaker for 5 min per wash.
Serum blocking
Dry the sections and use a hydrophobic pen to draw a circle around the tissue. Apply 3% BSA for 30 min to block nonspecific binding.
Primary antibody incubation
Prepare primary antibody: dilute the primary antibodies (DSPP 1:100; MPO 1:500) in PBS buffer. Add the diluted primary antibody to the sections, ensuring that they are lying flat in a humidified chamber. Incubate overnight at 4 °C.
Secondary antibody incubation
Wash the sections three times in PBS (pH 7.4) on a decolorization shaker for 5 min per wash. Add the secondary antibody (Cy3-labeled goat anti-mouse IgG) and incubate for 50 min at room temperature in the dark.
Stain cell nuclei with DAPI
Wash the sections three times in PBS (pH 7.4) on a decolorization shaker for 5 min per wash. Add the DAPI staining reagent and incubate for 10 min at room temperature in the dark.
Quench tissue autofluorescence and sealing
Wash the sections three times in PBS (pH 7.4) on a decolorization shaker for 5 min per wash. Add tissue autofluorescence quenching agent B and incubate for 10 min at room temperature in the dark. Use antifade mounting medium to seal the sections.
Goldner trichrome staining (Figure 2F)
Dewaxing and rehydration
Prepare reagents: Xylene (divide the liquid into two separate bottles labeled I and II), anhydrous ethanol (divide the liquid into two separate bottles labeled I and II), and 75% ethyl alcohol. Sequentially immerse the paraffin sections in the following sequence: xylene I for 20 min, xylene II for 20 min, anhydrous ethanol I for 5 min, anhydrous ethanol II for 5 min, and 75% ethyl alcohol for 5 min. Subsequently, rinse with tap water.
Nuclear staining
Mix Goldner staining solution A and Goldner staining solution B in equal proportions. Place sections in the mixed solution and stain for 20 min. Wash with tap water, then differentiate with 1% hydrochloric acid alcohol solution for 2 s. Rinse with tap water, then wash with distilled water.
Staining with Goldner staining solution C
Immerse sections in the first jar of Goldner staining solution C for 10 min, then quickly rinse with 0.2% glacial acetic acid (3 s per rinse).
Staining with Goldner staining solution D
Immerse sections in Goldner staining solution D for 3 min. Monitor the staining under a microscope; ensure that Goldner staining solution C fades from the collagen areas.
Staining with Goldner staining solution C
Immerse sections in the second jar of Goldner staining solution C for 5 min, then quickly rinse with 0.2% glacial acetic acid (3 s per rinse).
Staining with Goldner staining solution E
Immerse sections in Goldner staining solution E for 5 min. Then, differentiate the stain using three jars of 0.2% glacial acetic acid (2 s per jar). Dehydrate the sections using three jars of anhydrous ethanol (2 s, 3 s, and 5 s, respectively).
Clearing and sealing
Place sections in the third jar of anhydrous ethanol for 5 min. Clear the sections in xylene for 5 min. Finally, seal with neutral gum.
Perform microscopy examination, image acquisition, and analysis (Figure 1F)
HE staining: Nuclei will be stained blue, whereas cytoplasm will be stained red.
Immunofluorescence staining: DAPI (excitation wavelength 330–380 nm, emission wavelength 420 nm) will stain nuclei blue, whereas Cy3 (excitation wavelength 510–560 nm, mission wavelength 590 nm) will stain the target protein red.
Goldner trichrome staining: mineralized bone will be stained green, whereas non-mineralized tissue will be stained orange red.
Data analysis
HE staining results
Following above procedures (A–D, G), we constructed a mouse model of LPS-induced pulpitis (LPS surgery mice). To verify that pulpitis was induced by LPS, we established a solvent control group in which 1 mg/mL LPS was replaced with an equal volume of endotoxin-free water, and the same procedure was followed (solvent surgery mice).
We analyzed the HE staining results of LPS surgery mice and solvent surgery mouse models of pulpitis at 1 day (1 d), 10 d, and 30 d after surgery using Case Viewer and ImageJ. In the early stage of pulpitis, the primary manifestation is exudation. At 1 d post-surgery in LPS surgery mice (Figure 3A-A2), we observed a large number of necrotic tissues and extensive inflammatory exudate, including neutrophils, around the coronal cavity preparation (black arrow). Additionally, we noted many more neutrophils and a small amount of chronic inflammatory cell infiltrates at the junction between inflamed and normal tissue (black ellipse). These findings indicate that our model can be used to explore the early stages of pulpitis. After the early stage, pulpitis gradually transitions from exudation to hyperplasia, denoting the middle and later stages. In the middle and later stages, the primary manifestation is hyperplasia; this mainly includes fibroblast proliferation, blood vessel dilation, and the proliferation of chronic inflammatory cells. At 10 d post-surgery in LPS surgery mice (Figure 3B-B2), we observed vacuolar degeneration in some fibroblasts (blue arrow) and proliferation in others at the distal end of inflamed tissue (blue ellipse). Additionally, the pulp blood vessels were dilated and congested (black triangle). These observations suggest that our model can also be used to explore the middle and later stages of pulpitis. In the end stage, the final outcome typically comprises necrosis, although recovery is possible. At 30 d post-surgery in LPS surgery mice (Figure 3C-C2), we observed inflammation invading the coronal pulp cavity and most of the root pulp. The coronal pulp cavity and upper half of the root pulp were necrotic. Furthermore, necrotic tissue and inflammatory exudate, including neutrophils, were present around the coronal cavity preparation and right root canal (black arrow). These findings indicate that our model can be used to explore the end stage of pulpitis. Analysis with ImageJ revealed that the proportion of inflamed areas gradually increased with prolonged inflammation (Figure 3G). These results revealed obvious symptoms of pulpitis, confirming the successful establishment of our model.
The solvent control group did not exhibit a favorable inflammatory phenotype (Figure 3D–F). Comparisons of A with D, B with E, and C with F in Figure 3 indicate that LPS is necessary for the successful induction of pulpitis.
To investigate whether 1 mg/mL LPS is the optimal concentration for pulpitis induction, we established high-concentration (10 mg/mL) and low-concentration (0.1 mg/mL) groups, then followed the same procedure (Figure S2). The experimental results showed that pulpitis induced by 0.1 mg/mL LPS was too mild and obvious pulp repair could be seen, which could not reflect the development process of pulpitis. However, the inflammation induced by 10 mg/mL LPS was too heavy, and tissue disintegration occurred on the first day; pulpitis progressed too fast, it was not easy to observe the complete inflammatory process, the consumption of LPS reagent was large, and the cost was higher. In summary, the pulpitis model constructed with 1 mg/mL LPS exhibited the most complete pulpitis progression and was the most suitable concentration for constructing this model.
Figure 3. HE staining results for lipopolysaccharide (LPS) surgery mouse and solvent surgery mouse (1, 10, and 30 days post-surgery). A. 1 d post-surgery in LPS surgery mouse: Inflammation has not invaded the root pulp. A1: A large number of necrotic tissues and extensive inflammatory exudate, including neutrophils, are present around the coronal cavity preparation (black arrow). Many more neutrophils and a small amount of chronic inflammatory cell infiltrates are evident at the junction between inflamed and normal tissue (black ellipse). The pulp blood vessels are dilated and congested (black triangle). A2: The root pulp appears normal. B. 10 d post-surgery in LPS surgery mouse: Inflammation has invaded the coronal pulp cavity and upper portion of root pulp. B1: A large number of necrotic tissues and extensive inflammatory exudate, including neutrophils, are present around the coronal cavity preparation (black arrow). Many disintegrated neutrophils and substantial chronic inflammatory cell infiltrates are evident at the junction between inflamed and normal tissue (black ellipse). B2: Fibroblasts exhibit vacuolar degeneration (blue arrow) and proliferate at the distal end of inflamed tissue (blue ellipse). The pulp blood vessels are dilated and congested (black triangle). Necrotic tissue is visible in the upper portion of root pulp, whereas the lower portion of root pulp appears normal. C. 30 d post-surgery in LPS surgery mouse: Inflammation has invaded the coronal pulp cavity and most of the root pulp. The coronal pulp cavity and upper half of the root pulp are necrotic. C1: Necrotic tissue and inflammatory exudate, including neutrophils, are present around the coronal cavity preparation and right root canal (black arrow). C2: Many disintegrated neutrophils and substantial chronic inflammatory cell infiltrates are evident at the junction between inflamed and normal tissue (black ellipse). Fibroblasts are proliferating beneath the necrotic tissue (blue ellipse). The pulp blood vessels are dilated and congested (black triangle). D. 1 d post-surgery in solvent surgery mouse: Inflammation has not invaded the root pulp. D1: A small amount of inflammatory exudate is evident around the coronal cavity preparation (black arrow); intratissue hemorrhage is present, and pulp blood vessels under the inflammatory exudate are congested (black triangle). Some neutrophils and chronic inflammatory cell infiltrates can be observed (black ellipse). D2: The root pulp appears normal. E. 10 d post-surgery in solvent surgery mouse: Inflammation has invaded a small portion of the root pulp. E1: A large amount of inflammatory exudate, including neutrophils, is visible around the coronal cavity preparation (black arrow). Fibroblasts are secreting collagen fiber bundles to contain the inflammation (broken line). The pulp blood vessels are congested (black triangle). E2: Pre-mineralized tissue is present beneath the inflammatory exudate (broken line). The root pulp appears normal. F. 30 d post-surgery in solvent surgery mouse: Inflammation has invaded the root pulp. F1: A small amount of inflammatory exudate including neutrophils and chronic inflammatory cells are observed around coronal cavity preparation (black arrow). Fibrous matrix proliferates and fills part of the pulp cavity (broken line). The pulp blood vessels are congested (black triangle). F2: Some neutrophils and chronic inflammatory cell infiltrates are present in the root pulp (black ellipse). G. Analysis of the proportion of inflamed area at 1 d, 10 d, and 30 d post-surgery in LPS surgery mice (47% ± 10%, 74% ± 8%, and 89% ± 1%, respectively). The proportion of inflamed areas at 10 d post-surgery in LPS surgery mice significantly increased compared with 1 d post-surgery in LPS surgery mice (P < 0.01). The proportion of inflamed areas at 30 d post-surgery in LPS surgery mice significantly increased compared with 10 d post-surgery in LPS surgery mice (P < 0.05).
Immunofluorescence staining results
With the LPS-induced pulpitis model sections obtained in procedure C, we performed subsequent immunofluorescence staining (procedure E) with MPO as the staining indicator. Myeloperoxidase (MPO), also known as peroxidase, is derived from neutrophils, monocytes, and macrophages. In the process of inflammation, white blood cells release a large amount of MPO, so the detection of MPO expression can reflect the severity of inflammation. In Figure 4A, DAPI channel nuclei are shown in blue and MPO is shown in red. It could be observed that with the extension of LPS modeling time, the proportion of MPO positive areas increased significantly (P < 0.001, Figure 4B), indicating that the inflammation was gradually aggravated, suggesting that the modeling was successful.
Goldner trichrome staining results
With sections of LPS-induced pulpitis model obtained in procedure C, we performed subsequent Goldner trichrome staining (procedure F). Goldner trichrome staining can distinguish between mineralized bone (green) and non-mineralized tissue (orange red). In Figure 5, it can be seen that with the extension of modeling time, more and more mineralized tissues in the dental pulp of mice tended to form mature mineralized tissues, which successfully demonstrated the progress of pulpitis and proved the success of modeling.
Figure 4. Immunofluorescence staining results for no-surgery mouse and lipopolysaccharide (LPS) surgery mouse (1, 10, and 30 days post-surgery). A. Immunofluorescence staining results; red represents MPO and blue represents DAPI. B. Analyzing the proportion of positive areas in no-surgery, 1 d, 10 d, and 30 d post-surgery mice. MPO, myeloperoxidase.
Figure 5. Goldner trichrome staining results for no-surgery mouse and lipopolysaccharide (LPS) surgery mouse (1, 10, and 30 days post-surgery). No-surgery mouse shows no green in pulp cavity. 1 d post-surgery mouse shows punctate light green. 10 d post-surgery mouse shows thin cord-like and meshy light green in the coronal pulp. 30 d post-surgery mouse shows dense green in the coronal pulp diffusely and cord-like light green in distal pulp.
Validation of protocol
All four mice had induced pulpitis by LPS (Figure 3, 4, and 5), which sufficiently validated this protocol.
General notes and troubleshooting
Because the mouse’s oral cavity is small and it is difficult to obtain an appropriate mouth-opening device, we used dressing forceps to open the mouths of mice in this study. However, to prevent mandibular dislocation and adverse effects (e.g., incomplete mouth closure) related to the extended use of dressing forceps for mouth opening, straight ophthalmic forceps can be used for mouth opening after pulp exposure; this approach reduces damage to the maxillary and mandibular joints.
During the feeding period after modeling, the Esthet-X flow used to close the opening hole can fall off due to nibbling feed or bedding, which will lead to modeling failure. In order to avoid this situation, we use shavings cushion material and soft pasty feed after modeling. This operation can effectively avoid the shedding of Esthet-X flow.
Acknowledgments
This work was supported by the China National Natural Science Foundation (82071074); 2023-PUMCH-E-010.
Competing interests
The authors declare no competing interests.
Ethical considerations
All animal experiments were approved by the ethics committee of Capital Medical University (AEEI-2024-071).
References
Huang, H., Okamoto, M., Watanabe, M., Matsumoto, S., Moriyama, K., Komichi, S., Ali, M., Matayoshi, S., Nomura, R., Nakano, K., et al. (2023). Development of Rat Caries-Induced Pulpitis Model for Vital Pulp Therapy. J Dent Res. 102(5): 574–582.
Aubeux, D., Renard, E., Pérez, F., Tessier, S., Geoffroy, V. and Gaudin, A. (2021). Review of Animal Models to Study Pulp Inflammation. Front Dent Med. 2: e673552.
Krivanek, J., Soldatov, R. A., Kastriti, M. E., Chontorotzea, T., Herdina, A. N., Petersen, J., Szarowska, B., Landova, M., Matejova, V. K., Holla, L. I., et al. (2020). Dental cell type atlas reveals stem and differentiated cell types in mouse and human teeth. Nat Commun. 11(1): 4816.
Shi, X., Li, Z., He, Y., Jiang, Q. and Yang, X. (2017). Effect of different dental burs for experimental induction of pulpitis in mice. Arch Oral Biol. 83: 252–257.
He, Y., Gan, Y., Lu, J., Feng, Q., Wang, H., Guan, H. and Jiang, Q. (2017). Pulpal Tissue Inflammatory Reactions after Experimental Pulpal Exposure in Mice. J Endod. 43(1): 90–95.
Erdogan, O., Xia, J., Chiu, I. M. and Gibbs, J. L. (2023). Dynamics of Innate Immune Response in Bacteria-Induced Mouse Model of Pulpitis. J Endod. 49(11): 1529–1536.
Chung, M. K., Lee, J., Duraes, G. and Ro, J. (2011). Lipopolysaccharide-induced Pulpitis Up-regulates TRPV1 in Trigeminal Ganglia. J Dent Res. 90(9): 1103–1107.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Images of the surgical procedure
Figure S2. HE staining results for 0.1 mg/mL, 1 mg/mL, 10 mg/mL LPS surgery mouse (1 d and 10 d post-surgery) and no surgery mouse
Article Information
Publication history
Received: Apr 1, 2024
Accepted: Oct 5, 2024
Available online: Oct 23, 2024
Published: Jan 20, 2025
Copyright
© 2025 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Biological Sciences > Biological techniques
Medicine > Inflammation
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5,129 | https://bio-protocol.org/en/bpdetail?id=5129&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Culture and Characterization of Differentiated Airway Organoids from Fetal Mouse Lung Proximal Progenitors
ZZ Zhonghui Zhang *
CT Chengxu Tao *
QL Qiuling Li
(*contributed equally to this work)
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5129 Views: 247
Reviewed by: Valeria Fernandez ValloneSamantha Haller Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Aug 2023
Abstract
Developing a physiologically relevant in vitro model of the respiratory epithelium is critical for understanding lung development and respiratory diseases. Here, we describe a detailed protocol in which the fetal mouse proximal epithelial progenitors were differentiated into 3D airway organoids, which contain terminal-differentiated ciliated cells and basal stem cells. These differentiated airway organoids could constitute an excellent experimental model to elucidate the molecular mechanisms of airway development and epithelial cell fate determination and offer an important tool for establishing pulmonary dysplasia disease in vitro.
Key features
• Efficient isolation of proximal epithelial progenitors from mouse embryos.
• Differentiation of pulmonary airway organoids differentiated from tracheal progenitors, which recapitulates the process of airway cell differentiation.
• Airway organoids can be used to explore the molecular mechanisms of lung development and respiratory diseases.
Keywords: Airway organoids Fetal Proximal progenitors Lung development
Graphical overview
Overview of the culture of airway organoids from isolated E13.5 mouse proximal epithelial cells
Background
The pseudostratified airway epithelium is the first mechanical barrier of the respiratory system and maintains tissue homeostasis after injury. The airway epithelium consists of secretory club cells, terminally differentiated ciliated cells, basal stem cells, and a small population of goblet cells, neuroendocrine cells, and ionocytes [1]. All types of proximal airway epithelial cells differentiate from Sox2+ progenitors during embryonic lung development [2]. Deficiencies in early embryonic development of the airway epithelium are frequently associated with neonatal mortality, as well as a variety of life-threatening lung diseases in adulthood, including bronchopulmonary dysplasia, chronic obstructive pulmonary disease, and lung cancer.
Recent advances in in vitro 3D culture techniques enable the generation of organoids derived from stem or progenitor cells, which functionally and morphologically mimic the airway epithelium at a near-physiological level [3,4]. This approach has allowed researchers to elucidate the molecular mechanisms of lung development and the consequences of genetic alterations associated with various respiratory diseases [5,6]. Additionally, several research groups have developed lung organoid disease models for preclinical drug screening [7–10].
We previously reported a tracheal progenitor isolation and culture system that can generate airway organoids containing differentiated epithelial cells [11]. Here, we provide a detailed protocol to generate 3D airway organoids with hollow structures derived from isolated proximal epithelial progenitors of E13.5 mouse tracheas, the same source for generating airway organoids as previously reported [11]. Compared with the trans-well air-liquid interface (ALI) culture, this protocol requires fewer progenitors and the resulting 3D organoids are more suitable for in vivo transplantation for the study of diseases [12,13]. This robust in vitro 3D culture model provides an opportunity to explore the molecular mechanisms of early lung development and relevant respiratory diseases in an experimentally tractable system.
Materials and reagents
Biological materials
10–12-week-old adult male mice and sexually attracted female mice; obtained from the Animal Experimental Center of Anhui Medical University
Reagents
PBS (Thermo Fisher, catalog number: 20012027), store at RT
Ethanol absolute (Hushi, catalog number: 10009218), store at RT
Sucrose (Sigma-Aldrich, catalog number: V900116), store at RT
Triton X-100 (Sigma-Aldrich, catalog number: T8787), store at RT
Normal goat serum (Gibco, catalog number: 16210-064), shelf-life is 3 months, store at -20 °C
NaN3 (Sigma-Aldrich, catalog number: S2002), dissolve 2 g in 1 mL of sterile Milli-Q-treated water, store at -20 °C.
Caution: NaN3 is hazardous. Avoid inhalation, ingestion, and skin contact.
Tween-20 (Sigma-Aldrich, catalog number: P1379), add 10 µL of Tween-20 in PBS to 10 mL, store at RT
Optimal cutting temperature compound (OCT) (Sakura, catalog number: 4853), store at RT
Chloral hydrate (MACKLIN, catalog number: 302-17-0), dissolve 10 g in 90 mL of sterile Milli-Q-treated water, store at 2–8 °C
Matrigel basement membrane matrix, growth factor reduced (GFR) (Corning, catalog number: 354230), store at -80 °C
Note: Thaw Matrigel on an icebox in a 4 °C refrigerator, aliquot in 200 μL vials upon arrival, and minimize freeze and thaw cycles; it will polymerize at temperatures above 10 °C.
Paraformaldehyde (PFA) (MACKLIN, catalog number: 30525-89-4), dissolve 20 g of PFA in 500 mL of sterile PBS to prepare 4% PFA. Shelf-life is 2 months; store at 2–8 °C
Caution: PFA is hazardous. Avoid inhalation, ingestion, and skin contact. Use protective equipment and work in a fume hood.
Type I rat tail collagen (Corning, catalog number: 354236), dissolve 4 mg of Type I rat tail collagen in 1 mL of 0.02 N acetic acid to prepare 4 mg/mL Type I rat tail collagen, then add 50 μL of 4 mg/mL Type I rat tail collagen in 1.95 mL of 0.02 N acetic acid to prepare 0.1 mg/mL Type I rat tail collagen. Store at -20 °C
Acetic acid (Sigma-Aldrich, catalog number: 695092), dissolve 1 mL in 87 mL of sterile Milli-Q-treated water. Use a syringe filter to filter the acetic acid solution; store at -20 °C
Penicillin/streptomycin (PS) (Life Technologies, catalog number: 15140-163), with 10,000 units penicillin and 10 mg streptomycin per milliliter in 0.9% NaCl; store at -20 °C
DMEM (high glucose, L-glutamine, sodium pyruvate, sodium bicarbonate, phenol red, w/o HEPES) (VivaCell, catalog number: 113-0500), store at 2–8 °C
DMSO (Solarbio, catalog number: D8370), store at RT
Advanced DMEM/F-12 (Gibco, catalog number: C11330500BT), store at 2–8 °C
GlutaMAX, 0.4 M (Gibco, catalog number: 35050061), store at 2–8 °C
HEPES, 1 M (Gibco, catalog number: 15630-080), store at 2–8 °C
PrimocinTM, 50 mg/mL (InvivoGen, catalog number: ant-pm), store at -20 °C aliquoted. Avoid multiple freeze-thaw cycles
NaHCO3 (Sigma-Aldrich, catalog number: S6014), dissolve 0.75 g in 10 mL of sterile Milli-Q-treated water. Use a syringe filter to filter the solution; store at -20 °C
FBS (Biowest, catalog number: S1580-500), store at -20 °C
HBSS (Gibco, catalog number: 14175), store at RT
BSA (Sigma-Aldrich, catalog number: V900933), dissolve 0.1 g in 10 mL of PBS to prepare 1% BSA, store at 2–8 °C
EGF (Corning, catalog number: 354001), dissolve 100 μg in 4 mL of HBSS + 0.1% BSA. Shelf-life is 3 months; store at -20 °C aliquoted
CHIR99021 (Tocris, catalog number: 4423), dissolve 1 mg in 716.3 μL of DMSO. Shelf-life is 3 months; store at -20 °C aliquoted
Y-27632 (Tocris, catalog number: 1254), dissolve 1 mg in 624.5 μL of PBS. Shelf-life is 1 year; store at -20 °C aliquoted
Retinoic acid (Sigma-Aldrich, catalog number: R2625), dissolve 1 mg in 13.31 mL of DMSO. Make fresh each time and store at -20 °C
Note: Vortex retinoic acid stock before making the working solution. This is light sensitive; store in the dark.
Transferrin (Sigma-Aldrich, catalog number: T1147-100 mg), dissolve 100 mg in 20 mL of HBSS + 0.1% BSA. Shelf-life is 2 months; store at -20 °C aliquoted
A-8301 (AbMole, catalog number: M5037), dissolve 5 mg in 23.72 mL of DMSO. Shelf-life is 2 months; store at -20 °C aliquoted
Insulin (Sigma-Aldrich, catalog number: I6634), dissolve 100 mg in 50 mL of 4 mM HCl. Shelf-life is 28 days; store at -20 °C aliquoted
Noggin (Sino Biological, catalog number: 10267), dissolve 1 mg in 50 μL of PBS. Shelf-life is 1 year; store at -20 °C aliquoted
Cholera toxin (Sigma-Aldrich, catalog number: 9012-63-9), dissolve 1 mg in 4 mL of HBSS + 0.1% BSA. Shelf-life is 1 year; store at -20 °C aliquoted
Pronase (Sigma-Aldrich, catalog number: 9036-06-0), dissolve 15 mg of Pronase in 10 mL of Ham’s F12
Note: Use a 0.22 µm sterile filter to filter the Pronase solution and make fresh each time.
Ham’s F12 (Corning, catalog number: 10-080-cv), store at 2–8 °C
Anti-fluorescence quenching agent (Beijing Biotopped Technology, catalog number: Top0702). Shelf-life is 1 year; store at 2–8 °C. This is light sensitive; store in the dark
Antibodies
Anti-mouse Tp63 (Santa Cruz Biotechnology, catalog number: sc-25268), we recommend dilution at 1:400 with dilution buffer and storing at -20 °C aliquoted. Fresh dilution is prepared before staining
Anti-mouse acetylated α Tubulin (Abcam, catalog number: ab-24610), we recommend dilution at 1:400 with dilution buffer and storing at -20 °C aliquoted. Fresh dilution is prepared before staining
Anti-rabbit Krt5 (Abcam, catalog number: ab-64081), we recommend dilution at 1:250 with dilution buffer and storing at -20 °C aliquoted. Fresh dilution is prepared before staining
Goat anti-rabbit 555 (Life Technologies, catalog number: A21430), we recommend dilution at 1:1,000 with dilution buffer and storing at -20 °C aliquoted. This is light sensitive; store in the dark. Fresh dilution is prepared before staining
Goat anti-mouse 488 (Life Technologies, catalog number: A11029), we recommend dilution at 1:1,000 with dilution buffer and storing at -20 °C aliquoted. This is light sensitive; store in the dark. Fresh dilution is prepared before staining
DAPI (Sigma-Aldrich, catalog number: D9542), we recommend dilution at 1:1,000 with dilution buffer and storing at -20 °C aliquoted. This is light sensitive; store in the dark. Fresh dilution is prepared before staining
Solutions
Buffer solution (see Recipes)
Suspension medium (see Recipes)
mTEC/basic medium (see Recipes)
mTEC/Plus1 medium (see Recipes)
mTEC/Plus2 medium (see Recipes)
Blocking buffer (see Recipes)
Dilution buffer (see Recipes)
Recipes
Buffer solution (50 mL)
Shelf-life is 1 month; store at 2–8 °C.
Reagent Final concentration Volume
PBS n/a 49.5 mL
Penicillin/streptomycin 500 U/mL 500 μL
Suspension medium (50 mL)
Make fresh each time and store at 2–8 °C.
Reagent Final concentration Volume
DMEM n/a 47 mL
FBS 5% 2.5 mL
Penicillin/streptomycin 500 U/mL 0.5 mL
mTEC/basic medium (50 mL)
Shelf-life is 1 month; store at 2–8 °C.
Reagent Stock concentration Final concentration Volume
Advanced DMEM/F-12 n/a n/a 48.05 mL
HEPES 1 M 15 mM 0.75 mL
Penicillin/streptomycin 500 U/mL 500 U/mL 0.5 mL
GlutaMAX 0.4 M 4 mM 0.5 mL
NaHCO3 0.9 M 3.6 mM 0.2 mL
mTEC/Plus1 medium (3 mL)
Shelf-life is 1 month; store at 2–8 °C.
Reagent Stock concentration Final concentration Volume
MTEC/basic medium n/a n/a 2.82 mL
FBS 100% 5% 150 μL
EGF 25 μg/mL 25 ng/mL 3 μL
Insulin 2 mg/mL 10 μg/mL 15 μL
Transferrin 5 mg/mL 5 μg/mL 3 μL
Cholera toxin 250 μg/mL 0.1 μg/mL 1.2 μL
PrimocinTM 50 mg/mL 100 μg/mL 6 μL
mTEC/Plus2 medium (1 mL)
Make fresh each time and store at 2–8 °C.
Reagent Stock concentration Final concentration Volume
MTEC/Plus1 medium n/a n/a 996.4 μL
Retinoic acid 250 μM 50 nM 0.2 μL
Y-27632 5 mM 5 μM 1 μL
CHIR99021 3 mM 3 μM 1 μL
A8301 500 μM 500 nM 1 μL
Noggin 20 mg/mL 20 μg/mL 1 μL
Blocking buffer (10 mL)
Make fresh each time and store at 2–8 °C.
Reagent Stock concentration Final concentration Volume
PBS n/a n/a 7.98 mL
Normal goat serum 100% 20% 2 mL
Triton X-100 100% 0.1% 10 μL
NaN3 2 g/mL 0.2 mg/mL 1 μL
Tween-20 100% 0.05% 5 μL
Dilution buffer (10 mL)
Make fresh each time and store at 2–8 °C.
Reagent Stock concentration Final concentration Volume
PBS n/a n/a 9.78 mL
Normal goat serum 100% 2% 0.2 mL
Triton X-100 100% 0.1% 10 μL
NaN3 2 g/mL 0.2 mg/mL 1 μL
Tween-20 100% 0.05% 5 μL
Laboratory supplies
1 mL syringe (Jiufekang, catalog number: 045X15RWLB), containing 4.5-gauge syringe needle
24-well plates (Corning, catalog number: 353047)
35 mm culture dish (Sparkjade, catalog number: GF0006)
60 mm culture dish (Sparkjade, catalog number: GF0007)
1.5 mL centrifuge tube (Sparkjade, catalog number: NZ-86009)
Falcon, 50 mL centrifuge tube (Wocas, catalog number: NZ-86011)
Falcon, 15 mL centrifuge tube (Wocas, catalog number: NZ-86010)
Pipette tip, 10, 200, 1,000 µL (Wocas, catalog numbers: NZ-86001, NZ-86002, NZ-86003)
Parafilm M wrapping film (Thermo Fisher, catalog number: 1337416)
300-mesh cell strainers (Sangon biotech, catalog number: F513453)
Adhesion microscope slides (Citotest, catalog number: 80312-3161)
Syringe filter (SHIGOUYI, catalog number: SFMCE013022NA)
Dyeing tank (Biosharp, catalog number: BS-WB-20B)
Equipment
Cell culture hood (Nuaire, model: NU-425-400S)
Cell culture incubator (Nuaire, model: NU-5510E)
Cell counter (Biosharp, model: BS-QT-1103)
Cryostat (Yidi, model: YD-1900), set from 0 to -40 °C
Surgical scissors (Merck, catalog numbers: S3146)
Surgical forceps (Biolab, catalog numbers: Z168777 and Z225614)
Vortex mixer (Kylin-bell, model: VORTEX-5)
Ice machine (PHCbi, model: SIM-F140AY65-PC)
Autoclave (TOMY, model: SX-500)
Brightfield microscope (Leica, model: DMI3000B)
Cell culture benchtop centrifuge (Merck, model: MGL-16MA)
Micropipettes (Dragon, catalog numbers: 7010101004, 7010101005, 7010101006, 7010101009, 7010101016)
Pure water technology (Zhiang, model: Best-R)
37 °C water bath (Changzhou Yuexin, model: HH-1)
Stereomicroscope (Olympus, model: SZX16)
Multiphoton laser scanning microscope (Leica, model: TCS SP8 DVE)
Procedure
Obtaining mouse at 13.5 days of pregnancy
Cage one 10–12-week-old adult male mouse and two sexually attracted female mice together at about 6 p.m.
Note: House mice in the animal room and control the temperature (21 ± 2 °C) and humidity (50% ± 10%) with a 12/12 h light/dark cycle.
Check female vaginal plugs the next morning; mark the morning of the day when you detect a vaginal plug as E0.5.
Weigh mice every day. Pregnant females generally show a detectable increase in body weight from E6.5 onward, with a 0.5–2 g increase at E13.5 based on the number of embryos in the womb.
Separation of the proximal airways from E13.5 embryos
Before dissecting E13.5 mice, disinfect the stereomicroscope and operating table with 75% ethanol and sterilize the forceps and scissors by autoclaving to minimize the possibility of contamination during the experiment. In addition, prepare an ice box, 35 and 60 mm Petri dishes, and 1.5 and 50 mL tubes containing buffer solution (see Recipes section) (Figure 1).
Figure 1. Example of equipment and materials and dissection tools. A. Stereomicroscope. B. Materials and dissection tools needed for proximal airway dissection: tubes (50 mL, 1.5 mL) with buffer solution, Petri dishes (35 mm, 60 mm) with buffer solution, surgical scissors, and straight thin tip forceps.
Prepare surgical tools: surgical scissors, surgical forceps, and surgical bed, and select an E13.5 pregnant mouse from the laboratory animal house.
Intraperitoneally inject 0.3 mL of chloral hydrate and wait for 5 min. Clamp the mouse's back foot with forceps; if the mouse does not struggle and twitch, assume that the mouse does not feel pain (Figure 2A).
Figure 2. Representative pictures of E13.5 lung and trachea dissection. Each panel illustrates steps B2–B14. Scale bars: 1 cm (A–I); 100 μm (J–L).
Fix the pregnant mouse to the surgical bed using 4.5-gauge syringe needles (Figure 2B).
Disinfect the abdomen area with 75% ethanol (Figure 2B).
Pinch and lift the skin with forceps (Figure 2C).
Dissect along the mid-abdominal line with surgical scissors, starting from the pubic area and continuing to the end of the abdominal cavity. Inject PBS into the heart, revealing the abdominal organs (Figure 2D).
Note: This procedure helps to avoid accidental injury to the embryo located below.
Place the uterus with embryos with forceps on the 60 mm dish filled with buffer solution maintained on ice (Figure 2E).
Note: Sacrifice the mouse by cervical dislocation after placing the uterus with embryos on the dish.
Separate each embryo from the uterus with the intact yolk sac using forceps (Figure 2F).
Remove each embryo from the uterus and keep them in the dish with buffer solution on ice using forceps.
Place one embryo in a 35 mm culture dish containing 1 mL of buffer solution using forceps (Figure 2G).
Discard the yolk sac, amniotic membrane, and umbilical cord under the stereomicroscope using forceps. Keep the rest on ice (Figure 2H).
Remove the head and abdomen of the embryo, cut the rest under the liver, pull the chest skin of the embryo open, and open the rib-enclosed part of the chest cavity (Figure 2I–J).
Find the red heart part of the embryo mouse and identify the trachea and lung parts around the heart; then, remove the esophagus from the lung and carefully remove the remaining excess tissues with surgical forceps (Figure 2K).
Note: The stripped airway should remain as intact as possible. Work fast to prevent progenitor cell damage.
Remove the proximal airway with forceps and place in the 35 mm culture dish containing buffer solution at 4 °C (Figure 2L).
Next, dissect the remaining embryos placed on ice.
Isolation of trachea epithelial progenitors
Under the microscope, using fine-tipped forceps, clean off the connective tissue around the trachea in a 35 mm culture dish.
Open the trachea longitudinally using scissors and further dissect the connected tissue using forceps. Wash it three times in a 1.5 mL centrifuge tube containing 200 μL of buffer solution to get off the connected tissue around the proximal airway.
Put all cleaned tracheae into a 1.5 mL centrifuge tube containing 1 mL of Pronase solution using forceps, and digest at 4 °C overnight (approximately 14 h).
Note: Prolonged digestion with Pronase might be harmful for progenitor cells.
Transfer the undigested trachea to a new 1.5 mL tube using forceps and use 200 μL of mTEC/basic medium (see Recipes) to pipette up and down.
Note: After the airway is digested overnight, there is still some tissue around the airway, such as cartilage, and some progenitor cells in the remaining tissue.
Mix the cleaned solution with the mixture of cellular digestive solution. Centrifuge at 800× g for 5 min at 37 °C.
Discard the supernatant and add 1 mL of suspension medium (see Recipes).
Note: When separating the proximal airway progenitor cells of the embryonic lung, pay attention to the placement of the centrifuge tube in the centrifuge to prevent cell loss.
Use a 300-mesh cell strainer (i.e., 300 holes per inch) to filter the cell suspension into a new 15 mL tube and centrifuge at 1,000× g for 3 min.
Discard the supernatant.
Resuspend the cells in 1 mL of prewarmed mTEC/basic medium and transfer into a 1.5 mL tube.
Note: Prewarm the required volume of mTEC/Basic medium in a water bath (37 °C) for approximately 15 min before use.
Prepare type I rat tail collagen coated 35 mm dish.
Note: First, add 1 mL of collagen coating solution in a 35 mm dish, then aspirate the collagen coating solution and allow the membrane to air-dry for at least 5 min. Rinse both apical and basal surfaces with 1 mL of sterile PBS three times to remove free collagen. Allow the 35 mm dish coated surface to dry for 5 min before use and make fresh each time.
Place the cell suspension prepared in step C9 on a 35 mm dish coated type I rat tail collagen. Incubate at 5% CO2 at 37 °C for 3 h.
Collect the nonadherent cells in a new 15 mL tube carefully.
Centrifuge at 1,000× g for 5 min.
Aspirate the supernatant.
Resuspend the cells in 1 mL of mTEC/basic medium.
3D in vitro culture of proximal epithelial progenitors
Thaw Matrigel on ice overnight at 4 °C before use.
Note: When handling Matrigel, always keep it on ice at all times and work quickly. Matrigel will tend to polymerize as it warms up.
Add 100 µL of Matrigel in the bottom of a 24-well plate and place it at 37 °C for 30 min.
Centrifuge the proximal airway progenitor cells at 1,000× g for 10 min at 37 °C and discard the supernatant.
Carefully aspirate the supernatant and resuspend the cells in mTEC/Plus2 medium (see Recipes).
Cell counting: We use a cell counter to calculate the number of cells, and we control 800 cells per 100 µL of cell suspension.
Note: Adjust the cells to 20–50 cells per large grid to minimize counting errors.
Mix Matrigel with the cell suspension on ice at a ratio of 1:1.
Note: Put the resuspended cell–Matrigel mixture on ice to avoid Matrigel solidification.
Inoculate a ratio of 800 cells per well on the center of the Matrigel-coated 24-well plate and ensure cell growth in the 3D droplet shape of Matrigel. Place at 37 °C for 30 min.
After Matrigel has polymerized into a jelly-like texture, add 500 µL of mTEC/Plus2 medium along the wall of the 24-well plate.
Incubate at 5% CO2 at 37 °C for 18 days.
Note: Prewarm the mTEC/Plus2 medium in a 37 °C water bath for 10–20 min before use.
Change the culture medium every two days during the organoid culture and replace half of the mTEC/Plus2 medium each time.
Take pictures on days 1, 2, 3, 6, 9, 12, 15, and 18.
Proximal epithelial progenitor cells start to form lumen-like structures on the second day of culture, which gradually grow larger during subsequent cultures, eventually forming hollow, spherical airway organoids (Figure 3).
Figure 3. Top view of 3D culture of airway organoids differentiated from mouse lung proximal progenitors. The pictures show a representative picture of organoids culture on days 1, 2, 3, 6, 9, 12, 15, and 18. Scale bars: 25 μm.
Identification of airway organoids by immunofluorescence staining
Organoid fixation
Prepare 1% BSA solution.
Carefully aspirate the mTEC/Plus2 medium.
Add 1 mL of pre-cooled PBS to each well.
Coat the 15 mL tube and pipette tip using 1% BSA solution to wash the tube wall.
Transfer organoids in the 24-well plates to the 15 mL centrifuge tube using the pipette tip that was washed with 1% BSA solution, and then pipette up and down the 24-well plate with 1 mL of 1% BSA solution. Transfer to the 15 mL centrifuge tube.
Add 10 mL of pre-cooled PBS to the 15 mL centrifuge tube, centrifuge at 70× g for 3 min at 4 °C, and carefully aspirate the supernatant. Repeat step E1e.
Add 1 mL of 4% (wt/vol) PFA to fix the organoids at 4 °C for 30 min. Pipette and mix them every 10 min, paying attention to controlling the pipetting force to prevent the organoid structure from being destroyed.
After fixation, add 10 mL of 0.1% Tween-20 and suspend organoids. Then, centrifuge at 70× g for 3 min at 4 °C and carefully aspirate the supernatant.
Add 2 mL of PBS.
Note: Organoids in PBS can be stored at 4 °C for up to a week.
Frozen sections
Centrifuge at 70× g for 3 min at 4 °C before soaking the fixed organoids overnight in PBS with 10% sucrose at 4 °C.
Use a cutting pipette tip to place the organoids on the sample table and add OCT; then, directly freeze in a pre-cooled -20 °C frozen cryostat.
Slice blocks of frozen organoids to a thickness of 8 μm under a cryomicrotome and attach the tissue to an adhesive microscope slide.
Store the frozen slices at -20 °C for immunofluorescence staining.
Immunofluorescence staining
Thaw frozen sections in a sealed environment to avoid damage by frozen water crystals.
Incubate with 0.5% Triton X-100 in PBS for 15 min.
Note: This step helps with permeabilization, especially for sections thicker than 8 μm.
Block with blocking buffer (see Recipes) for 30 min at RT.
Note: This step will help to reduce background staining; a longer incubation may be more effective.
Clean around the tissue after blocking with wiping paper.
Note: The wiping paper should be free of scraps.
Dilute the antibody with dilution buffer (see Recipes) in different proportions and mix well in 1.5 mL centrifuge tubes. Then, add to the tissue drop by drop and cover with parafilm M wrapping film to prevent evaporation. Keep it still during incubation at 4 °C overnight to avoid membrane movement.
Note: When performing immunofluorescence staining, add sterile Milli-Q treated water to the bottom of the dyeing tank to prevent evaporation from drying the tissue. Also, keep it bubble-free when placing the parafilm on the tissue.
On the next day, use the dyeing tank to soak the tissue with 0.5% Triton X-100 in PBS three times for 5 min each time.
Dilute the corresponding fluorescent secondary antibody and DAPI dye with dilution buffer as shown in the antibodies list, then add to the tissue drop by drop and incubate at RT for 2 h.
Note: Avoid exposure to light after the addition of secondary antibody and DAPI.
Soak in the dyeing tank with 0.5% Triton X-100 in PBS three times at RT for 5 min each time.
After adding 10 μL of anti-fluorescence quenching agent, add a coverslip, wrap the samples in tin foil, and scan tissues with a multi-photon laser scanning microscope.
Note: The samples wrapped in tin foil can be stored at 4 °C for up to one week.
As shown in Figure 4, immunofluorescence staining characterization of airway organoids shows that the organoids cultured for 18 days are multilayered cellular structures, which express the terminally differentiated ciliated cell-specific marker Ac-tubulin on the luminal side and the basal cell-specific markers (Tp63 and Krt5) on the peripheral side (Figure 4).
Figure 4. Representative sectional immunofluorescent images showing that organoids express basal cell markers (Krt5, red; Tp63, green) and differentiated ciliated cells (Ac-tub, green). DAPI stains the nuclei (blue). Scale bars: 25 μm.
Validation of protocol
The reproducibility of the protocol has been described within the corresponding article: Li et al. [11].
General notes and troubleshooting
Troubleshooting (Table 1)
Table 1. Troubleshooting suggestions
Problem Possible reason Possible solution
Few progenitor cells were isolated from a single airway. The airway is prone to rupturing during the stripping process. 1. Increase the amount of digested tissues.
2. Constantly shake the digestive fluid that contains the airways.
Clogging of mesh. After overnight digestion, undigested tissues, such as cartilage, remain. Refilter using a new 300-mesh cell strainer.
Organoids fail to form lumen-like structures.
1. The initial seeding density of tracheal progenitors is too high.
2. The various factors added to the medium are not preserved properly and lose potency.
1. Reduced density of tracheal progenitor cells.
2. If lumen-like structures fail to form, the experiment must be attempted again from the beginning.
Trachea epithelial progenitors and organoids appear to drop to the bottom of the Matrigel droplet.
1. The Matrigel has been repeatedly frozen and thawed.
2. Matrigel and cell mixture placed at 37 °C for too short.
1. Matrigel must be aliquoted, stored strictly in accordance with the manufacturer's instructions, and used within the expiration date.
2. Extend 20 min of incubation of Matrigel and cell mixture at 37 °C.
Acknowledgments
We thank the staff of the facility of the Institute of Health Sciences & Technology, Anhui University, for providing technical support. This work was supported by grants from the National Natural Sciences Foundation of China (32270880).
This protocol was derived from the previous publication Li et al. [11].
Competing interests
The authors declare that they have no conflicts of interest with the contents of this article.
Ethical considerations
This study was approved by the animal welfare committee of Anhui University (IACUC(AHU)-2022-012).
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Li, Q., Jiao, J., Heng, Y., Lu, Q., Zheng, Y., Li, H., Cai, J., Mei, M. and Bao, S. (2023). Prmt5 promotes ciliated cell specification of airway epithelial progenitors via transcriptional inhibition of Tp63. J Biol Chem. 299(8): 104964.
Bukowy-Bieryłło, Z. (2021). Long-term differentiating primary human airway epithelial cell cultures: how far are we? Cell Commun Signaling. 19(1): 63.
Barkauskas, C. E., Chung, M. I., Fioret, B., Gao, X., Katsura, H. and Hogan, B. L. M. (2017). Lung organoids: current uses and future promise. Development. 144(6): 986–997.
Article Information
Publication history
Received: Aug 7, 2024
Accepted: Oct 7, 2024
Available online: Oct 22, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Developmental Biology > Cell growth and fate > Differentiation
Stem Cell > Organoid culture
Cell Biology > Cell isolation and culture > 3D cell culture
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5,130 | https://bio-protocol.org/en/bpdetail?id=5130&type=0 | # Bio-Protocol Content
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Rapid Method for Estimating Polyhydroxybutyrate Accumulation in Bacteria Using Sodium Hypochlorite
Ingrid E. Redersdorff *
AR Ailen N. Rodríguez *
ME Mariana Escobar
CS Claudia A. Studdert
MS M. Karina Herrera Seitz
(*contributed equally to this work)
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5130 Views: 195
Reviewed by: Andrea GramaticaDaniel Segura Segura GonzalezAlejandro Avilés-Reyes
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Original Research Article:
The authors used this protocol in Journal of Applied Polymer Science Feb 2024
Abstract
This protocol outlines the use of the previously described sodium hypochlorite extraction method for estimating the accumulation of polyhydroxybutyrate (PHB) in bacteria. Sodium hypochlorite (NaClO) is widely used for PHB extraction as it oxidizes most components of the cells except PHB. We assessed the feasibility of using NaClO extraction for the estimation of PHB accumulation in bacterial cells (expressed as a percentage w/w). This allowed us to use a simple spectrophotometric measurement of the turbidity of the PHB extracted by NaClO as a semiquantitative estimation of PHB accumulation in the marine microorganisms Halomonas titanicae KHS3, Alteromonas sp., and Cobetia sp. However, this fast and easy protocol could be used for any bacterial species as long as some details are considered. This estimation exhibited a good correlation with the accumulation measured as dry cell weight or even with the accumulation measured by crotonic acid and HPLC quantifications. The key advantage of this protocol is how fast it allows an estimation of PHB accumulation in Halomonas, Alteromonas, and Cobetia cultures (results are available in 50 min), enabling the identification of the appropriate moment to harvest cells for further extraction, polymer characterization, and accurate quantification using more reliable and time-consuming methods. This protocol is very useful during bacterial cultivation for a quick evaluation of PHA accumulation without requiring (i) large volumes of cultures, (ii) a long time for analysis compared to dry cell weight, (iii) preparation of standard curves with sulfuric acid hydrolysis for crotonic acid quantification, or (iv) specific equipment and/or technical services for HPLC quantification.
Key features
• Fast and easy method for bacterial PHB content estimation in cultures of different marine microorganisms. It can be used in other PHB-accumulating bacteria.
• Useful to explore culture conditions to achieve maximal PHB accumulation.
• Useful to follow the kinetics of both PHB accumulation and mobilization throughout culture development.
• In cultures with high (50%–70% dry cell weight) or very low (<15%) PHB accumulation, differences are visible by the naked eye before spectrophotometric measurement.
Keywords: Polyhydroxybutyrate Sodium hypochlorite oxidation Halomonas sp. Alteromonas sp. Cobetia sp.
Graphical overview
Rapid estimation of polyhydroxybutyrate content in bacteria using modified sodium hypochlorite treatment
Background
Polyhydroxyalkanoates (PHAs) are bacterial-origin polymers that share several characteristics with hydrocarbon-derived plastics. Among PHAs, polyhydroxybutyrates (PHBs) are the most common type, known for their biocompatibility and biodegradability. PHAs were first described in 1927 by Lemoigne [1] in bacteria from the genus Bacillus, and their presence in the cell was associated with the observation of Sudan Black–stained lipid inclusions. As early as 1958, Williamson and Wilkinson [2] described a method still widely used for the extraction and purification of PHAs from cells—treatment with an alkaline solution of sodium hypochlorite (NaClO). When Bacillus cells were suspended in such a solution, they were almost completely dissolved, including spores if present. Williamson and Wilkinson determined the best conditions (pH, temperature, incubation time) for the hypochlorite treatment and observed that the turbidity of the cell suspension started to decrease from the beginning of the treatment and reached a constant value when the cell lysis was complete. Moreover, the residual turbidity was proportionally higher in cell suspensions containing larger amounts of lipid inclusions as judged by microscopic observation, and they were able to show that the residual turbidity was strongly correlated with the concentration of PHA (µg/mL) remaining after the treatment. Hypochlorite treatment of Halomonas titanicae KHS3 cells that accumulated PHAs, followed by two washes with distilled water and one wash with ethanol, rendered a material that, upon NMR analysis, was found to be pure polyhydroxybutyrate [3]. PHB content could then be quantified by measuring the dry weight of hypochlorite-resistant material relative to the total dry biomass or, more precisely, through complete acid hydrolysis of the PHB material for spectrophotometric determination. However, neither of those possibilities allowed a rapid and continuous evaluation of the onset of the accumulation stage or the moment at which the cells reach a certain level of PHB content. It was then considered that the old turbidimetric measurements could be used to standardize a rapid, simple, and inexpensive method for the estimation of PHB content in growing cultures of PHB-accumulating strains. In the study conducted by Williamson and Wilkinson in 1958 [2], it was found that high cell density suspensions resulted in excessive residual turbidity, possibly due to incomplete cell lysis. As part of our protocol, we set a maximum cell density of OD600nm = 2 for the assay. By using a simple spectrophotometric measurement of the remaining turbidity after NaClO hydrolysis, our fast protocol resulted in a semiquantitative estimation of the percentage of PHB accumulation expressed as weight/weight in the marine microorganisms Halomonas titanicae KHS3, Alteromonas sp., and Cobetia sp. This estimation exhibited a reliable estimation of the accumulation, providing values that correlated to PHB weight/dry cell weight or even with the accumulation measured by crotonic acid and HPLC quantifications. Moreover, when propionic acid was added to the culture media, Halomonas titanicae KHS3 accumulated the co-polymer poly-hydroxy-butyrate-valerate (PHBV), and it was also possible to estimate PHBV accumulation using the described protocol. Therefore, although not tested here, it is highly probable that this protocol will also work for other PHAs and bacterial species.
Materials and reagents
Biological materials
Environmental strains isolated from seawater and identified as Halomonas titanicae KHS3, Alteromonas sp., and Cobetia sp.
Reagents
Dipotassium phosphate (K2HPO4) (Merck, catalog number: 105104)
Monopotassium phosphate (KH2PO4) (Fluka, catalog number: 104873)
Ammonium sulfate [(NH4)2SO4] (Merck, catalog number: 101217)
Sodium chloride (NaCl) (J.T. Baker, catalog number: 3624-19)
Magnesium sulphate heptahydrate (MgSO4·7H2O) (Anedra, catalog number: 10034-99-8)
Ferric chloride hexahydrate (FeCl3·6H2O) (Fluka, catalog number: 44944)
Sodium hypochlorite (NaClO) [5.5% (w/v) commercially available household bleach] *
Yeast extract (YE) (Oxoid, catalog number: LP0021)
Glycerol (Gly) (BioPack, catalog number: 1620.08)
Distilled water
Bacteriological agar (Oxoid, catalog number: LP0011)
*Note: Store at room temperature. Undiluted household bleach has a shelf life of six months to one year; afterward, it degrades and loses oxidative activity.
Solutions
H1 minimal medium (see Recipes)
Yeast extract 10% (YE) (see Recipes)
Glycerol 75% (Gly) (see Recipes)
Saline solution (NaCl 2%) (see Recipes)
Recipes
Note: All solutions must be autoclaved before use.
H1 minimal medium (200 mL)
Reagent Final concentration Quantity or Volume
K2HPO4 11.2 g/L 2.24 g
KH2PO4 4.8 g/L 0.96 g
(NH4)2SO4 2 g/L 0.4 g
NaCl 20 g/L 4 g
MgSO4·7H2O (1 M) 1 mM 1 mL*
FeCl3·6H2O (0.5% (w/v)) 1.85 µM 0.1 mL*
H2O n/a 200 mL
Total n/a 200 mL
*Add after autoclaving.
Note: Different carbon sources are used depending on the desired culture condition: for PHB accumulation condition, 3% glycerol; for non-PHB accumulation condition, 0.25% yeast extract. To prepare plates, add 1.5% bacteriological agar to the medium. This protocol utilizes marine halotolerant bacteria, which are cultivated in H1 minimal medium. If non-halotolerant bacteria are used, appropriate media must be prepared.
Yeast extract (YE) 10% (w/v) (20 mL)
Reagent Final concentration Quantity or Volume
Yeast extract 100 g/L 2 g
H2O n/a 20 mL
Total n/a 20 mL
Glycerol (Gly) 75% (v/v) (100 mL)
Reagent Final concentration Quantity or Volume
Glycerol 945 g/L 75 mL
H2O n/a 25 mL
Total n/a 100 mL
Saline solution (NaCl 2%) (100 mL)
Reagent Final concentration Quantity or Volume
NaCl 20 g/L 2 g
H2O n/a 100 mL
Total n/a 100 mL
Note: For the cell suspension solution when using halophilic or halotolerant microorganisms. Otherwise, use a solution with the appropriate composition (saline/buffer solution similar to the growth medium).
Laboratory supplies
Microcentrifuge tube 2 mL (Henso Medical Co. Ltd., catalog number: HSN1411)
Micropipette tips 200 μL (Henso Medical Co. Ltd., catalog number: HSN1402)
Micropipette tips 1,000 μL (Henso Medical Co. Ltd., catalog number: HSN1402)
Flask 250 mL (Pyrex, catalog number: 4442-250)
Glass Petri dishes (Pyrex, catalog number: 3160-60)
Glass reagent bottle 500 mL (Duran, catalog number: 218014459)
Equipment
Micropipette P1000 (100–1,000 μL) (Gilson, model: P1000)
Micropipette P200 (20–200 μL) (Gilson, model: P200)
Glass alcohol burner (Everglass, catalog number: EVG1381)
Mini centrifuge (Eppendorf, model: MiniSpin plus)
Cell density meter (Biochrom, model: UltrospecTM 10 Classic)
Cuvettes quartz glass (Hellma, model: 6040-UV-10-531)
Orbital incubator shaker (Amerex Instruments, model: Gyromax 727)
Autoclave 13.5 L (Ficoinox, model: SL9000)
Procedure
Bacterial growth under accumulation and no-accumulation (control) conditions
Spread a fresh H1 minimal medium plus 0.25% yeast extract plate of H. titanicae or the desired microorganism and incubate at the appropriate temperature and time. Since this protocol could be used for any bacterial strain, the selected culture media will depend on the bacterial species.
Start a fresh 3 mL overnight culture inoculated from the fresh plate from step A1.
Inoculate a 50 mL culture in a 250 mL flask with an initial OD600nm of approximately 0.05, using either accumulation medium (H1 medium with 3% glycerol) or non-accumulation medium (H1 medium with 0.25% yeast extract).
Monitor cell growth by measuring optical density (OD600nm) using 1 mL of sample for spectrophotometric determination. Collect samples at desired intervals to evaluate PHB accumulation.
Evaluation of PHB accumulation
Determine OD600nm of culture to be evaluated.
Harvest an appropriate volume of culture. The volume of samples needed for PHB evaluation at each growth point will depend on the OD600nm values of the culture. At each growth stage, collect enough culture to obtain 2 mL of cell suspension with OD600nm between 1.7 and 2.
Example: For a culture with an OD600nm of 0.45, it will be necessary to harvest between 7.55 and 8.9 mL of culture to obtain 2 mL of cell suspension with a final OD600nm between 1.7 and 2.0.
(final volume desired of cell suspension) × (final desired OD600nm) = (culture volume needed) × (OD600nm of culture)
2 mL × 2 = X mL × 0.45
Once the required volume of culture is determined, centrifuge at 9,700× g for 5 min to pellet the cells. Remove the supernatant by pouring off the remaining culture medium and gently withdrawing the residue with a P200 pipette.
Resuspend the cell pellet in 2 mL of 2% NaCl solution. Alternatively, cells can be resuspended in 2 mL of fresh medium without a carbon source. This step should be adapted to each bacterial strain requirement. For example, non-halophilic microorganisms’ cells could be resuspended in a low ionic-strength buffer solution.
Divide the cell suspension into two fractions, each of 700 µL, in microcentrifuge tubes. Label the tubes as dH2O and NaClO.
Add 150 µL of distilled water to the tube labeled dH2O and 150 µL of sodium hypochlorite [from a 5.5% (w/v) stock solution, commercially available household bleach] to the tube labeled NaClO. The final concentration of NaClO will be 1%. Gently mix to homogenize.
Incubate at 37 °C for 30 min.
After incubation, gently homogenize and measure the OD600nm for each tube within 15 min after completion of incubation.
The percentage of accumulated PHA is estimated using the formula described below in the Data analysis section (see examples in Figure 1).
Figure 1. Estimation of polyhydroxybutyrate (PHB) content by Na-hypochlorite and dry weight. Values obtained with the described NaClO method are compared with the ones obtained after treating the cells with Na-hypochlorite for 30 min at 37 °C, followed by two washes with water and one wash with ethanol. % PHB (dry weight) content was then calculated as: [(mg/mL dry PHB)/(mg/mL dry cells)] × 100
Data analysis
The following formula is used to estimate the percentage of accumulated PHB after Na-hypochlorite incubation:
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600
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(
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×
100
Validation of protocol
This protocol has been used and validated in the following research article:
Rodríguez et al. [3]. Characterization of polyhydroxybutyrate production from Halomonas titanicae KHS3 and manufacturing of electrosprayed nanoparticles. Journal of Applied Polymer Science. 141(6): e54928. (Supplementary information S1)
The percentage of PHB estimated with hypochlorite was graphically correlated with the percentage using other methods (Figure 2). The values used for this analysis (Table 1) were obtained in multiple experiments in different conditions.
Figure 2. Graphical correlation between the percentage of PHB estimated with hypochlorite and other methods. The black circles indicate the correlation of PHB estimation vs. the percentage of PHB by dry weight; the gray circles indicate a correlation with the percentage of PHB obtained from quantification on HPLC. The R2 was 0.9733. Obtained from Rodríguez et al. [3].
Table 1. Correlation between the content of PHB estimated with hypochlorite and dry weight. The values were obtained in multiple experiments in different conditions. *Obtained by HPLC quantification.
% PHA (hypochlorite) % PHA (dry weight)
11.39 9.17
49 48
5.99 4.07
4.37 5.50
15 11.11
28.20 31.35
7 2
83.50 87.50
12 15.80
77.60 72.20
13 6
58.77 48
75.41 70
63.15 66
72.20 72.38
6.30 6.04
46.80 49.70
44 37.50
37 30*
42.60 35*
30.33 31.35*
38.60 34.80*
Dry cell weight was determined by the gravimetric method using 20 mL of culture and harvesting cells by centrifugation at 7,600× g for 10 min at 4 °C. Supernatants were discarded, and cell pellets were washed twice with distilled water. Pellets were dried at 105 °C up to constant weight.
Polymer extraction was performed as described previously by Williamson and Wilkinson [2] with slight modifications. This method is based on the resistance of PHA to sodium hypochlorite treatment. Alkaline sodium hypochlorite solution was added to the culture to a 1% w/v final concentration and incubated at 37 °C for 1 h. Then, the mixture was centrifuged at 9,700× g for 10 min at 4 °C, and the polymer pellet was collected and washed twice with distilled water and finally washed with ethanol 96%. The pellet was dried at 65 °C up to constant weight.
General notes and troubleshooting
General notes
When using this protocol with new microorganisms, verify that the extracted compound is a PHB by nuclear magnetic resonance (NMR) or Fourier-transform infrared spectroscopy (FTIR). The correlation between this PHB estimation method and other PHB quantification methods should be verified for each different microorganism to be tested.
This protocol was evaluated on extreme halophilic haloarchaea, but no reliable results were obtained.
Troubleshooting
Problem 1: False positive.
Possible cause: Undiluted household bleach has a shelf life of six months to one year; beyond this period, it degrades and loses its oxidative activity.
Solution: Use fresh hypochlorite solution. Check that non-accumulating cultures are clarified after NaClO treatment.
Problem 2: Values of accumulation are much higher than expected.
Possible cause: If the optical density of the initial suspension is too high, it is possible that the concentration of NaClO or the incubation time were not enough for the complete dissolution of cellular content.
Solution: Be certain that the optical density of the initial cell suspension is between 1.7 and 2.
Problem 3: Too low OD600nm values in the control sample (“dH2O”).
Possible cause: Sometimes, bacterial cells do not sediment properly after centrifugation, making it difficult to obtain an appropriate measurement of OD600nm values for the control sample (“dH2O”). This could lead to OD600nm values lower than 0.5 for this control sample, and this sample is excluded from the analysis.
Solution: Higher centrifugation forces could improve bacterial cell harvesting.
Acknowledgments
This protocol was possible thanks to the early observations of Williams and Wilkinson [2] about the resistance of the PHB inclusions to treatment with alkaline hypochlorite and the measurement of turbidity as a means to assess the accumulation.
This study was funded by the National Scientific and Technical Research Council (CONICET). We also extend our gratitude to the Institute of Biological Research, National University of Mar del Plata (IIB, UNMDP), and the Institute of Agrobiotechnology of the Litoral (IAL, Santa Fe) for their valuable support.
Competing interests
There are no conflicts of interest or competing interests.
References
Lemoigne, M. (1927). Études sur l’autolyse microbienne. Origine de l’acide ß-oxybutyrique formé par autolyse. Ann Inst Pasteur. 41: 148.
Williamson, D. H. and Wilkinson, J. F. (1958). The Isolation and Estimation of the Poly-β-hydroxy-butyrate Inclusions of Bacillus Species. J Gen Microbiol. 19(1): 198–209.
Rodríguez, A. N., Escobar, M., Redersdorff, I. E., Studdert, C. A., Abraham, G. A., Cortez Tornello, P. R. and Herrera Seitz, M. K. (2023). Characterization of polyhydroxybutyrate production from Halomonas titanicaeKHS3 and manufacturing of electrosprayed nanoparticles. J Appl Polym Sci. 141(6): e54928.
Article Information
Publication history
Received: Aug 13, 2024
Accepted: Oct 10, 2024
Available online: Oct 22, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY license (https://creativecommons.org/licenses/by/4.0/).
How to cite
Category
Microbiology > Microbial biochemistry > Other compound
Biochemistry > Other compound
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5,131 | https://bio-protocol.org/en/bpdetail?id=5131&type=0 | # Bio-Protocol Content
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Application of a Dual Optogenetic Silencing-Activation Protocol to Map Motor Neurons Driving Rolling Escape Behavior in Drosophila Larvae
AS Ankura Sitaula
YH Yuhan Huang
AZ Aref Zarin
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5131 Views: 346
Reviewed by: Ivan Sanchez DiazRupkatha Banerjee Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Proceedings of the National Academy of Sciences of the United States of America Dec 2023
Abstract
Drosophila larvae exhibit rolling motor behavior as an escape response to avoid predators and painful stimuli. We introduce an accessible method for applying optogenetics to study the motor circuits driving rolling behavior. For this, we simultaneously implement the Gal4-UAS and LexA-Aop binary systems to express two distinct optogenetic channels, GtACR and Chrimson, in motor neuron (MN) subsets and rolling command neurons (Goro), respectively. Upon exposure to white LED light, Chrimson permits the influx of positive ions into Goro neurons, leading to depolarization, whereas GtACR mediates chloride influx into MNs, resulting in hyperpolarization. This method allows researchers to selectively activate certain neurons while simultaneously inhibiting others within a circuit of interest, offering a unique advantage over current optogenetic approaches, which often utilize a single type of optogenetic actuator. Here, we provide a detailed protocol for the dual silencing-activation approach using GtACR and Chrimson optogenetic channels and present a robust methodological framework for investigating the neuromuscular basis of rolling in larvae. Our cost-effective and scalable approach utilizes readily accessible equipment and can be applied to study other locomotor behaviors in Drosophila larvae, thereby enhancing our understanding of the neural circuit mechanisms underlying sensorimotor transformation.
Key features
• Enables real-time manipulation of neural activity, providing insights into the immediate effects of neuronal activation and silencing on larval behavior.
• The protocol is adaptable to different experimental setups, allowing researchers to extend its application to other sensory modalities or behavioral assays.
• Offers a standardized approach to studying nociceptive behaviors.
Keywords: Drosophila Rolling Larvae Motor neurons Optogenetics Cost-effective
Graphical overview
Experimental setup and procedure for the rolling assay. A) 69E06-LexA; Aop-Chrimson::tdTomato, UAS- GtACR1-eGFP female virgins are crossed with MN-Gal4 male flies, resulting in larval offspring carrying all four components essential for concurrent silencing and activation of neurons of interest. In these larvae, GtACR1-eGFP is selectively expressed in the motor neurons (MNs) of interest, whereas Aop-Chrimson::tdTomato is only expressed in Goro command neurons. B) Upon exposure to white LED light, Chrimson causes an influx of cations (Ca2+ and Na+) into Goro neurons, resulting in depolarization (activation) of these command neurons. In contrast, when exposed to white LED light, GtACR1 triggers an influx of chloride anions (Cl-), thereby hyperpolarizing (silencing) the MNs of interest. C) Both GtACR1 and Chrimson are sensitive to ambient light; therefore, the larvae should be raised in the dark until they are ready for experimentation.
Background
Noxious stimuli instinctively trigger escape behaviors in animals, prompting rapid motor responses to avoid harm. Understanding the neural mechanisms governing escape behaviors is essential for elucidating how animals respond to environmental threats. When subjected to harmful mechanical stimuli or heat, Drosophila larvae execute nocifensive escape behavior characterized by bending into a C-shape, followed by rolling and rapid forward crawling [1]. This behavior is triggered by the activation of class IV (cIV) dendritic arborization sensory neurons, which are polymodal nociceptor sensory neurons distributed throughout the body wall [2,3]. Rolling can also be induced optogenetically by stimulating nociceptive sensory neurons or interneurons within the central nervous system, including the Goro command neuron [4–6].
The body muscles of Drosophila larvae are co-innervated by type Is (small boutons, phasic firing) and a single type Ib motor neuron (MN) (big boutons, tonic firing). Type Ib MNs typically innervate one muscle, whereas Type Is MNs innervate multiple target muscles [7,8]. Here, we present a detailed protocol describing how we used a dual-optogenetic method to identify muscles and MN types essential for larval rolling escape behavior [9]. Current approaches to studying neural circuits often involve either optogenetic activation or inhibition of neural targets, limiting our ability to take full advantage of light-activated ion channels to explore circuits of interest. This protocol leverages the LexA-Aop system to express Chrimson, a cationic channelrhodopsin, in Goro neurons (69E06-LexA) to induce the larval rolling escape behavior. Concurrently, we utilize MN-Gal4 driven GtACR1, a light-sensitive chloride channel activated by the same white light as Chrimson, to acutely inhibit specific subsets of MN subsets. This dual optogenetic strategy enables the targeted and acute silencing of different MNs using different MN-Gal4 lines during the same temporal window of Goro activation. This setup, including a system to rapidly control light exposure, ensures consistent behavioral responses, thus enhancing the reproducibility and precision of behavioral studies in larvae. By implementing MN silencing in conjunction with rolling assays, we recently elucidated the neuromuscular basis of larval escape behavior in Drosophila [9].
This approach can be extended to investigate multifactorial motor neuron diseases, such as amyotrophic lateral sclerosis (ALS), which results from a combination of genetic and environmental factors that remain incompletely understood. The neural activation approach (69E06-LexA; LexA-Aop-Chrimson) can be integrated with reverse genetic screens to identify genes critical for the maintenance of motor circuit health and function, thereby elucidating novel genetic defects underlying motor neuron diseases, such as ALS. Optogenetic activation of the GORO neuron induces continuous rolling behavior in Drosophila larvae, which potentially leads to more rapid larval exhaustion than default forward crawling. The exhausted larvae may provide a more sensitive background to identify genes essential for neural function and maintenance compared to non-threatened larvae that perform forward crawling and have the opportunity to take brief pauses between crawling bouts to recover from subtle defects in motor circuit malfunction. To conduct these reverse genetic experiments, researchers can substitute UAS-GtACR1 with the UAS-RNAi line to knock down the candidate gene in the neurons of interest. Alternatively, 69E06-LexA; LexA-Aop-Chrimson activation could be employed in larvae carrying null mutant alleles for any candidate gene.
Materials and reagents
Biological materials (Drosophila stocks)
Commercially available Drosophila stocks
Type Is MN driver: w;;27E09-Gal4 (Bloomington Drosophila Stock Center, catalog number: 49227)
Type Ib MNs of ventral longitudinal (VL) and dorsal oblique (DO) muscles driver: w; exex-Gal4 (HB9-Gal4) (Bloomington Drosophila Stock Center, catalog number: 83004)
Lab-generated Drosophila stocks
Goro neuron driver: w; 69E06-LexA; Aop-Chrimson::tdTomato, UAS- GtACR1-eGFP [9]
Type Is and Ib MN1 driver: w; 27E09-Gal4; RRa-Gal4 [9]
Type Ib MNs of lateral transverse (LT) muscles driver: w; BH1-Gal4 [9]
Five type Ib MNs of dorsal longitudinal (DL) muscles driver: w; CQ-Gal4 [9]
Five type Ib MNs of DL muscles driver:: Unc4AD;vGlut DBD split Gal4 line [9]
Five Type Ib and Ib MN1 driver: w; CQ-Gal4; RRa-Gal4 [9]
Type Ib MNs of VL and DO muscles driver: w; vGlutAD;NKx6 DBD split-Gal4 [9]
Reagents
All trans-retinal (ATR) (Sigma-Aldrich, catalog number: R2500)
Agarose (Genesee Scientific, catalog number: 20-102GP)
Yellow cornmeal (Flystuff, catalog number: B078SZR5MS)
Drosophila Agar (Apex, catalog number: B078T16PNG)
Active dry yeast (SAF Instant, catalog number: B0049WLQ30)
Molasses (Oasis Supply, catalog number: B00M1ZYPXA)
Propionic acid (Genesee Scientific, catalog number: 20-271)
Tegosept (Genesee Scientific, catalog number: 20-259)
Ethanol (Sigma-Aldrich, catalog number: 64-17-5)
Solutions
Drosophila media (see Recipes)
10 mM ATR stock solution (see Recipes)
Recipes
Drosophila media
Reagent Quantity or Volume
Cornmeal 67 g
Drosophila agar 16 g
Active dry yeast 27 g
Molasses 67 mL
Propionic acid 5 mL
Tegosept 17 mL
Distilled water 1,100 mL
Begin by mixing cornmeal, agar, active dry yeast, molasses, and water in a large cooking vessel.
Place the vessel in a medium-to-high-heat setting and bring the fly food mixture to a boil.
Stir the mixture every 5–10 min to ensure even mixing.
Once boiling, reduce the heat slightly and let the mixture simmer gently for 30 min.
After 30 min of simmering, turn off the heat and allow the cooking vessel to cool until it is warm to the touch.
Carefully add the measured quantity of propionic acid and Tegosept to the warm mixture.
Transfer the well-mixed food into the Droso-Filler and pour the fly food into polypropylene vials.
Note: Any standard fly food will be suitable for this experiment; it does not need to be restricted to our recipe.
10 mM ATR stock solution
Reagent Quantity or Volume
ATR 1 mg
Ethanol 0.35 mL
Dissolve 1 mg of ATR in 0.35 mL of ethanol to create a 10 mM stock solution.
Store the stock solution in the dark at -20 °C.
Laboratory supplies
Petri dish 10 cm Ø (e.g., Sigma-Aldrich, catalog number: CLS430167)
Spatula (Genesee Scientific, catalog number: 93-131)
Wash bottle containing distilled water (e.g., Depepe, catalog number: B07DB1HCKP)
Paintbrush (Soucolor, catalog number: B07YDDF26Y)
Wide tip forceps (DR instruments, catalog number: B008RBLO8Q)
Cellulose acetate fly plugs (VWR, catalog number: 89168-886)
Polypropylene vials (VWR, catalog number: 75813-144)
Droso-Filler (Genesee Scientific, catalog number: 59-168)
Equipment
External light source 18 W LED (remote control included) (Oeegoo, catalog number: B086JP5Y91)
Extension cord (Husky, catalog number: HD#342-576)
Microscope lens smartphone adapter (Celticbird, catalog number: B0BXWPL7H4)
Stereomicroscope (ZEISS, model: Stemi-305)
Stereomicroscope (ZEISS, model: Stemi-508)
Smartphone camera operating at 30 frames per second (fps) or more.
Note: Any smartphone brand (iPhone, Samsung, or Google Pixel) offering a high frame rate and excellent optical quality can be used. The smartphone camera lens should be aligned with the eyepiece of the microscope. A resolution of at least 12 megapixels is recommended, along with a wide-aperture lens (f/1.5 to f/1.8) to ensure optimal light capture during imaging. The default camera application on a smartphone may be used for video recording. Alternatively, camera applications such as ProCamera (iOS), Open Camera (Android), or Camera FV-5 (Android) can also be used to capture high-quality images.
Software and datasets
ImageJ (https://imagej.net/)
FFmpeg (https://ffmpeg.org/) (version 2022-08-25-git-9bf9d42d01)
Procedure
Larval preparation
Prepare the larvae by raising parent flies on Drosophila media (see Recipes) in the dark at 25 °C, ensuring optimal growth conditions.
Note: Drosophila media is compatible with ATR administration.
Prepare food supplemented with ATR for the optogenetic experiments.
Final concentration of ATR needs to be 1 mM (see Recipes).
Note: In our experiments, 100 µL of 1 mM ATR was administered to the food vial containing larvae using a pipette.
Transfer the parents to a fresh vial and thoroughly distribute the appropriate volume of ATR within the food provided to the larvae in the original vial in a dimly lit room. For the no-ATR control condition, omit the addition of ATR to the larvae's food.
Allow the larvae to incubate for 24 h in the dark after administering ATR, as it acts as the chromophore essential for light-sensitive proteins like channelrhodopsins.
Note: The ATR can be added as early as after the eggs are laid to ensure the larvae are exposed to ATR for at least 24 h. For L3 larvae, we recommend adding ATR at the early L2 stage. For L1–L2 stages, we recommend adding ATR during late embryonic stages and assay the late L1 to early L2 stages.
Imaging setup
Replace the original 10× eyepiece of the Stemi 305 scope with a microscope phone adapter featuring a 16× built-in eyepiece to attach a phone for image capture (Figure 1). For imaging of L1 stage larvae, utilize a Stemi 508 dissection scope.
Install the smartphone to the phone adapter.
Secure the smartphone vertically to the microscope using a phone adapter that firmly attaches to the microscope to ensure stability and prevent movement during imaging.
Adjust the position of the smartphone so that its camera aligns precisely with the optical output of the microscope.
Utilize the smartphone’s camera settings to fine-tune the zoom and focus, ensuring the image is sharp and the area of interest is centered in the view.
Note: The images used in our experiments were captured using the default camera application of iPhone 14 with a 12 MP camera operating at 30 fps.
Set up the external light for the experiment.
Plug in the Oeegoo 18W square LED ceiling light before the experiment and set it up to ensure that it is a white light at maximum brightness.
Turn off the light using the remote; it will retain the settings when turned on again.
Position the light beneath the imaging microscope, ensuring that it illuminates the agarose pad placed atop it within the Petri dish (Figures 1 and 2).
Note: This assay should be conducted in a room with minimal light exposure to ensure accurate results and to reduce potential interference.
Figure 1. Step-by-step demonstration of the rolling assay setup. The procedure begins with the removal of the microscope lens (2), followed by the installation of an adapter (3) to secure the smartphone. Once the smartphone is placed in position (4) for imaging, an LED light source is installed (5) to provide illumination. Finally, the setup is completed with the placement of an agarose plate (6), ready for the rolling assay.
Figure 2. Experimental setup for the rolling assay. A phone camera is attached to the stereomicroscope with an agarose-filled Petri dish placed on top of the external light source (A) off and (B) on.
Optogenetic induction
Using a spatula, gently collect the food or media containing the larvae (Figure 3).
Figure 3. Step-by-step demonstration of larval preparation for optogenetic induction. Larval preparation begins by scraping the food from the vial (1) to retrieve the larvae. Next, the media is diluted in distilled water (2) to locate individual larvae. Following this, a single larva is picked and rinsed in distilled water to remove any residual food (3), placed on an agarose pad, and gently dried with a brush (4).
Slightly dilute the collected media in distilled water to facilitate the identification and location of individual larvae.
Isolate each larva individually using fine-tipped forceps and transfer it into a separate Petri dish containing distilled water.
Rinse each larva in the distilled water for 1–2 s to remove any residual food or debris.
Carefully transfer it onto a 1.5% agarose pad positioned within a Petri dish to provide a stable and uniform surface for imaging purposes.
Pat dry the larva using a paintbrush and provide a 30-s period of undisturbed rest in darkness to facilitate acclimation and minimize potential influences of light exposure on their behavior.
Note: Ensure the larva is dorsal side up at the beginning of the experiment.
Begin the video recording process and ensure continuous filming for the duration of the experiment.
Induce optogenetic activation of the larva's rolling behavior by turning on Oeegoo 18W square LED ceiling light and exposing them to 30 s of intense white light.
Note: Avoid subjecting each larva to more than three repeated optogenetic inductions to avoid stress and potential fatigue, with at least 2–3 min of rest between inductions.
Transfer the recorded videos to your computer. The rolling response should be analyzed manually by researchers blinded to the experimental conditions.
Data analysis
Transfer the video from your phone to your computer using a USB cable or cloud service. The video file size can range from 30 to 60 MB depending on the resolution and length of the video.
Open the video using FFmpeg software to convert it to a .avi file.
Open the converted video with ImageJ software.
Note: To analyze videos in ImageJ, the computer should have at least 8 GB of RAM and 500 MB of free storage and be running Windows 10, macOS 10.13+, or Linux.
Observe larvae's movements and spot both dorsal tracheae (Figure 4 and Video 1). A full 360° roll is defined as the larva starting from the dorsal side up, rolling completely, and returning to the dorsal side up position. Identify a full roll by observing the disappearance and reappearance of both tracheae (Video 1 and Video 2).
Figure 4. Sill images of L3 larvae showing 0°, 90°, 180°, 270°, and 360° of an outward roll in wild-type (GORO>Chrimson) rolling and defective rolling in HB9>UAS-GtACR1-eGFP, and Unc4AD-vGlutDBD>UAS-GtACR1-eGFP animals. Silencing Ib motor neurons (MNs) of ventral longitudinal (VL) and dorsal oblique (DO) muscles (HB9>UAS-GtACR1-eGFP) results in a significant inability to roll, approaching complete dysfunction. Silencing of dorsal longitudinal (DL) muscles (Unc4AD-vGlutDBD>UAS-GtACR1-eGFP) leads to animals being unable to complete a full 360° roll continuously. Black lines indicate dorsal tracheae. The left and right tracheae are both visible when the dorsal side is facing upward and invisible when the ventral side is facing upward.
Note: The larvae in the figure appear to be at different larval stages because of varying camera zoom levels during the experiment.
Video 1. Video of a wild-type third-instar larvae showing the rolling response with a 360° rolling. Wild-type rolling is initiated by bending to one lateral side while the dorsal plane is facing upward (identified by both tracheae being visible, labeled with black lines), followed by continuous body rotation.
Video 2. Examples of defective rolling. The video depicts a defective rolling in HB9>UAS-GtACR1 third-instar larva struggling to initiate rolling and Unc4AD-vGlutDBD>UAS-GtACR1-eGFP larva failing to complete a full 360° roll continuously.
Consider a roll successful if the larva completes at least one 360° rotation, demonstrating the ability to rotate in a single direction from the dorsal side up back to the dorsal side up.
Criteria for failed rolling: First, confirm that the larva can crawl in an upright position (dorsal up) in the dark or with minimum light without triggering the Chrimson response. Count the rolling as a failure if, upon light stimulation, the larva responds to light but cannot bend into a C-shape, bends but cannot initiate lateral rolling, or cannot complete at least one 360° roll. A successful 360° roll contains the C-bending at a dorsal-up position (both tracheae completely visible), continuous rolling (inward or outward the curve) in a single direction without changing the direction of the curve, and the tracheae become invisible as the larva rolls to ventral up and visible again as the larva returns to dorsal up (Figure 4, Video 1, and Video 2).
Calculate the percentage of larvae that were able to perform rolling to determine the success rate for each genotype.
Calculate rolling frequency by counting the number of full 360° rolls the larva executes in 30 s (use the frame count shown by ImageJ to measure the time precisely).
Calculate the rolling duration by counting the number of frames required to complete rolling within the 30s-time frame in ImageJ. The frame count can be found in the top-left corner of the ImageJ window. Divide the number of frames by 30 to get the rolling duration in seconds as the videos are recorded at 30 fps. For instance, if a larva requires 120 frames to complete four rolls within the 30s-time frame, divide 120 by 30 to yield a rolling duration of 4 s. It is important to acknowledge that certain larvae may experience interruptions in their rolling motion, resulting in brief pauses between successive rolls. In such cases, it is recommended to exclude these interrupted frames from the calculation of rolling duration. However, these interruptions can still provide valuable insights into larvae's behavior and may be noted separately as an additional parameter for analysis.
Note: Rolling duration can be calculated using Excel by dividing the total number of frames by 30.
Import all values into a programming environment of your choice and plot bar graphs showing the percentage of animals of different genotypes that are able to complete at least one complete roll to visualize the rolling success. Also, use the scatterplot or boxplot tool to visualize the rolling duration, frequency, and number of interruptions during rolling graphically.
Statistical analyses: The distribution of rolling parameters, such as rolling frequency and duration, often does not meet normality criteria, making parametric tests like Student’s t-test unsuitable. For comparing two groups, the non-parametric Wilcoxon Rank-Sum Test is preferred. When comparing more than two genotypes, the Kruskal-Wallis test serves as a non-parametric alternative to one-way ANOVA. For multiple comparisons, corrections should be made, typically using Dunn’s method as a post-hoc test. These statistical analyses can be conducted using common software like MATLAB, R, Jupyter Notebook, or similar (Figure 5).
Figure 5. Experimental results showing the percentage of L1–L2 animals of different genotypes that are able to complete at least one complete roll when a different subset of motor neurons (MNs) is acutely silenced using GtACR1. Silencing MN Is (R27E09>UAS-GtACR1-eGFP) or in combination with Ib MN1 (R27E09+RRa>UAS-GtACR1-eGFP) had little or no effect on rolling performance. Silencing Ib MNs of lateral transverse (LT) muscles (BH1>UAS-GtACR1-eGFP) leads to a slightly reduced chance of successful rolling. Silencing Ib MN of dorsal longitudinal (DL) muscles caused mild (CQ>UAS-GtACR1-eGFP) to moderate (Unc4AD-vGlutDBD>UAS-GtACR1-eGFP) defect to rolling; however, silencing both Ib and Is targeting DL muscles (CQ+RRa>UAS-GtACR1-eGFP) lead to severe rolling failure. Silencing Ib MNs of ventral longitudinal (VL) and dorsal oblique (DO) muscles (HB9>UAS-GtACR1-eGFP and vGlutAD-Nkx6DBD>UAS-GtACR1-eGFP) leads to near-complete failure in rolling.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Cooney et al. [9]. Neuromuscular basis of Drosophila larval rolling escape behavior. PNAS (Figure 4, panel B, C).
General notes and troubleshooting
General notes
While conducting optogenetics experiments, it is crucial to maintain uniform conditions across all genotypes. Variations in factors such as light luminance and ATR concentration can lead to inconsistent results. Therefore, standardizing these parameters ensures reliable and reproducible outcomes.
The dual activation and silencing technique using Chrimson and GtACR1 in this protocol can be applied to other experiments to investigate various neural circuits and behaviors. The protocol can also be expanded to include genetic tools like RNAi.
Troubleshooting
Problem 1: The smartphone camera cannot focus on the larva upon light stimulation.
Possible cause: The focus distance is not set up correctly in advance.
Solution: Most smartphone cameras feature auto-focusing when there is sufficient light. Before recording the experimental animals, adjust the distance between the objective and the gel pad with a wild-type larva with the LED light on until the camera stably focuses on the larva. Afterward, turn off the light to prepare the experimental larva without changing the camera settings on the phone. The camera should auto-focus on the rolling larva when the LED light is turned on.
Problem 2: The larva rolls out of the field of the recording.
Possible cause: The objective magnification level of the stereomicroscope is too high.
Solution: Reduce the zoom level. Alternatively, gently move the agarose gel pad between rolls to keep the animal in the center of the field. Do not move the gel pad before each roll is complete.
Problem 3: Difficulty locating the larva with the camera (especially for small, transparent L1 larvae).
Possible cause: The magnification level is low, or the agarose gel pad is too big for L1 larvae.
Solution: Increase the magnification level of the stereomicroscope. If there is difficulty navigating the gel when using a high magnification power, draw a small red mark with a Sharpie marker on the center of the gel. Then, put the larva near the mark, locate and focus on the red mark (and the larva) at a low magnification level, and zoom in.
Acknowledgments
Work in the Zarin lab is supported by Texas A&M grant. This protocol is derived from the original work of Cooney and Huang [9].
Competing interests
The authors declare no competing interest.
References
Tracey, W., Wilson, R. I., Laurent, G. and Benzer, S. (2003). painless, a Drosophila Gene Essential for Nociception. Cell. 113(2): 261–273.
Hwang, R. Y., Zhong, L., Xu, Y., Johnson, T., Zhang, F., Deisseroth, K. and Tracey, W. D. (2007). Nociceptive Neurons Protect Drosophila Larvae from Parasitoid Wasps. Curr Biol. 17(24): 2105–2116.
Grueber, W. B., Jan, L. Y. and Jan, Y. N. (2002). Tiling of the Drosophila epidermis by multidendritic sensory neurons. Development 129(12): 2867–2878.
Ohyama, T., Schneider-Mizell, C. M., Fetter, R. D., Aleman, J. V., Franconville, R., Rivera-Alba, M., Mensh, B. D., Branson, K. M., Simpson, J. H., Truman, J. W., et al. (2015). A multilevel multimodal circuit enhances action selection in Drosophila. Nature. 520(7549): 633–639.
Yoshino, J., Morikawa, R. K., Hasegawa, E. and Emoto, K. (2017). Neural Circuitry that Evokes Escape Behavior upon Activation of Nociceptive Sensory Neurons in Drosophila Larvae. Curr Biol. 27(16): 2499–2504.e3.
Burgos, A., Honjo, K., Ohyama, T., Qian, C. S., Shin, G. e., Gohl, D. M., Silies, M., Tracey, W. D., Zlatic, M., Cardona, A., et al. (2018). Nociceptive interneurons control modular motor pathways to promote escape behavior in Drosophila. eLife. 7: e26016.
Peron, S., Zordan, M. A., Magnabosco, A., Reggiani, C. and Megighian, A. (2009). From action potential to contraction: Neural control and excitation–contraction coupling in larval muscles of Drosophila. Comp Biochem Physiol A Mol Integr Physiol. 154(2): 173–183.
Clark, M. Q., Zarin, A. A., Carreira-Rosario, A. and Doe, C. Q. (2018). Neural circuits driving larval locomotion in Drosophila. Neural Dev. 13(1): 6.
Cooney, P. C., Huang, Y., Li, W., Perera, D. M., Hormigo, R., Tabachnik, T., Godage, I. S., Hillman, E. M. C., Grueber, W. B., Zarin, A. A., et al. (2023). Neuromuscular basis of Drosophila larval rolling escape behavior. Proc Natl Acad Sci USA. 120(51): e2303641120.
Article Information
Publication history
Received: Jun 21, 2024
Accepted: Oct 7, 2024
Available online: Oct 23, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Category
Neuroscience > Basic technology > Optogenetics
Neuroscience > Behavioral neuroscience > Sensorimotor response
Neuroscience > Sensory and motor systems > Animal model
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Quantitative Analysis of Kinetochore Protein Levels and Inter-Kinetochore Distances in Mammalian Cells During Mitosis
NW Neeraj Wasnik *
MS Mahima Singhal *
SK Sukirti Khantwal *
SM Sanghamitra Mylavarapu *
SM Sivaram V. S. Mylavarapu
(*contributed equally to this work)
Published: Vol 14, Iss 23, Dec 5, 2024
DOI: 10.21769/BioProtoc.5132 Views: 255
Reviewed by: Rajesh RanjanShanmugaPriyaa Madhukaran Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Oct 2021
Abstract
The mammalian kinetochore is a multi-layered protein complex that forms on the centromeric chromatin. The kinetochore serves as the attachment hub for the plus ends of microtubules emanating from the centrosomes during mitosis. For karyokinesis, bipolar kinetochore-microtubule attachment and subsequent microtubule depolymerization lead to the development of inter-kinetochore tension between the sister chromatids. These events are instrumental in initiating a signaling cascade culminating in the segregation of the sister chromatids equally between the new daughter cells. Of the hundreds of conserved proteins that constitute the mammalian kinetochore, many that reside in the outermost layer are loaded during early mitosis and removed around metaphase-anaphase. Dynamically localized kinetochore proteins include those required for kinetochore-microtubule attachment, spindle assembly checkpoint proteins, various kinases, and molecular motors. The abundance of these kinetochore-localized proteins varies at prometaphase, metaphase, and anaphase, and is thus considered diagnostic of the fidelity of progression through these stages of mitosis. Here, we document detailed, state-of-the-art methodologies based on high-resolution fluorescence confocal microscopy followed by quantification of the levels of kinetochore-localized proteins during mitosis. We also document methods to accurately measure distances between sister kinetochores in mammalian cells, a surrogate readout for inter-kinetochore tension, which is essential for chromosome segregation.
Key features
• Immunostaining of cultured and suitably fixed adherent mammalian cells growing as monolayers.
• Confocal fluorescence imaging for the purpose of fluorescence quantification.
• 2D and 3D image reconstruction and analysis of the acquired images using appropriate background correction and normalization.
• Quantification of the inter-sister kinetochore distances using 3D image reconstruction.
Keywords: Mitosis Kinetochore Prometaphase Metaphase Confocal fluorescence microscopy Fluorescence quantification
Graphical overview
Key steps involved in fluorescence quantification of spindle assembly checkpoint (SAC) protein loading at the kinetochores and in the calculation of inter-sister kinetochore distances. High-resolution confocal image datasets of cells subjected to appropriate treatment as per the experimental need (drugs, inhibitors, gene-specific siRNA, etc.) are used for quantifying the levels of SAC proteins at the kinetochores, measuring inter-sister kinetochore distances, or both. Quantification and analysis of both parameters are performed using image analysis software such as Fiji (open source) or the IMARIS software suite. Figure created with BioRender.com.
Background
The spindle assembly checkpoint (SAC) is the primary mechanism that ensures equal sister chromatid segregation [1–3]. The SAC monitors two major parameters during mitosis. The first is the end-on attachment of centrosome-derived microtubule plus ends to the kinetochores (Kt-MT attachment). The establishment of bipolar (amphitelic) Kt-MT attachment of a sister chromatid pair to opposite centrosomes, followed by the depolymerization of these kinetochore microtubules (kMTs), leads to poleward pulling of the kinetochores and the generation of inter-kinetochore tension, the second parameter sensed by the SAC. All other stochastically occurring forms of Kt-MT mis-attachment (monotelic, merotelic, and syntelic) fail to elicit inter-kinetochore tension [1].
The SAC operates through SAC proteins resident on the fibrous corona, the outermost and most transient kinetochore layer. The SAC generates a diffusible “wait-anaphase” signal triggered by the lack of attachment or mis-attachment at even one kinetochore, stalling the entire cell in metaphase and providing time for the rectification of Kt-MT mis-attachments [3]. Inter-kinetochore tension stabilizes Kt-MT end-on attachments and also initiates a molecular signaling cascade instrumental for the separation of sister chromatids toward the opposite spindle poles (centrosomes) due to kMT depolymerization. Inactivation or silencing of the SAC, achieved primarily through dynein-based poleward “stripping” of SAC proteins from kinetochores along spindle microtubules upon Kt-MT attachment or tension, is essential for anaphase onset [1,3].
The level of enrichment of SAC proteins, their receptors, and other associated proteins at the kinetochores can serve as biochemical markers of the exact stage of mitosis or be used as indicators of specific defects in mitotic progression [4]. High-resolution confocal microscopy has been used to reliably quantify these levels and correlate them with the stage of mitosis, mainly because the size of human kinetochores is not optically diffraction-limited [5]. Kinetochore protein level quantification methods were initially developed on individual, two-dimensional confocal micrographs of single z-planes, taking into account local background fluorescence corrections around the kinetochores [5] and have been used extensively since. In recent years, this concept has been built upon to evolve widely used methods employing modern microscopy analysis software packages and three-dimensional image reconstructions [6–8]. Similar tools have been used to quantify inter-kinetochore distances at prometaphase and metaphase. These distance measurements have proven to be reliable correlates of inter-kinetochore tension, especially at late metaphase prior to segregation [1]. This article details state-of-the-art methodologies for the quantification of levels of kinetochore proteins and distance measurements between sister kinetochores. The methods described can be adapted to most cultured cells/cell lines that can be imaged using high-resolution confocal fluorescence microscopy.
Materials and reagents
Reagents
Immunofluorescence labeling
Primary antibodies against the protein(s) of interest (the Mad1 example is shown here):
Mad1 (Thermo Fisher Scientific, catalog number: PA5–28185). Other alternatives compatible with immunofluorescence staining may be used after empirical optimization.
CREST (to stain kinetochores) (Antibodies Incorporated, catalog number: 15–234). Other alternatives staining the kinetochore or the centromere that are compatible with immunofluorescence staining may be used after empirical optimization for specificity.
α-Tubulin, DM1 alpha (Sigma, catalog number: T9026)
Secondary antibodies tagged with fluorophore:
Alexa Fluor® 488 AffiniPure Donkey Anti-Rabbit IgG (Jackson ImmunoResearch Inc., catalog number: 711–545-152)
Alexa Fluor® 594 AffiniPure donkey Anti-Mouse IgG (Jackson ImmunoResearch Inc., catalog number: 715–585-150)
CyTM 5 AffiniPure Donkey Anti-Human IgG (Jackson ImmunoResearch Inc., catalog number: 709–175-149)
Note: Other alternative fluorophore-conjugated secondary antibodies compatible with immunofluorescence staining may be used after empirical optimization.
Sodium chloride (NaCl) (SRL, catalog number: 64072); may be procured commercially from other standard sources
Potassium chloride (KCl) (SRL, catalog number: 38630); may be procured commercially from other standard sources
Disodium hydrogen phosphate (Na2HPO4) (SRL, catalog number: 1949144); may be procured commercially from other standard sources
Potassium dihydrogen phosphate (KH2PO4) (SRL, catalog number: 1649201); may be procured commercially from other standard sources
Triton X-100 (Sigma, catalog number: T8787–100ML)
Bovine serum albumin (BSA) (pH 6–7) (SRL, catalog number 9048–46-8); may be procured commercially from other standard sources
4',6-diamidino-2-phenylindole (DAPI), 5 mg/mL stock solution made in dimethyl sulfoxide (DMSO) (Thermo Scientific, catalog number: 62247)
Note: Alternatively, chromatin dyes for immunofluorescence such as Hoechst 33342 may be used after empirical optimization.
ProLong DiamondTM mounting media (Thermo Scientific, catalog number: P36970). Alternative antifade mounting media with a refractive index compatible with the lens and the lens immersion medium may be used after empirical optimization
Solutions
1× phosphate buffered saline (PBS) (see Recipes)
Blocking solution (see Recipes)
Recipes
1× phosphate buffered saline (PBS)
Components Amount/volume added Working concentration
NaCl 8 g 137 mM
KCl 0.2 g 2.7 mM
Na2HPO4 1.44 g 10 mM
KH2PO4 0.24 g 2 mM
Deionized water Make up the volume to 1 L
Blocking solution for immunofluorescence staining
Components Amount/volume added Working concentration
PBS 50 mL 1×
Triton X-100 250 µL 0.5%
BSA 0.5 g 1%
Weigh 0.5 g of BSA and add to 45 mL of 1× PBS in a 50 mL tube.
Place the tube on a rocker and mix at slow speed until completely dissolved. Avoid frothing of the solution.
Caution: High-speed mixing can denature proteins including BSA, of which frothing is a sign.
Add 250 μL of Triton X-100 to the solution.
Slowly mix the solution on a rocker to dissolve all the components completely.
Add 1× PBS to make up the volume to 50 mL and gently mix to homogeneity.
Laboratory supplies
Clean glass slides and coverslips
Critical: The coverslips should be properly cleaned and ideally be of category #1.5 (even thickness between 0.16 and 0.19 mm) to minimize chromatic aberration and refractive index mismatches between the lens, immersion medium, and sample mounting medium.
Equipment
Confocal image acquisition
Leica TCS SP8 laser scanning optical confocal microscope with HCX PL APO 63X-1.4 NA oil-immersion objective and a HyD (hybrid) or photomultiplier tube (PMT) detector.
Note: Equivalent confocal microscopes from standard commercial sources may be used, as long as they offer the following key features:
Plan apochromat (PL APO) confocal grade objectives with a high numerical aperture (1.2 or above).
Laser illumination in at least three distinct colors.
Emission pinhole(s) designed to block out-of-focus emission light from adjacent z-planes from being captured in the images.
In the case of raster laser scanning confocal scan-heads, high-sensitivity detectors such as PMTs, hybrid detectors, or avalanche photodiodes (APDs) with a quantum efficiency in the visible spectrum around 50 % may be used. Alternatively, for spinning disc confocal microscopes, high sensitivity cameras such as scientific complementary metal oxide semiconductor (sCMOS) cameras or electron multiplying charge-coupled device (EMCCD) cameras, with a quantum efficiency in the visible spectrum of around 80% or above, may be used.
Software and datasets
For image analysis
IMARIS software suite, version 8.3, Bitplane. Higher versions may be used.
Fiji (open-source image analysis software) (https://fiji.sc, Schindelin et al. [9])
For statistical analysis
GraphPad Prism (GraphPad Software, USA). Other standard software packages may be used.
Procedure
Cell fixation and immunofluorescence labeling
Note: Prior to fixation and immunofluorescence labeling, cells may be subjected to treatment as per experimental needs. For instance, siRNA-mediated protein depletion or using specific inhibitors against the protein of interest and subsequent scoring for mitotic phenotype(s) may suggest its role in cell division.
After treatment of the cells as per experimental requirements, remove the coverslips containing the cells and place them in a fresh 6-well plate with the surface containing the cells facing upward for further processing.
Wash the cells 2–3 times with 2 mL of 1× PBS each time.
Add 2–3 mL of pre-chilled (at -20 °C) absolute methanol (AR grade or HPLC grade) to the coverslips to fix the cells.
Immediately place the 6-well plate at -20 °C for a minimum of 30 min to complete the fixation process.
Note: The coverslips may be stored at -20 °C submerged in absolute methanol for an extended period of time without letting the methanol dry out.
Note: An alternate method of fixation has been described in the literature for mammalian cell kinetochores, using 4% formaldehyde and a buffer consisting of PIPES-HEPES-EDTA and magnesium chloride (PHEM) [10]. This method may be optimized and alternatively used. If this method of fixation is used, the following immunostaining steps must be immediately performed, since aldehyde fixation does not remain stable over long periods. Additionally, only 4% paraformaldehyde in 1× PBS may also be used for fixation prior to immunostaining [7, 8, 11].
Rehydrate the cells by adding adequate 1× PBS and wash 3–4 times (2 mL each time) to ensure complete rehydration.
Use a clean, fine-tipped pair of forceps to lift the coverslips and gently place them cell-side up on a fresh ParafilmTM sheet inside a humidified chamber prepared by lining a large cell culture dish with a damp paper towel and the lid closed.
Caution: All following steps are performed in the humidified chamber.
Add approximately 50–100 μL of blocking solution on each coverslip to cover the entire surface and leave at room temperature for 1 h. Only enough blocking solution should be added to form a large drop covering the entire coverslip.
Prepare the primary antibody solution by diluting all required primary antibodies (caution: raised in different animals) in blocking buffer in a fresh microcentrifuge tube (Mad1: 1:200, CREST: 1:100, α-tubulin: 1:800) at room temperature.
After 1 h of blocking, gently aspirate off the blocking solution from the edge of the coverslip to remove the blocking solution, add approximately 40–50 μL of diluted primary antibody to each coverslip, and incubate for 1 h at room temperature.
Note: 40 μL of the diluted antibody solution is sufficient to cover a 12 mm circle coverslip.
Remove the antibody solution and wash the cells thrice with blocking solution.
Dilute fluorophore-conjugated secondary antibodies (Alexa Fluor® 488 anti-rabbit IgG: 1:800, Alexa Fluor® 594 anti-mouse IgG: 1:800, and CyTM 5 anti-human IgG: 1:800) in a single microcentrifuge tube containing blocking solution and add 40–50 μL to each coverslip. Incubate at room temperature for 1 h.
Aspirate off the secondary antibody solution and wash the cells with 1× blocking solution thrice.
Wash cells with 1× PBS once to remove the blocking solution.
Add 50 μL of diluted DAPI (1:10,000 of a 5 mg/mL stock made in DMSO) and incubate for a maximum of 2 min at room temperature.
Aspirate off the DAPI solution and wash the cells thrice with 1× PBS.
Wash the cells once with de-ionized water to remove the PBS.
Note: The salts in the PBS can cause noise during fluorescence/confocal imaging and must therefore be removed.
Label a clean, frosted glass slide on the frosting using a lead pencil. Place 5 µL of ProLong DiamondTM mounting media on it. Gently lift the coverslips using forceps and drain any excess liquid by touching the bottom edge of the vertically held coverslip on a piece of lint-free tissue paper. Place the coverslip on the drop of mounting media (cell-side down) by laying it down gently from one edge and letting the weight of the coverslip spread the mounting media under it as it settles. Remove any trapped air bubbles between the coverslip and the slide by pushing them outward with gentle pressure on the coverslip using the tip of the forceps.
Allow the mounted coverslips to air-dry overnight at room temperature, protected from light on a flat, horizontal surface.
Caution: This step shouldnotbe performed in the humidified chamber.
The dried coverslips may be imaged immediately on a confocal microscope or stored at -20 °C in a slide box for later use.
Image acquisition
Acquire high resolution images of prometaphase and metaphase cells on a Leica TCS SP8 laser scanning optical confocal microscope (or equivalent) using an HC PL APO 63×-1.4 NA oil-immersion objective and a HyD (hybrid) or PMT detector.
Keep the imaging parameters constant for all experimental conditions using the Leica LASX software (or other acquisition software, as applicable).
Keep the zoom constant at 4.5× for imaging mitotic cells (both metaphase and prometaphase).
Use simultaneous scanning sequences for two different wavelength channels for image acquisition. Ensure that the emission wavelength of one fluorophore does not cross-excite any other fluorophore. For example, in scan sequence 1, use laser lines for 405 nm and 594 nm, and in scan sequence 2, use laser lines for 488 nm and 647 nm.
Note: Alternatively, image acquisition can be performed using sequential scanning to eliminate the possibility of inadvertent fluorophore cross-excitation. However, the scanning sequence should begin at the highest wavelength (647 nm) and end at the lowest wavelength (405 nm).
Critical: An important consideration during imaging for fluorescence quantification is to never expose the fluorophore tagged to the protein of interest while focusing or locating cells prior to image acquisition. This is important to prevent bleaching of the fluorophore when the laser hits the sample and may therefore impact accurate quantitative measurements.
Adjust the oversaturation and undersaturation of the emission signals using the Quick Look-up Table (QLUT) or an equivalent feature to stay within the dynamic range of the detectors (Table 1).
Table 1. Imaging parameters used during acquisition
Checklist of imaging parameter settings to be ensured before imaging
S. no. Setting name Imaging parameters Remarks
1 Scan Mode xyz
2 Imaging resolution 1024 × 1024 The images should be acquired at high resolution for effective 2D and 3D quantification.
3 Scan Direction X Bidirectional For faster coverage of the imaged field.
4 Scan Speed 200 Hz Higher frequency (scan speed) enables faster coverage of the imaged field but can compromise image resolution. Caution: For kinetochore fluorescence quantification through fixed cell imaging, image resolution is more important than imaging speed. The software should clearly be able to tell the kinetochore apart from the background (this must be manually ascertained).
5 Magnification 63× Other companies use Plan Apochromat lenses of 60×/64× magnification, for which the parameters would remain similar.
6 Objective name and numerical aperture HC PL APO CS2 63×/1.40 NA (numerical aperture) oil immersion lens A higher NA lens provides better resolution.
7 Immersion Oil (Leica type F immersion liquid, catalog number: 11513859) An equivalent immersion oil/immersion liquid from other manufacturers may also be used.
8 Zoom [Region of Interest (ROI) imaging] 4.5 Keep the zoom factor such that only the dividing cell is imaged (zoom factor was kept at 4.5 in the Leica TCS SP8 microscope). In a laser scanning microscope, this reduces the area of the field that needs to be scanned, leading to faster imaging.
10 Pinhole diameter 0.5 Airy Units (AU) The pinhole diameter is usually set at 1 AU for most confocal microscopes. For kinetochore imaging, which usually shows bright fluorescence emission, a lower pinhole diameter (such as 0.5 AU) increases the x-y resolution. Caution: Lower pinhole diameters cannot be used for weak fluorescence intensity/signals. A good primary antibody that gives a strong and specific fluorescence signal must be used for immunostaining.
11 Scanning sequence 1 488 nm for antigen to be quantified and 647 nm for CREST The two fluorophores can be simultaneously excited, and the emission signals simultaneously captured on two separate detectors.
12 Scanning sequence 2 405 nm for DAPI and 594 nm for microtubules The two fluorophores can be simultaneously excited, and the emission signals simultaneously captured on two separate detectors.
Use an imaging pixel format of 1024 × 1024 and a scan speed of 200 Hz for image acquisition. The higher pixel format enables higher image resolution and subsequent quantification of fluorescence intensities with greater accuracy.
If required, reduce the pinhole diameter to 0.5 Airy units (AU) for better structural definition of the kinetochores.
Note: Reducing the pinhole omits low emission intensities and can therefore be used only when the emission signal is strong.
Keep the z-step size at 0.3 μm.
Start image acquisition after ensuring all settings are accurate.
Acquire z-sections (optical depth section images) of each cell in steps of 0.3 μm each, starting from the first out-of-focus plane on one side of the cell, optically sectioning through the cell, and ending at the first out-of-focus z-plane on the other end of the cell.
Image analysis
Quantification of SAC proteins at the kinetochore can be achieved using 3D reconstruction of the z-slices in the IMARIS software suite. Alternatively, this can also be performed from 2D images as described in Hoffman et al. [5] using Fiji, Metamorph, or other software packages. Here, we describe the quantification of fluorescence intensities at the kinetochore both in 2D as well as in 3D. The schematics of the workflow are represented as flowcharts for fluorescence intensity measurements in 2D (Figure 1) and 3D (Figure 2).
Figure 1. Schematic flowchart for fluorescence intensity measurement in 2D. Open the image in Fiji, select the z-plane with the brightest signal for SAC protein, zoom into the kinetochore area, draw a circular ROI (0.45 μm diameter) around the CREST signal, draw a circular ROI (0.63 µm diameter) around the CREST signal for background correction, measure fluorescence intensity (Analyze > Measure), and export all intensity values as an MS Excel spreadsheet for further analyses. Figure created with BioRender.com.
Figure 2. Schematic flowchart for fluorescence intensity measurement in 3D. Launch IMARIS software suite, open the image file, select the image to be analyzed, select the channel to be used for analysis, crop the cell in 3D, open the spot function, assign the spot size and add new spot, check the background subtraction box and click the next button, click the statistical spreadsheet followed by detailed button, and export all intensity values as an MS Excel spreadsheet for further analyses. Figure created with BioRender.com.
The quantification of SAC protein levels at the kinetochore using 2D imaging (Fiji) is illustrated in Figure 3 with the following steps:
For quantification of SAC proteins at the kinetochore, consider cells that have clearly defined kinetochore immunofluorescence (as visualized using the CREST stain).
Open the selected cells in Fiji. Choose the z-plane(s) containing the visibly brightest signal for Mad1 (the SAC protein example illustrated here) by going through the entire z-stack and selecting the channels for CREST (resident kinetochore protein) and Mad1 for quantification.
Zoom into the kinetochore (chromosomal) region of the cell. It is important to maintain the same zoom for all channels to ensure the regions of interest (ROI) across channels have the same XY dimensions.
Draw a circular ROI of diameter 0.45 μm encompassing the CREST signal using the circular drawing tool from the Fiji menu bar and designate it as the inner circle.
Critical: The average size may be empirically determined for individual cell lines using several control kinetochores across multiple cells to capture the CREST (kinetochore) signal and the signal of the loaded kinetochore SAC protein. SAC proteins at kinetochores may not always be seen overlaid on the same exact pixels as the kinetochore antigen. The loading of these proteins can accumulate significantly on laterally adjacent pixels attached to the kinetochore, especially in prometaphase.
Draw a larger circle of 0.63 μm diameter (approximately double the area of the inner circle) around the inner circle and designate it as the outer circle. The fluorescence intensity from the outer circle will be used for calculating the local background during quantification.
Figure 3. Fluorescence quantification of SAC protein loading at the kinetochore in 2D using Fiji image analysis software. The window on the far left is the confocal micrograph showing kinetochores (pseudo-colored in magenta). An ROI is drawn around the kinetochore using the tool shown in the red box. The diameter of each circular ROI is assigned in the window labeled Specify. The box labeled Scaled units (microns) is checked to convert the measurement of the circular ROI to micrometers. Each ROI is added to the ROI manager as shown in the window on the far right and designated as either outer circle or inner circle. The yellow dotted box represents ROIs drawn around each kinetochore to be quantified; the inset of the same represents the zoomed image.
Once all ROIs are drawn, measure the fluorescence intensities using the Analyze > Measure tool on the Fiji menu bar. This generates a data sheet for all fluorescence intensities and can be exported as an MS Excel spreadsheet for further calculations.
Paste the ROIs from the CREST channel on the Mad1 channel and obtain fluorescence measurements for the Mad1 channel as described in step C6.
After exporting the fluorescence intensities from all channels to an MS Excel spreadsheet, perform the following calculations.
For calculating the fluorescence intensity of the region between the inner and the outer circles (FS), subtract the integrated fluorescence of the inner circle (FI) from the integrated fluorescence of the outer circle (FO).
F
S
=
F
O
-
F
I
Next, calculate the area of the region between the inner and outer circles (AS) by subtracting the area of the inner circle (AI) from the area of the outer circle (AO).
AS = AO - AI
Calculate the background intensity (FBackground) using the equation below:
FBackground = FS (AI/AS)
The background corrected fluorescence intensity of CREST is obtained by subtracting the background intensity (FBackground) from the intensity of the inner circle (FI).
FCREST = FI - FBackground
Calculate the final fluorescence intensity of Mad1 (FMad1) in the same manner by following steps C10–12 for the Mad1 channel.
Normalize the Mad1 intensity to the CREST intensity by dividing the background-corrected Mad1 intensity by the background-corrected CREST intensity for each kinetochore.
Mad1:CREST = FMad1/FCREST
The normalized ratio of Mad1:CREST is plotted as a scatter plot using GraphPad Prism.
Note: Other kinetochore resident proteins such as CENP-A and CENP-C (Cmentowski et al [12]) are also commonly used as kinetochore normalization markers. The underlying assumption is that the kinetochore structure and organization (inner and outer kinetochore layers) do not change upon SAC protein loading on the outermost fibrous corona in mitosis.
Quantification of inter-kinetochore distance (using Fiji)
Sequence of steps for fluorescence intensity measurement in 2D: Open image in Fiji > select z-plane with brightest signal for SAC protein > Analyze > line draw tool > draw line joining centroids of sister kinetochores > analyze > measure > export fluorescent intensities values as MS Excel spreadsheet for further analyses.
Inter-kinetochore distance is measured using Fiji in 2D.
Consider only those cells for quantification that have a tightly formed metaphase plate and in which most kinetochore pairs are aligned almost parallel to the spindle axis (the line connecting the two spindle poles). This is indicative of the late metaphase stage, where most kinetochores have captured microtubules and sufficient inter-kinetochore tension has been generated.
Select a single z-plane containing the highest number of visibly brightest CREST signals by moving across the z-stack.
Select the line draw tool from the Analyse tab on the Fiji menu bar and draw a straight line joining the centroids of a pair of kinetochores on the sister chromatids. This is illustrated in Figure 4.
The morphological features that characterize mammalian kinetochores are a faint, comet-like tail of CREST intensity connecting the two kinetochores. A pair of sister kinetochores (located on sister chromatids) is determined by moving through the z-stack. Adjacent CREST signals that come into and go out of focus in the same frame while moving through the z-stack are presumed to be a pair and therefore may be considered for distance measurements.
The distance between the pair of kinetochores is measured by selecting the Analyze > Measure tool on the Fiji menu bar. This generates a data sheet for inter-kinetochore distance in micrometers and can be exported as an MS Excel spreadsheet.
Twenty pairs of kinetochores per cell over a total of 20 cells per replicate are analyzed in this manner to obtain statistically significant data.
The distance measurements between kinetochore pairs from a set of experiments are plotted as a scatter plot using the GraphPad Prism software.
Figure 4. Quantification of inter-kinetochore distance in 2D using Fiji image analysis software. The window on the far left is the confocal micrograph showing kinetochores (pseudo-colored in magenta). A line connecting the centers of each pair of kinetochores (ROI) is drawn using the line tool (red box). The yellow dotted box highlights the representative lines drawn to connect the centers of two kinetochores, whose inset represents the zoomed form. Each ROI is added to the ROI manager (window on the far right) for calculating the inter-kinetochore distance.
Quantification of SAC protein(s) at the kinetochore in 3D
A flowchart depicting fluorescence intensity measurement steps in 3D is shown in Figure 2.
The IMARIS software suite is used for fluorescence quantification of SAC proteins at the kinetochore.
Launch the IMARIS software suite and open the acquired images using the Open tab on the menu bar as indicated in Figure 5 (yellow box).
A window containing all the acquired images appears. Open the image file from the list in the dialogue box as shown in figure 6.
Upon opening the selected image file, a 3D rendered image containing the xyz coordinates and all channels for that image (405 nm, 488 nm, 594 nm, and 647 nm) is displayed as below (Figure 7). Since channels for DAPI (405 nm) and microtubules (594 nm) are not considered for quantification of SAC proteins at the kinetochore, turn off these channels. Now, only signals for CREST in magenta (594 nm) and Mad1 (SAC protein) in green (488 nm) are displayed (Figure 8).
Select the Crop 3D option from the menu bar for cropping a cell in 3D. Carefully select the cell of interest using the selection tool, ensuring the exclusion of unwanted neighboring or background cells. Apply the cropping function by clicking OK to display only the 3D-cropped image (Figure 9).
Select the Spot function tab and press add new spot button as indicated in Figure 10 (yellow box). Once the Spot function is activated, a new window appears, as indicated in Figure 10 (yellow box).
Figure 5. The graphical user interface (GUI) of the IMARIS software suite. The yellow dotted box and inset show the menu bar functions for opening the confocal image files for initiating the analysis.
Figure 6. GUI of the IMARIS software suite showing the acquired confocal image files, enabling the user to select an image for analysis and open the selected file from the displayed dialogue box (yellow dotted box).
Figure 7. 3D reconstruction of the acquired confocal image displaying the xyz spatial coordinates and fluorescence channel information: Mad1 (green), CREST (magenta), DAPI (blue), and microtubules (red).
Figure 8. 3D cropping of the selected image. Click the edit button on the menu bar and click on the Crop 3D button from the dropdown menu for cropping the cell of interest in 3D. This function retains the x, y, z information of the selected image upon cropping.
Figure 9. 3D cropping of the selected image. The white dotted box represents the region containing the cell of interest to be cropped in 3D. The OK button at the bottom right (yellow dotted box) enables the cropping of the cell of interest.
Figure 10. Activating the spot function on the IMARIS GUI
The channel for CREST (magenta) is selected, and the spot size of 0.45 μm is designated for kinetochores of HeLa cells. Also, the background subtraction box is checked to ensure that the software automatically subtracts the background fluorescence in the processed image (figure 11).
Figure 11. Assigning the spot size for kinetochores. Kinetochores are assigned a spot size of 0.45 μm in the selected channel for CREST (magenta) in HeLa cells. The average size may be empirically determined for individual cell lines using several control kinetochores across multiple cells to capture the CREST (kinetochore) signal and the signal of the loaded kinetochore SAC protein.
The software automatically picks CREST-positive spots and displays them as grey balls. Spurious spots may also be occasionally picked by the software but can be manually deleted by adjusting the slider for the intensity (Figure 12, yellow inset box at the bottom). Conversely, spots that are clearly identified as kinetochores but may have been missed by the software can be added manually using the slider shown in the yellow dotted box in Figure 12.
Figure 12. Kinetochore spots of 0.45 μm diameter as rendered in grey by the spot function on IMARIS. The yellow dotted box displays the interactive slider for selectively removing non-specific spots.
Once all kinetochores have been selected, click the Next button to convert the grey spots to magenta as indicated in Figure 13.
Figure 13. Finalizing the selected kinetochore spots
Click the statistical spreadsheet button followed by the detailed option button to simultaneously obtain the intensity sum values for the selected kinetochores (CREST) as well as the SAC protein loaded on the kinetochores (Mad1) (Figure 14, yellow dotted box).
Figure 14. Fluorescence intensity sum values of the CREST and Mad1 channels are displayed in the yellow dotted box.
Integrated intensity sum values for each spot for CREST and Mad1 are exported as an MS Excel spreadsheet for further calculations, as indicated in Figure 15.
Figure 15. Screenshot of MS Excel spreadsheet displaying the quantified integrated fluorescence intensity measurements extracted for CREST and Mad1 as measured by IMARIS.
For quantifying the level of Mad1 at the kinetochore, the ratio of the fluorescence intensities of Mad1:CREST is calculated for each spot by dividing the fluorescence intensity of Mad1 by that of CREST. These values are then sorted from high to low and the top 20 kinetochore intensity values are considered for each cell. These calculations are performed for at least 20 cells per replicate to obtain robust, statistically significant data.
Quantification of inter-kinetochore distance in 3D
Import the acquired raw image file into the IMARIS software suite. Access the file menu located in the upper left corner. Click on Open file to open the confocal image files containing all the acquired images and select and open the individual z-image to be quantified (single confocal plane). Click the OK button to open the selected image (yellow box, Figure 16).
Figure 16. IMARIS GUI displaying the confocal image file containing all acquired images. An individual image can be selected and opened by clicking on the image followed by clicking OK.
Select the Slice mode from the software menu bar. Since only the kinetochore channel (CREST, 647 nm) is required for quantification of inter-kinetochore distances, turn off all the other channels by un-checking the individual boxes as shown in Figure 17 (yellow dotted box).
Figure 17. The presented image depicts a 2D confocal microscopy reconstruction, visualized using x, y, and z spatial coordinates. The image displays various fluorophore markers: Mad1 (green), CREST (magenta), DAPI (blue), and microtubules (red).
Crop the cell of interest by selecting Crop 3D from the Edit button on the menu bar as described in step E5.
Zoom into the chosen metaphase cell such that the individual kinetochores are clearly visible. The morphological features that characterize mammalian kinetochores are a faint, comet-like tail of CREST intensity connecting the two kinetochores. A pair of kinetochores of sister chromatids is determined by moving through the z-stack. Adjacent CREST signals that come into and go out of focus at the same time while moving through the z-stack are presumed to be a pair and therefore may be considered for distance measurement (Figure 18).
Figure 18. The yellow box depicts a metaphase cell containing pairs of sister kinetochores aligned at the metaphase plate
Click on the center of each kinetochore pair. A line joining the two sister kinetochores appears as shown in Figure 19. The inter-kinetochore distance (here, 0.922 μm) appears in the dialogue bar in the upper-left corner (yellow box). Repeat for each kinetochore pair for all experimental conditions and record the values in an MS Excel spreadsheet. A minimum of 20 kinetochore pairs from each cell and 20 cells from each replicate are measured to obtain statistically significant data.
Figure 19. A specific z-plane is identified where two CREST spots from a pair of kinetochores are clearly discernible within the yellow dotted box. This z-plane is selected for the quantification of interkinetochore distance. The measured distance between the kinetochores of a pair is displayed in the box demarcated by the white dotted line (top left).
The measurements obtained following the described method are represented as a scatter plot using the GraphPad Prism software.
Data analysis
Statistical analysis
A minimum of 20 kinetochore pairs from each cell and 20 cells from each replicate are analyzed for robust statistics as indicated in steps D7, E12, and F5. The distribution of the data is tested for normality using tests such as the D’Agostino–Pearson through the GraphPad Prism software. Parametric tests are used for normal data distribution (Student’s t-test/one-way ANOVA), while non-parametric tests are used for non-normal data distribution (Mann-Whitney U/Kruskal–Wallis) to statistically analyze the data and calculate statistical significance using the Prism software. Graphs are generated using the GraphPad Prism software. Error bars can be represented as standard deviation (SD) or standard error of mean (SEM) from at least three independent experiments (biological replicates)
Validation of protocol
Validation of quantitative analyses of fluorescence intensity
The methods described in this paper are widely accepted and have been used in the fields of kinetochore and mitosis biology for over twenty years [2, 5–8, 12–15]. However, users of either method may choose to cross-validate them. For instance, data using 2D quantification [13] and 3D quantification [6,7] of SAC proteins (Zw10, Mad1) upon depletion of the same gene (hLIC1) in the same cell type (HeLa) show reproducible phenotypes across the two methods.
General notes and troubleshooting
General notes
Cell fixation and immunofluorescence labeling
Mitotic cells are very loosely adherent to the substratum (coverslips). Therefore, very gentle washing using 1× PBS is recommended to avoid detachment of the mitotic cells.
Chilled methanol-fixed coverslips can be stored at -20 °C for an extended time as long as they stay immersed in chilled methanol throughout. However, if cells are fixed using aldehydes (4% formaldehyde or PHEM buffer), it is recommended that immunofluorescence staining be performed immediately.
After fixation using chilled methanol, rehydrate the cells adequately using 1× PBS for optimal immunostaining.
Prior to mounting the stained coverslips in mounting medium, it is crucial to remove all traces of PBS, as the salt crystals formed on the coverslips during the drying process may auto-fluoresce upon exposure to laser light and interfere during image acquisition.
Analysis from unevenly stained samples should be avoided. It is advisable to repeat the experiment and stain again to obtain even staining.
Image acquisition
If high background fluorescence is observed, a new experiment with a reduced concentration of the primary antibody is recommended for staining (needs to be empirically optimized). This is likely to reduce the non-specific signal.
If antibody signals remain non-specific, the use of fluorescently tagged constructs can be explored, with appropriate negative controls to rule out tag-induced artifacts.
Imaging parameters should be kept constant for all experimental conditions to ensure unbiased fluorescence quantification and data interpretation.
A zoom factor of 4.5 is optimal for imaging mitotic cells at sub-cellular resolution when using a 63× magnification plan apochromat lens.
While setting up simultaneous scanning sequences for multiple fluorophores, it is critical to ensure that the excitation and emission wavelengths of the two fluorophores are sufficiently far apart. This will minimize the chances of inadvertent cross-excitation of the lower energy fluorophore by the fluorescence emission of the higher energy fluorophore.
It is important to never expose the fluorophore channel of interest (which is to be quantified) to the laser while locating or focusing the field of cells to prevent any photo-bleaching of the fluorophore. This may significantly impact fluorescence quantification and data interpretation.
Oversaturation as well as undersaturation of fluorescence signals should be avoided during image acquisition. The Quick Look Up Table (QLUT, for Leica confocal microscopes) or an equivalent feature in other machines should be used to optimize the excitation light intensity and the detector gain voltage, in order to stay within the dynamic range of the detector.
The confocal pinhole aperture is normally set at 1 Airy unit. However, for strong emission signals, it may be adjusted to a lower value (such as 0.5 Airy units) to increase the lateral (x-y) resolution of the kinetochores being imaged.
In case a low emission signal-to-noise ratio (SNR) is observed, the following steps can be taken to improve the SNR: i) Increase detector gain voltage (detector sensitivity), not exceeding 700–800 V. Try not to increase the laser illumination intensity to avoid bleaching. ii) If required, increase the laser illumination intensity through the acoustic-optical tunable filter (AOTF) to a level where the SNR improves but bleaching does not happen.
Acknowledgments
The authors express gratitude to the Regional Centre for Biotechnology, India, for infrastructure, cell culture, and imaging facilities. N.W., M.S., and S.K. are supported by doctoral research fellowships from the Indian Council of Medical Research (ICMR) India, the Council for Scientific and Industrial Research (CSIR) India and the Department of Biotechnology (DBT) India, respectively, while S.M. is supported by a Women-in-Science (WOS-A) grant from the Department of Science and Technology (DST) India. This work was supported by funding to SVSM from the Department of Biotechnology (DBT; grant BT/PR6420/GBD/27/435/2012), the Science and Engineering Research Board (grants EMR/2016/007842 and CRG/2021/006858) and the Regional Centre for Biotechnology.
Author contributions: N.W., M.S., S.K., and S.M. conducted the experiments and analyzed data. N.W., M.S., and S.K. prepared figures. S.M. wrote and edited the manuscript. SVSM conceived, designed, supervised the study and edited the manuscript.
Competing interests
Authors declare no conflicts of interest or competing interests.
Ethical considerations
This study did not require any animal or human ethics clearances. The study was performed with due approval from the institutional biosafety committee of the Regional Centre for Biotechnology.
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Article Information
Publication history
Received: Apr 8, 2024
Accepted: Sep 30, 2024
Available online: Nov 4, 2024
Published: Dec 5, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Cell Biology > Cell imaging > Confocal microscopy
Cell Biology > Cell-based analysis
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5,133 | https://bio-protocol.org/en/bpdetail?id=5133&type=0 | # Bio-Protocol Content
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Microdissection and Single-Cell Suspension of Neocortical Layers From Ferret Brain for Single-Cell Assays
LD Lucia Del-Valle-Anton
SA Salma Amin
VB Víctor Borrell
Published: Vol 14, Iss 24, Dec 20, 2024
DOI: 10.21769/BioProtoc.5133 Views: 259
Reviewed by: Pilar Villacampa Alcubierre Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Mar 2024
Abstract
Brain development is highly complex and dynamic. During this process, the different brain structures acquire new components, such as the cerebral cortex, which builds up different germinal and cortical layers during its development. The genetic study of this complex structure has been commonly approached by bulk-sequencing of the entire cortex as a whole. Here, we describe the methodology to study this layered tissue in all its complexity by microdissecting two germinal layers at two developmental time points. This protocol is combined with a step-by-step explanation of tissue dissociation that provides high-quality cells ready to be analyzed by the newly developed single-cell assays, such as scRNA-seq, scATAC-seq, and TrackerSeq. Altogether, this approach increases the resolution of the genetic analyses from the cerebral cortex compared to bulk studies. It also facilitates the study of laboratory animal models that recapitulate human cortical development better than mice, like ferrets.
Key features
• Microdissection of individual germinal layers in the developing cerebral cortex from living brain slices.
• Enzymatic and mechanical dissociation generates single-cell suspensions available for high-throughput single-cell assays.
• Protocol optimized for embryonic and early postnatal ferret cortex.
Keywords: Cerebral cortex Microdissection Single-cell suspension Ventricular zone Outer subventricular zone Splenial gyrus Lateral sulcus Ferret Cell concentration Cell viability
Graphical overview
Background
The field of cerebral cortex development has witnessed milestone breakthroughs over the last 15 years, including the discovery of new types of progenitor cells such as basal radial glia (bRG) [1–5] and truncated radial glia (tRG) [6–8]. The discovery of new neural progenitor cell types has derived from more detailed characterizations of these cells, revealing for example that they dynamically modify their morphology along development, defining cell morphotypes [9–11]. The lineage relationships between this wide repertoire of progenitor cell morphotypes are also variable and dynamic, in line with the complexity of intrinsic and extrinsic factors regulating this process [9]. This complexity is greater in mammals with a folded cortex, like humans or ferrets, compared to species with a smooth cortex, like mice [8].
Traditionally, transcriptomic analyses of the developing cerebral cortex were possible only in bulk, considering it a relatively simple and homogeneous structure in spite of being clearly layered and anatomically subdivided. However, the emergence of single-cell sequencing technologies (i.e., scRNA-seq) has allowed the transcriptomic characterization of individual cell types with unprecedented detail and the discovery of unsuspected cell diversity also in the cerebral cortex. Nevertheless, scRNA-seq analyses of the developing cortex are still largely analyzed in bulk, relying on a limited number of specific, presumed marker genes to identify cell types. An increasing number of studies question if such marker genes are reliable for studying cell diversity in the developing cortex, particularly neural progenitor cells, and if they are faithful across mammalian species [8,9,12–15]. This protocol represents a milestone for studies of cortex development, where microdissection of individual layers of the developing cortex allows grasping its full complexity. By taking advantage of the extensive knowledge of the biology of cortical progenitor cells, the microdissection of germinal layers presented here allows distinguishing progenitor cell populations based on their layer of residency (i.e., apical versus basal progenitors), independent from the use of marker genes that must be assumed specific to particular cell types. Furthermore, the generation of a single-cell suspension from these microdissected tissues enables the use of massively parallel single-cell assays, including scRNA-seq or scATAC-seq, as well as TrackerSeq for cell lineage tracing. Hence, the combination of microdissection of individual germinal zones with single-cell approaches vastly increases the power of these analyses compared to studies of bulk tissue. Last, this protocol facilitates the study of animal models that are less common and more complex than mice, such as ferrets, with features of cortical development similar to humans. Unfortunately, the dissociation of cortical tissue into individual cells bears inherent limitations, such as the impossibility of cell morphology assessment, which currently limits the association of the new data with previous characterizations of progenitor cell morphotypes.
The specific details and conditions presented in this protocol have been optimized for the cerebral cortex of developing ferrets; nevertheless, it can also be implemented to study other layered brain regions, such as the hippocampus, olfactory bulb, or superior colliculus. The study of these structures by microdissection of individual layers, combined with the ever-increasing possibilities of single-cell assays, will significantly contribute to redefining cell type diversity in the brain.
Materials and reagents
Biological materials
Pregnant sable ferret (Mustela putorius furo) (Euroferrets, Denmark)
Reagents
Blue ultrasound gel (Transonic, catalog number: TOG04)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S3264)
Potassium chloride (KCl) (VWR, catalog number: 26759.291)
Potassium dihydrogen phosphate (KH2PO4) (VWR, catalog number: 26925.295)
MACS neural tissue dissociation kit (P) (Miltenyi Biotec, catalog number: 130-092-628)
Leibovitz’s 1× L-15 medium with L-glutamine, without phenol red (Gibco, catalog number: 21083027)
SeaPlaque agarose (Lonza, catalog number: 50100)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906)
1× Phosphate buffered saline (PBS) without CaCl2/MgCl2 pH 7.4 (Gibco, catalog number: 10010023)
RNaseZap RNase decontamination solution (Invitrogen, catalog number: AM9780)
Anesthetic/analgesic cocktail: xylazine (Karizoo, trade name: Xylasol 20 mg/mL, catalog number: 579700.7), ketamine (VetViva Richter, trade name: Ketamidor 100 mg/mL, catalog number: 580393.7), buprenorphine (VetViva Richter, trade name: Bupaq 0.3 mg/mL, catalog number: 578816.6)
Saline solution (NaCl 0.9%) (Interapothek, catalog number: 208978)
1× Hank’s balanced salt solution (HBSS) without CaCl2/MgCl2/phenol red (Gibco, catalog number: 14175095)
Ethanol absolute (VWR, catalog number: 20821.365P)
0.4% Trypan blue stain (Invitrogen, catalog number: T10282)
Solutions
10× Phosphate buffered saline (PBS) stock solution (see Recipes)
1× PBS (see Recipes)
Cell collection medium (see Recipes)
Recipes
10× PBS stock solution (1 L)
Reagent Final concentration Quantity or Volume
NaCl 1.37 M 80 g
Na2HPO4 100 mM 14.4 g
KCl 27 mM 2 g
KH2PO4 18 mM 2.4 g
Distilled H2O n/a 1 L*
Total n/a 1 L
*Dilute reagents in 800 mL, bring solution pH to 7.4, and top off to 1 L.
1× PBS (1 L)
Reagent Final concentration Quantity or Volume
PBS 10× 1× 100 mL
Distilled H2O n/a 900 mL
Total n/a 1 L
Cell collection medium (50 mL)
Reagent Final concentration Quantity or Volume
BSA 0.04% (w/v) 0.02 g
PBS (without CaCl2/MgCl2, pH 7.4) 1× 50 mL
Total n/a 50 mL
The medium should be freshly prepared and filtered on the day of the experiment.
Laboratory supplies
Glass Pasteur pipettes (Fisher Scientific, catalog number: FB50253)
0.5 mL PCR tubes (Eppendorf, catalog number: 0030124537)
2 mL DNA LoBind tubes (Eppendorf, catalog number: 0030108078)
15 mL centrifuge tubes (Corning, catalog number: 430791)
50 mL centrifuge tubes (Corning, catalog number: 430829)
40 μm PluriStrainer mini cell strainers (PluriSelect, catalog number: 43-10040-40)
0.22 μm syringe filter (VWR, catalog number: 514-1263)
50 mL sterile syringe (BD, catalog number: 300865)
Pipetman P1000 tips (Daslab, catalog number: 162222)
Pipetman P200 tips (Daslab, catalog number: 162001)
Pipetman P20 tips (Daslab, catalog number: 162001)
Permanent marker pen (Sarstedt, catalog number: 95.954)
100 mm × 15 mm sterilized Petri dishes (JetBiofil, catalog number: TCD010100)
Ice buckets
500 mL PYREX griffin low-form beaker (Corning, catalog number: 1000J-500)
1 mL sterile syringe (ENFA, catalog number: JS1)
10 mL sterile syringe (BD, catalog number: 307736)
Microlance 3 hypodermic needles 25 G × 5/8'', 16 × 0.5 mm (BD, catalog number: 300600)
Microlance 3 hypodermic needles 30 G × 1/2'', 13 × 0.3 mm (BD, catalog number: 304000)
Polypropylene laboratory tray (Vitlab, catalog number: 72298)
Single-edge razor blade (KBC, catalog number: 1-909-11036)
Double-edge razor blade (Gillette Platinum, catalog number: 21377003)
Peel-A-Way embedding molds truncated - T12 (Polysciences, catalog number: 18986-1)
Peel-A-Way embedding molds rectangular - R30 (Polysciences, catalog number: 18646B-1)
Super Glue-3 (Loctite, catalog number: 2640076)
LDPE Pasteur pipette (Fisher Scientific, catalog number: 747760)
20 μL ZEROTIP pipette micro tip with filter (JetBiofil, catalog number: PMT252020)
2 mL floating tube rack (Sigma-Aldrich, catalog number: R7776-5EA)
Digital timer (Scharlau, catalog number: 038-000566)
Pipetman P10 tips (Daslab, catalog number: 163030)
0.2 mL PCR tubes (Eppendorf, catalog number: 0030124332)
Hand tally counter (VWR, catalog number: 710-0935)
Countess cell counting chamber slides (Invitrogen, catalog number: C10228)
Equipment
Ultrasound system (SonoSite, model: MicroMaxx)
Distilled water (H2O) (Milli-Q, model: IQ-7000)
pH meter (Crison, model: Basic 20+)
Autoclave (Tuttnauer, model: 2540M)
Bunsen burner (Carl Roth, catalog number: CK29.1)
Drying oven (POL-EKO, model: SLW400)
Straight, sharp tips, fine dissecting scissors (Fine Science Tools, catalog number: 14060-09)
Straight, super fine tips dissecting forceps (Fine Science Tools, catalog number: Dumont #5SF 11252-00)
Micro spoon spatulas (RSG Solingen, catalog number: 80.025.150)
Micro flat-ended spatula (RSG Solingen, catalog number: 80.038.150)
15° straight, sharp pointed tips, microsurgical stab knives (MSP, catalog number: 72-1501)
Straight, sharp/blunt tips dissecting scissors (Fine Science Tools, catalog number: 14001-18)
Pipetman P1000 (Gilson, catalog number: F144059M)
Pipetman P200 (Gilson, catalog number: F144058M)
Pipetman P20 (Gilson, catalog number: F144056M)
Laminar flow hood (Telstar, model: CytoStar)
Water bath at 42 °C (MRC, catalog number: WBO-100)
Dissecting scope with incident illumination (Leica, model: M60)
Vibratome (Leica, model: VT1000 S)
Dissecting scope with transmitted illumination (Leica, model: MS5)
Vortex mixer (Labnet)
Water bath at 37 °C (MRC, catalog number: WBO-200)
Microcentrifuge (Labnet, model: Spectrafuge 24D)
Blaubrand Neubauer counting chamber (Brand, catalog number: 718605)
Pipetman P10 (Gilson, catalog number: F144055M)
Inverted microscope (Leica, model: DM IL)
Countess II FL automated cell counter (Invitrogen, catalog number: AMQAF1000)
Procedure
Material preparation before the day of the experiment
Perform an abdominal ultrasound on the jill 27–30 days after mating to verify a successful pregnancy and the number of kits. Plan the number of samples to be isolated during the experiment (see General note 1).
Prepare 1 L of 10× PBS stock solution, autoclave it, and store at room temperature (RT).
Fire-polish glass Pasteur pipettes to round off sharp edges without significantly decreasing the size of the opening. Prepare one glass Pasteur pipette per dissected sample. Autoclave.
Critical: Dry autoclaved fire-polished pipettes for 2 days in the oven to remove any remaining moisture inside them, as microdissected tissue pieces may stick to wet pipettes.
Autoclave dissecting material: two fine dissecting scissors, three super fine dissecting forceps, two micro spoon spatulas, one micro flat-ended spatula, and 1 L of 1× PBS solution.
Reconstitute, aliquot, and store MACS kit reagents to be ready to use according to manufacturer’s instructions.
Material preparation on the day of the experiment
Sterilize microsurgical stab knives, sharp/blunt tip dissecting scissors, 0.5 and 2 mL tubes, 15 and 50 mL centrifuge tubes, 40 μm strainers, 0.22 μm syringe filter, 50 mL syringe, P1000, P200, and P20 pipettes and tips, and a marker. Do this by exposure to UV light inside a laminar flow hood for 30 min.
Aliquot 500 mL of cold (4 °C) L-15 medium in 50 mL centrifuge tubes inside the flow hood and store at 4 °C.
Prepare 50 mL of fresh 4% low melting-temperature agarose in autoclaved 1× PBS. Keep melted agarose in a water bath at 42 °C.
Prepare 50 mL of fresh cell collection medium and filter inside the flow hood using a sterile 0.22 μm syringe filter attached to a sterile 50 mL syringe.
Clean lab benches with RNase decontamination solution. Four benches will be required: one to obtain embryonic day 34 (E34) ferret embryos by cesarian section or to keep postnatal day 1 (P1) ferret kits, one to extract the brains, and two neighboring benches, one to slice the brains and the other to microdissect the brain slices.
Place equipment on clean benches: dissecting scope equipped with incident light source and two empty 100 mm Petri dishes for brain extraction, vibratome with crushed ice inside its cooling bath, and dissecting scope equipped with transmitted illumination for slice microdissection.
Prepare five ice buckets: one big and deep that contains a 500 mL beaker with 350 mL of autoclaved 1× PBS, one big and flat that can hold up to six Petri dishes of 100 mm diameter, one small and flat with space for two Petri dishes of 100 mm diameter, one small for 2 mL tubes, and one big and deep to replace melted ice in the vibratome’s cooling bath.
Brain extraction and cleaning
Time frame to complete this section: ~1 h 20 min
Inject the cocktail of anesthetic/analgesic to overdose the timed-pregnant jill (if obtaining samples from E34 embryos), or the P1 kit (if obtaining samples from early postnatal ferrets).
Administer xylazine (0.05–0.1 mL/kg), ketamine (0.05–0.15 mL/kg), and buprenorphine (0.03–0.06 mL/kg) in a single injection via intramuscular (IM) administration. Wait until there is no sensory response.
Note: For pregnant jills, wait a minimum of 6 min to ensure a complete anesthetic/analgesic effect.
Administer 16.67% (v/v) sodium pentobarbital diluted in saline solution via intraperitoneal (IP) administration. Wait 10 min for full effect.
Note: Both steps C1a and C1b are carried out for pregnant jills and P1 kits, with identical doses per animal’s body weight.
Place the deeply anesthetized pregnant jill on a tray.
Perform a cesarean section using sterilized sharp/blunt-tip dissecting scissors, extract the uterine horns, and place them inside the 500 mL beaker with ice-cold PBS. Cut the diaphragm and the heart right atrium of the jill for final euthanasia.
Extract the embryos by cutting open the uterine horns longitudinally and piercing the amniotic sac using the autoclaved fine dissecting scissors. Keep the embryos in ice-cold PBS.
Fill the 100 mm Petri dishes for brain extraction with ice-cold L-15 aliquoted medium.
Decapitate the E34 embryo or P1 kit by cutting along the base of the skull with the fine or the sharp/blunt-tip dissecting scissors, respectively.
Place the head in the 100 mm Petri dish filled with ice-cold L-15 medium under the dissecting scope with incident light.
Hold the head still for brain extraction with the autoclaved super fine dissecting forceps, by piercing the eyes rostro-caudally (Figure 1A).
Figure 1. Craniotomy for brain extraction. A. Immobilize the head by carefully piercing the eyes with the super fine dissecting forceps. B. Cut the snout open by making incisions on both sides of the head. C. Shell out the olfactory bulbs by carefully inserting a micro-spoon spatula into the rostral side of the opened skull.
Using a new pair of autoclaved fine dissecting scissors, cut along the dorsal midline through the skin and membranous skull, from the back of the head to the snout. For this operation, place the sharp tips of the dissecting scissors upward and the blade slightly parallel to the surface of the skull, to avoid piercing or cutting the brain’s surface.
Note: To keep the surface of the brain intact, the skin layer can be cut first and peeled away to the sides; then, cut both pieces out and proceed with the membranous skull.
At the level of the snout, extend the midline cut to both sides of the head, to open the skull and access the brain (Figure 1B).
Insert a micro spoon spatula rostrally to detach the olfactory bulbs (OBs) from the skull (Figure 1C).
Turn the head over to extract the brain from the skull. Use the micro spoon spatula to carefully release it from any remaining tissue attachments and the fine dissecting scissors to cut the cranial nerves on the ventral side.
Transfer the extracted brain to the other 100 mm Petri dish filled with ice-cold L-15 using the micro spoon spatula.
Remove the meninges (these appear as a transparent film) using two new autoclaved super fine dissecting forceps. Pierce them at the level of the OBs and carefully pull them out rostrocaudally, taking care to avoid touching the brain’s surface. This should be done on both brain sides (dorsal and ventral) sequentially.
Critical: Avoid pulling the meninges with excessive force, or else tissue will deform and damage, particularly at the caudal cortex. Incomplete removal of the meninges may also cause deformation of the tissue during vibratome cutting.
Note: Stabilize the floating brain by pinching the remnants of the spinal cord and cerebellum.
Split the brain along the midline into its hemispheres using a single-edged razor blade. Make a clean cut in one fell swoop. Then, move the blade back and forth without lifting it until the hemispheres separate completely.
Remove the choroid plexus from the lateral ventricle and any meninges debris from the caudal cortex using the super fine dissecting forceps.
Critical: Avoid pulling the choroid plexus with excessive force to prevent tissue deformation. Incomplete removal of the choroid plexus may also cause deformation of the tissue during vibratome cutting.
Clamp the remnants of the spinal cord and cerebellum with one of the dissecting forceps. Close the tips from the second forceps and move them down along the edge of the clamping forceps to cut out the remaining.
Depending on the single-cell assay to carry out, and hence the required cell concentration (see General note 1), repeat steps C5–17 to obtain other E34 ferret embryos or steps C1 and C5–17 for other P1 kits.
Brain embedding and slicing
Time frame to complete this section: ~1 h 50 min
Label a histology embedding mold (whose size is in accordance with the brain size) with the animal number and L (left) or R (right) according to the brain hemisphere.
Fill the mold with freshly prepared 4% low-melting temperature agarose.
Transfer one brain hemisphere into the mold using the micro-spoon spatula, with the minimum accompanying liquid.
Rotate the brain hemisphere inside the liquid agarose with a P200 pipette tip by describing circles around the tissue in all directions, but without touching it.
Critical: Multiple turns of the tissue in all directions within the agarose are essential to build a complete coating around it and to avoid its popping out during vibratome cutting later on.
Before the agarose solidifies, place the brain hemisphere horizontally at the bottom of the mold with its lateral side facing down and its medial side facing up.
Place the mold on ice until it solidifies completely.
Extract the solidified agarose block containing the brain hemisphere from the mold and place it on the lid of a 100 mm Petri dish. The medial side of the hemisphere should face the base of the pyramid. Using the single-edged razor blade, cut the sides of the agarose block into the shape of a truncated square pyramid.
Repeat steps D1–7 for the other hemispheres.
Note: Up to six hemispheres from E34 ferret embryos or up to four hemispheres from P1 kits can be vibratome-cut in one go.
Glue the truncated pyramids on the vibratome’s specimen disk. To harden the glue quickly and improve the truncated pyramid attachment, add drops of ice-cold L-15 to the base of the pyramids with an LDPE Pasteur pipette. Given that the medial side of the hemisphere is facing the pyramid’s base, the lateral side of the cerebral cortex should face upward and will be the first to be cut.
Critical: The corpus callossum on the ventral side of the brain has a harder consistency than the cortical cell bodies. Therefore, the softer tissue should be cut before to avoid being pushed by the callossum.
Tighten the specimen disk to the buffer tray, fill it with ice-cold L-15 medium, place the knife holder with an unused blade from the double-edged razor blades, set the sectioning parameters on the vibratome, and place new ice inside the cooling bath.
Label 100 mm Petri dishes (as many as brain hemispheres to be cut), fill them with ice-cold L-15 medium, and place them on the big flat ice bucket.
Cut the brain hemispheres from E34 embryos or P1 kits.
Make 300 μm thick slices at a vibratome speed of 2, frequency 5 to 6.
Use two micro spatulas (one spoon and one flat-ended) to collect each slice and place them in their corresponding Petri dishes.
Check the slices under the dissecting scope equipped with transmitted light until the regions of interest (ROIs) [the prospective splenial gyrus (SG) and lateral sulcus (LS), in our case] come into sight.
Microdissection
Time frame to complete this section: ~2 h 15 min for E34 embryos or ~1 h 40 min for P1 kit.
Set a 100 mm Petri dish with the identified ROIs in the small flat ice bucket.
Replace the Petri dish set aside with a new dish, so brain slices from that hemisphere can still be collected during the microdissection.
Place the Petri dish with the identified ROIs under the dissecting scope with transmitted illumination. Using two microsurgical stab knives, carefully drag the slice of interest by its surrounding agarose and place it at the center of the field of view.
Microdissect the first ROI: prospective SG (Figure 2A).
Hold the slice by the surrounding agarose with one knife and use the other knife to make a horizontal cut at the most caudal end of the cortex, right next to the hippocampal formation.
Without lifting the knife that holds the agarose, use the second knife to make a second cut at a 45° angle with respect to the previous cut.
Critical: These two cuts will detach the prospective SG from the agarose. This piece of tissue is very small, so do not miss sight of it.
Critical: A lack of precision when making the horizontal cut next to the hippocampal formation, or at a 45º angle, may lead to the isolation of unwanted cells from neighboring cortical areas, which may be difficult to discard during subsequent analyses.
Figure 2. Neocortical areas and layers to microdissect. A. Example of live parasagittal slices from the brains of embryonic day (E) 34 and postnatal day (P) 1 ferrets with lateral sulcus (LS) and splenial gyrus (SG) outlined. Ctx, cortex; Str, striatum; Hpc, hippocampus; Th, thalamus. B, C. Tissue microdissections of the SG (B) and LS (C) from the two developmental stages with neocortical layers depicted. CP/MZ, cortical plate/marginal zone; IZ, intermediate zone; SVZ, subventricular zone; OSVZ, outer subventricular zone; ISVZ, inner subventricular zone; VZ, ventricular zone. Figure modified from [8].
Set aside the slice missing the prospective SG and focus on the dissected SG at the highest magnification of the dissecting scope.
Microdissect the most apical cortical layer from the first ROI, the ventricular zone (VZ) (Figure 2B).
Use one knife to hold the dissected slice piece by an unwanted part of the tissue, in our case the top half.
Carefully make a clean cut in one fell swoop of the layer that is most cell-dense and opaque, extending from the side of the cortex limiting with the ventricular cavity (see General note 4). This covers approximately 3/10 of the cortex total thickness in E34 embryos and 1/10 in P1 kits.
Critical: The VZ is slightly concave, so make your cut such that it contains the whole thickness of the VZ. Excess tissue can be trimmed off afterward.
Critical: Cut with moderate force to avoid pushing away the dissected thin layer and possibly losing it in the medium.
Turn the VZ around, prick the layer, and trim away any unwanted tissue.
Cut the dissected VZ into small pieces.
Note: Increasing the surface area of the dissected tissue will enable a larger contact between the tissue and the dissociating enzyme later on.
Critical: Do not mix the VZ pieces with the previously trimmed chunks.
Use a P20 pipette with low retention, RNase-free filter-tip to aspirate the VZ pieces with as little medium as possible. Transfer the pieces to a 2 mL tube, label it, and put it in the small ice bucket.
Critical: Check the pipette tip under the dissecting scope to make sure that the pieces have not stuck to its walls or opening. If that is the case, see the Troubleshooting section, problem 1.
Microdissect the second cortical layer from the first ROI, the outer subventricular zone (OSVZ) (Figure 2B).
Note: The OSVZ emerges at E34, therefore this layer is not yet present in E34 ferret embryos but is developed in P1 kits (Figure 3).
Again, use one knife to hold the dissected slice piece by the top half of the tissue.
Trim away the remnants of the cell-dense layer at the apical side.
Note: This layer is the inner subventricular zone (ISVZ). At P1, it has the thickness of the VZ; however, part of it has probably been dissected with the VZ and was later trimmed off.
Carefully make a clean cut in one fell swoop of the bottom half still left of the cortex's total thickness. It is a translucent layer, although denser than the layer directly on top [the intermediate zone (IZ)], especially its upper third. The lower 2/3 of this layer displays cells radially organized in ribbons (see General note 4).
Cut the dissected OSVZ into small pieces.
Note: Increasing the surface area of the dissected tissue will enable a larger contact between the tissue and the dissociating enzyme later on.
Critical: Do not mix the OSVZ pieces with the previously trimmed ISVZ.
Figure 3. Layer and cellular composition along cortical development. Layer composition at embryonic day (E) 34, when apical radial glia cells (aRGC) in the ventricular zone (VZ) self-consume to generate basal progenitors. At this stage, the first basal radial glia cells (bRGC) that initiate the outer subventricular zone (OSVZ) are produced. At postnatal day (P) 1, the bRGC from the OSVZ self-amplifies and, contrary to the inner subventricular zone (ISVZ), arises independently from aRGC. IPC, intermediate progenitor cell; tRGC, truncated radial glia cell; MZ, marginal zone; CP, cortical plate; IZ, intermediate zone; SVZ, subventricular zone. Figure modified from [16].
Use the P20 pipette with low retention, RNase-free filter-tip to aspirate the OSVZ pieces with as little medium as possible. Transfer the pieces to a 2 mL tube, label it, and put it in the small ice bucket.
Critical: Check the pipette tip under the dissecting scope to be sure that the pieces have not stuck to its walls or opening. If that is the case, see the Troubleshooting section, problem 1.
Select the lowest magnification in the dissecting scope, focus the slice from which the prospective SG was microdissected, and trim away the following piece of cerebral cortex with a similar size as the microdissected SG.
Microdissect the second ROI, prospective LS (Figure 2A).
Hold the slice by the surrounding agarose with one knife and use the other one to make a vertical cut over the cortex overlying the hippocampal region immediately caudal to the dentate gyrus.
Critical: This cut will detach the prospective LS from the agarose, so do not mistake it with the previous discarded piece of tissue.
Focus on the dissected LS at the highest magnification of the dissecting scope.
Repeat steps E6–7 to isolate the most apical cortical layer from the second ROI, VZ from LS (Figure 2C).
Repeat steps E8–9 to isolate the second cortical layer from the second ROI, OSVZ from LS (Figure 2C).
Microdissect the layers from all the vibratome slices containing the ROIs: 2–3 slices per hemisphere from E34 embryos or 4–5 slices per hemisphere from P1 kits.
Note: Pool the VZ pieces from 2–3 E34 embryos in the same 2 mL tube to obtain the optimal cell concentration range; see General note 1.
Single-cell suspension
Time frame to complete this section: ~1 h 20 min
For each sample to be processed, prepare 1,950 μL of enzyme mix 1 from the MACS kit inside the laminar flow hood by adding the following components:
Buffer X, 1,900 μL
Enzyme P, 50 μL
Vortex and prewarm the mixture in a water bath at 37 °C for 20 min before use.
For each sample to be processed, pre-wet a 40 μm strainer sitting in a 2 mL tube with 1× HBSS.
Note: Wetting the strainer membrane helps to reduce cell adhesion.
Carry the samples on ice to the flow hood.
Add 1,950 μL of prewarmed enzyme mix 1 to each sample inside the hood. Incubate them in a water bath at 37 °C for 20–30 min, gently finger flickering the tubes from time to time (do not vortex).
Note: Incubation time depends on tissue size and enzyme activity. While flickering the tubes, look at the tissue's appearance and stop the dissociation reaction when the pieces are broken into much smaller portions. Recommended incubating times: E34 pooled VZ pieces for 27 min and P1 tissue pieces from VZ or OSVZ for 24 min.
For each sample to be processed, prepare 30 μL of enzyme mix 2 from the MACS kit inside the flow hood by adding the following components:
Buffer Y, 20 μL
Enzyme A, 10 μL
Add 30 μL of enzyme mix 2 to each sample inside the hood. Gently invert the tubes to mix the content and stop the dissociation reaction (do not vortex).
Critical: Be careful not to spill the liquid when introducing the pipette tip inside the 2 mL tube, as it is nearly full.
For each sample to be processed, pre-coat a 2 mL tube with cell collection medium and label it.
Note: Coating the tube’s interior surface helps to reduce cell adhesion.
Replace the 2 mL tubes where pre-wet strainers are sitting by the pre-coated 2 mL tubes. Discard the 2 mL tubes containing filtered HBSS.
Using the autoclaved, fire-polished glass Pasteur pipettes, pipette E34 and P1 samples slowly up and down 15 and 10 times, respectively, to mechanically dissociate the tissue. Avoid forming air bubbles.
Critical: Check that the tissue pieces are not left stuck inside the pipette.
Using a P1000 pipette, suspend the cells from the E34 and P1 samples by slowly pipetting up and down 5 and 15 times, respectively. Avoid forming air bubbles.
Filter cell suspensions to the pre-coated 2 mL tubes through the 40 μm pre-wet strainer.
Critical: Absorb with the pipette any liquid held within the back of the strainer to avoid losing any cells.
Discard the 40 μm strainers and centrifuge cell suspensions at 92× g for 5 min at RT.
Carefully aspirate the supernatant and resuspend the cell pellet in 600 μL of cell collection medium.
Critical: Save the supernatant aside on ice in case you accidentally aspirated the cells together with the supernatant; see Troubleshooting section, problem 2.
Centrifuge cell suspensions at 92× g for 5 min at RT.
Carefully aspirate the supernatant and resuspend the cell pellet in 50 μL of cell collection medium.
Critical: Save the supernatant aside on ice in case you accidentally aspirated the cells together with the supernatant; see Troubleshooting section, problem 2.
Note: The required volume of cell collection medium might vary according to the single-cell assay further applied.
Cell concentration and viability
Time frame to complete this section: ~30 min
Place the samples on ice. For each sample to be processed, measure cell concentration and viability from a homogenous cell suspension using a hemocytometer.
Critical: Pipette the samples very slowly up and down to distribute the cells evenly, otherwise their concentration and viability will be under or overestimated.
Clean the hemocytometer with 70% ethanol and dry it well.
Add 10 μL of cell suspension to 10 μL of 0.4% trypan blue solution in a 0.2 mL tube. Mix it gently.
Load 10 μL of trypan blue–stained cell suspension into the hemocytometer chamber.
Using an inverted microscope and a hand tally counter, focus on the grid and count live, unstained cells in the four sets of 16 squares from the chamber.
Following the same guidelines, count dead, stained cells.
Calculate cell concentration (average cells per set × 104 × 2 dilution) and percentage viability [live cell concentration/(live cell concentration + dead cell concentration)] from the samples; see General note 2.
Optionally, re-measure cell concentration and viability from each sample using an automatic cell counter.
Critical: Pipette the samples very slowly up and down to distribute the cells evenly; otherwise, their concentration and viability will be under or overestimated.
Load 10 μL of trypan blue–stained cell suspension into a cell counter chamber slide.
Insert the slide into the automatic counter.
Copy cell concentration and percentage viability results.
Average manual and automatic measurements.
Proceed immediately to the first step from the chosen single-cell assay protocol.
Critical: Only apply debris-free single-cell suspensions with cell viability over 90%. If the suspension contains clustered cells and/or debris, see Troubleshooting section, problem 3.
Data analysis
Data processing and analyses that followed this protocol can be found in the Materials and Methods section from [8]. Briefly, single cells from our single-cell suspensions were isolated by microfluidics, and their cDNA libraries were prepared according to Chromium Single Cell 3' (10× Genomics) single-cell assay. After sequencing, single-cell libraries were aligned to the reference ferret genome (MusPutFur1.0/GCF_000215625.1), cell barcodes were filtered, and the quality of cells was assessed according to the standard thresholds in the field. Then, samples containing high-quality cells were normalized, their cell-cycle differences and percentage of mitochondrial genes were regressed out, and finally they were integrated. Cell clusters from the integrated samples were identified, their resolution and quality were assessed, and cortical cell types were labeled according to their gene expression. Finally, multiple analyses were carried out, such as analysis of differential gene expression, cluster and functional pathways enrichment, trajectory analyses, or study of genes related to human malformations of cortical development. This data was also integrated into datasets similarly obtained from other species.
Validation of protocol
This protocol has been used and validated in the following research article:
Del-Valle-Anton et al. [8]. Multiple parallel cell lineages in the developing mammalian cerebral cortex. Science Advances (Figure 1, panel B; Figure S1).
This protocol was successfully applied in a total of 18 samples: E34 VZ SG n = 3, E34 VZ LS n = 3, P1 VZ SG n = 3, P1 VZ LS n = 3, P1 OSVZ SG n = 3, P1 OSVZ LS n = 3. Each E34 sample comprised microdissected tissue of two to four ferret embryos from the same litter (n = 4 litters), and P1 samples were composed of one animal per litter (n = 5 litters). RNA integrity number (RIN) average score of 7.6 was obtained from single-cell suspensions. Bioanalyzer cDNA traces for library construction by means of Chromium Single Cell 3' (10× Genomics) experiments reflected a successful cDNA amplification (average of 5,873.84 pg/μL for undiluted samples) (Figure 4). Other high-quality parameters for single-cell libraries were met, such as steep barcode-rank distributions (which distinguish cell-containing droplets from ambient RNA), strong positive correlations between detected genes and unique molecular identifiers (UMIs) (Pearson correlation value = 0.95), high libraries complexity (> 0.8 log10 genes identified per UMI), and low percentage of mitochondrial genes per cell (average of 6%).
Figure 4. Representative readout from Agilent 2100 Bioanalyzer. A, B. Electrophoresis gel image (A) and electropherogram traces (B) of two amplified cDNA samples. bp, base pair; FU, fluorescence unit. C. Samples quality-control results after cDNA amplification.
The neocortical layer microdissection from the developing ferret brain shown in this protocol has been partially used and validated in the following research articles:
Singh et al. [17]. Gene regulatory landscape of cerebral cortex folding. Science Advances (Figure 1, panel A; Figure 2, panel A).
Martínez-Martínez et al. [18]. A restricted period for formation of outer subventricular zone defined by Cdh1 and Trnp1 levels. Nature Communications (Figure 5).
de Juan Romero et al. [19]. Discrete domains of gene expression in germinal layers distinguish the development of gyrencephaly. The EMBO Journal (Figure 1, panels A–E).
Landmarks for prospective SG and LS identification in embryonic and early postnatal ferret brains used in this protocol have been previously validated in the following research articles:
Borrell et al. [20]. In vivo gene delivery to the postnatal ferret cerebral cortex by DNA electroporation. Journal of Neuroscience (Figure 2; Figure 3, panels A, B; Figure 5, panels A, D; Figure 6).]
Reillo et al. [3]. A role for intermediate radial glia in the tangential expansion of the mammalian cerebral cortex. Cerebral Cortex (Figures 3–5).]
Cell lineage tracing, dye tracing, and tracking of radial fibers scaffold in the developing ferret caudal cortex were the techniques used for the faithful identification of these tissue landmarks.
General notes and troubleshooting
General notes
For obtaining the optimal cell concentration range for one reaction of Chromium Single Cell 3' (10× Genomics) experiment, 2–3 E34 ferret embryos or one P1 ferret kit are required. The recommended starting cell concentration might vary between single-cell assays.
By means of this protocol, the obtained cell concentration range for E34 samples is 7.3 × 105–5.4 × 106 cells/mL, for P1 samples from VZ is 6.4 × 105–2.9 × 106 cells/mL, and from OSVZ is 7.2 × 105–2.6 × 106 cells/mL.
After animal anesthesia, follow the protocol non-stop. A shorter experimental time, combined with tissue maintenance on ice, produces better outputs. The average experimental time for two researchers working together is 5 h.
Turn the light intensity from the dissecting scope up and down to facilitate distinguishing between the different neocortical layers.
During tissue dissociation, treat cells very gently to avoid cell death: finger flicker and invert the tubes gently, do not vortex the cells, make sure that the openings of the glass Pasteur pipettes are not too small, pipette slowly, and avoid forming air bubbles.
This protocol has been implemented to obtain a single-cell suspension from E34 or P1 ferret neocortical layers. If applied in more advanced ferret developmental stages, longer incubation time with the dissociating enzyme and larger mechanical dissociation should probably be used to counteract the stiffness caused by myelin. If, on the other hand, it is applied at earlier stages of development, shorter incubation time and milder mechanical dissociation might probably be recommended to avoid cell death.
Troubleshooting
Problem 1: Dissected tissue pieces have stuck to the walls or opening of the pipette tip while transferring them to the 2 mL tube.
Possible cause: The tissue has become sticky after being cut for a while.
Solution: Introduce the tip inside a Petri dish with ice-cold L-15 medium, focus on the stuck tissue with the dissecting scope, and pipette up and down with force to detach them. Do not miss sight of them to avoid losing the pieces due to their small size.
Problem 2: Single-cell suspension contains very few cells.
Possible cause: Accidental suction of the cells together with the supernatant after centrifuging.
Solution: Centrifuge the supernatants resulting from the first and second centrifugation steps at 92× g for 2 min, carefully aspirate the supernatant from both of them, resuspend the cell pellets in 40 μL of cell collection medium, and proceed to check cell concentration and viability.
Problem 3: Single-cell suspension contains clustered cells and/or debris.
Possible cause: Large size of microdissected tissue and/or low enzymatic dissociation.
Solution: Filter cell suspension again through a 40 μm pre-wet strainer to another pre-coated 2 mL tube. This second filtration can only be done once, otherwise cell membranes are compromised. In future experiments, cut the microdissected layer into smaller pieces and/or increase tube flickering during the dissociation reaction.
Acknowledgments
This work was supported by Spanish Research Agency (AEI) grants PGC2018-102172-B-I00, PID2021-125618NB-I00 and European Research Council grant UNFOLD (101118729) to V.B., who also acknowledges financial support from the AEI through the “Severo Ochoa” Programme for Centers of Excellence in R&D (CEX2021-001165-S). L.D.-V.-A. was recipient of an FPI contract from the Spanish Research Agency (AEI). This protocol was first implemented, described and validated in [8].
Competing interests
The authors declare that they have no competing interests.
Ethical considerations
All animals were treated according to Spanish and European regulations. The protocol was conducted in agreement with the Institutional Animal Care and Use Committee (IACUC) from the Spanish Council for Scientific Research (CSIC). The experimental protocol was performed at Instituto de Neurociencias on healthy ferret embryos or postnatal kits (Mustela putorius furo). Pregnant sable ferrets were housed at the Animal Research Facility (SEA) of the Miguel Hernández University (UMH), where specialized and accredited personnel were responsible for their wellbeing. They were kept in a 12/12 h light/dark cycle with standard chow and water freely accessible.
References
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Article Information
Publication history
Received: Jul 15, 2024
Accepted: Oct 5, 2024
Available online: Oct 29, 2024
Published: Dec 20, 2024
Copyright
© 2024 The Author(s); This is an open access article under the CC BY-NC license (https://creativecommons.org/licenses/by-nc/4.0/).
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Neuroscience > Development > Morphogenesis
Neuroscience > Neuroanatomy and circuitry > Cortex
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