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4,729 | https://bio-protocol.org/en/bpdetail?id=4729&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Development of a Mouse Model of Hematopoietic Loss of Y Chromosome
SS Soichi Sano
KW Kenneth Walsh
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4729 Views: 619
Reviewed by: Gal HaimovichChris TibbittRAVI THAKUR
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Original Research Article:
The authors used this protocol in Science Jul 2022
Abstract
This protocol describes the generation of chimeric mice in which the Y chromosome is deleted from a proportion of blood cells. This model recapitulates the phenomenon of hematopoietic mosaic loss of Y chromosome (mLOY), which is frequently observed in the blood of aged men. To construct mice with hematopoietic Y chromosome loss, lineage-negative cells are isolated from the bone marrow of ROSA26-Cas9 knock-in mice. These cells are transduced with a lentivirus vector encoding a guide RNA (gRNA) that targets multiple repeats of the Y chromosome centromere, effectively removing the Y chromosome. These cells are then transplanted into lethally irradiated wildtype C57BL6 mice. Control gRNAs are designed to target either no specific region or the fourth intron of Actin gene. Transduced cells are tracked by measuring the fraction of blood cells expressing the virally encoded reporter gene tRFP. This model represents a clinically relevant model of hematopoietic mosaic loss of Y chromosome, which can be used to study the impact of mLOY on various age-related diseases.
Graphical overview
Keywords: Hematopoietic mosaic loss of Y chromosome (mLOY) Cas9-knockin mice gRNA-encoding lentivirus Bone marrow reconstitution Heart failure Clonal hematopoiesis
Background
Mosaic loss of Y chromosome (mLOY) is the most prevalent post-zygotic mutation in humans (Forsberg et al., 2014). More than 40% of men exhibit appreciable mLOY in hematopoietic cells by the age of 70 years (Thompson et al., 2019). The Y chromosome contains a relatively small number of protein-coding genes, and it can be viewed as a gene desert. Thus, while the phenomenon of mLOY was identified more than half a century ago, it has been regarded as a benign age-related alternation or as a biomarker for aging. However, recent research has found that mLOY in blood cells elevates the risk of all-cause mortality and a variety of human disorders, such as solid cancers, Alzheimer’s disease, and cardiovascular disease (Forsberg et al., 2014; Dumanski et al., 2016; Thompson et al., 2019; Sano et al., 2022). However, due to the descriptive nature of this epidemiological research, it cannot be determined whether mLOY in blood plays a causal role in the development of disease. To address this gap in knowledge, experimental studies can provide insights as to whether mLOY is causally linked to age-related illness.
The reconstitution of the hematopoietic system by the bone marrow transplantation (BMT) of gene-modified cells has been widely employed in mice to study the regulatory mechanisms that control the blood system. Recently, we employed CRISPR/Cas9-mediated gene editing to create mice with targeted mutations in genes to model the phenomenon of clonal hematopoiesis of indeterminate potential. In this method, lineage-negative cells isolated from the bone marrow of Cas9-transgenic mice are transduced with lentivirus vectors that encode gRNAs ex vivo. Hematopoietic lineage-negative cells are comprised of stem/progenitor cells that lack the expression of specific cell surface markers associated with various blood cell lineages. The lineage-negative cell pool has the ability to differentiate into multiple blood cell types. The resulting gene-edited cells are then transplanted into conditioned wildtype mice, achieving an editing efficiency of 90% within the transduced hematopoietic stem and progenitor cells (HSPC). More recently, we employed a similar CRISPR/Cas9 approach to ablate the entire Y chromosome within hematopoietic cells (Sano et al., 2022). In this application, gRNAs were designed to specifically target the repetitive sequences within the Y chromosome, allowing the creation of mice that lacked the Y chromosome in a portion of leukocytes.
Materials and reagents
BD Luer-LokTM disposable syringes, single use, 10 mL (BD, catalog number: 309604)
BD Luer-LokTM disposable syringes, single use, 50 mL (BD, catalog number: 309653)
BD PrecisionGlideTM needle, 18 G (BD, catalog number: 305195)
Insulin syringe, 29 G (EXLINT, catalog number: 26028)
CorningTM FalconTM 50 mL conical centrifuge tubes (Fisher Scientific, catalog number: 14-959-49A)
CorningTM FalconTM 15 mL conical centrifuge tubes (Fisher Scientific, catalog number: 14-959-53A)
Falcon® 100 mm TC-treated cell culture dish (Life Sciences, catalog number: 353003)
Falcon® 6-well clear flat-bottom TC-treated multi-well cell culture plate, with lid (Life Sciences, catalog number: 353046)
Corning® Costar® ultra-low attachment multiple well plate, 6 well (Millipore Sigma, catalog number: CLS3471)
Polypropylene centrifuge tubes (Beckman, catalog number: 326823)
FisherbrandTM sterile cell strainers, 70 μm (Fisher Scientific, catalog number: 22-363-548)
Mouse Millex-HV syringe filter unit
B6 mice (C57BL/6J, The Jackson Laboratory, stock number: 000664, bred in our laboratory or purchased)
Rosa26Cas9 knock-in mouse [B6(C)-Gt(ROSA)26Sorem1.1(CAG-cas9*,-EGFP)Rsky/J, The Jackson Laboratory, stock number: 028555, bred in our laboratory]
pLKO5.sgRNA.EFS.tRFP (Addgene, catalog number: 57823)
psPAX2 (Addgene, catalog number: 12260)
pMD2.G (Addgene, catalog number: 12259)
HEK 293T cell line (ATCC, catalog number: CRL-3216)
Collagen from calf skin (Millipore Sigma, catalog number: C9791)
PEI MAX (Polysciences, catalog number: 24765-1)
Phosphate-buffered saline (PBS), pH 7.4 (Thermo Fisher Scientific, catalog number: 10010023)
Dulbecco’s modified Eagle’s medium (DMEM), high glucose (Millipore Sigma, catalog number: D5796)
RPMI-1640 medium (Millipore Sigma, catalog number: R8758)
StemSpanTM SFEM (STEMCELL Technologies, catalog number: 09600)
Recombinant murine thrombopoietin (TPO) (Peprotech, catalog number: 250-03)
Recombinant murine stem cell factor (SCF) (Peprotech, catalog number: 315-14)
Polybrene infection/transfection reagent (Merck Sigma, catalog number: TR-1003-G)
Penicillin-streptomycin (10,000 U/mL) (Thermo Fisher Scientific, catalog number: 15140122)
Isoflurane liquid inhalation 99.9% (Henry Schein, catalog number: 1182098)
Lineage Cell Depletion kit (Miltenyi Biotec, catalog number: 130-090-858)
Lenti-X qRT-PCR Titration kit (Clontech, catalog number: 631235)
Circular Glass coverslips (12 mm) (Fisher, catalog number: 12-545-80P)
Rubber Cement (Amazon)
Microscope slides (Fisher, catalog number: 12-544-2)
Coplin Jars (glass) (Fisher, catalog number: 08816)
Coplin Jars (plastic) (Market Lab, catalog number: ML9801)
Coverslips, 22 × 50 mm (Fisher, catalog number: 12-518-105EP)
Diamond Tip Scribe (Fisher, catalog number: 08-675)
DNA probes (direct labeled) and validated (Vysis or Cytocell, XqF4/YqA1)
20× SSC (solution) (Fisher, catalog number: BP1325-1)
Purified water, D.I.U.F. (Fisher, catalog number: W2-4)
2× SSC (pH = 7.2) (Cell Line Genetics, in house)
70% ethanol solution (Cell Line Genetics, in house)
85% ethanol solution (Cell Line Genetics, in house)
100% ethanol, 200 Proof (Fisher, catalog number: BP2818-4)
1:20 VectaShield with DAPI (Cell Line Genetics, in house)
1% Formaldehyde (Cell Line Genetics, in house)
0.01 N HCl (Cell Line Genetics, in house)
PBD (phosphate buffered detergent) (Cell Line Genetics, in house)
Pepsin stock solution (Cell Line Genetics, in house)
Recombinant murine SCF (Peprotech, catalog number: 250-03)
Recombinant human IL-11 (Peprotech, catalog number: 200-11)
Recombinant human Flt3-Ligand (Peprotech, catalog number: 300-19)
Recombinant murine IL-3 (Peprotech, catalog number: 213-13)
IMDM (Gibco, catalog number: 12440053)
Doxycycline hyclate (Sigma-Aldrich, catalog number: D9891)
Colcemid Solution (Thermo Fisher, 15212012)
Cell culture media (see Recipes)
Wash buffer for cells (see Recipes)
MACS buffer (see Recipes)
Equipment
Forceps (Fine Science Tools)
Scissors (Fine Science Tools)
Eppendorf® centrifuge 5424R (Millipore Sigma, catalog number: 05-401-205)
Ultracentrifuge (Beckman, model: L8-70M)
Freezer, 4 °C
Deep freezer, -80 °C
Applied BiosystemsTM QuantStudioTM 6 Flex Real-Time PCR System (Thermo Fisher Scientific, catalog number: 4485699)
MACS® MultiStand (Miltenyi Biotec, catalog number: 130-042-303)
LS Columns (Miltenyi Biotec, catalog number: 130-042-401)
QuadroMACSTM Separator (Miltenyi Biotec, catalog number: 30-090-976)
RS-2000 X-ray irradiator (Rad Source Technologies, Inc.)
BD LSRFortessa flow cytometer (BD Biosciences)
Fluorescence microscope (Olympus, catalog number: BX-41)
Appropriate filters matched to excitation and emission spectra of fluorophores used (Chroma, model: Vysis Probes: Spectrum Orange/Spectrum Green, Cytocell Probes: Rhodamine/FITC)
TruTemp DNA micro heating system (Abbott Molecular, model: ThermoBrite)
Stir plate (Fisher, model: Isotemp)
pH meter (Fisher, catalog number: AE150)
Balance (OHaus, catalog number: EP613c)
Water bath (Fisher, model: Isotemp 205)
Quick Spin centrifuge (Fisher, model: Quick Spin)
Software
FlowJoTM software (BD Biosciences)
GraphPad Prism9 (https://www.graphpad.com/scientific-software/prism/)
BioRender (https://biorender.com/)
Procedure
Generation of lentivirus
Lentiviruses are produced by co-transfecting psPAX2, pMD2.G, and the lentivirus pLKO5.0.sg.EFS.tRPF into HEK 293T cells. Our standard procedure is to purify lentivirus particles from the cell culture supernatant of twelve 6-well plates (i.e., 72 wells in total). The volume of supernatant is approximately 216 mL (3 mL/well × 72 wells). The culture supernatant is collected 48 h after transfection and concentrated by ultracentrifugation. The lentiviral titer is measured using a commercially available qPCR-based assay. Optimized methods for high-titer lentivirus preparation and storage are discussed elsewhere (Cante-Barrett et al., 2016; Sano et al., 2019).
Seeding of HEK 293T cells
In advance, we usually prepare ten 10 cm dishes with 90% confluent HEK 293T cells.
Prepare an approximately 1:200 solution of commercially available 0.1% collagen from calf skin in PBS (i.e., final concentration of 0.0005%). We usually prepare 40 mL of 0.0005% collagen solution for one batch of lentivirus generation by adding 200 μL of 0.1% collagen solution to 40 mL of 1× PBS.
Add 500 μL of 0.0005% collagen solution to each well of 6-well plates and incubate at 37 °C in 5% CO2 for more than 30 min (coating wells with collagen). The collagen solution used for coating wells can be reused several times. The collected solution should be kept at 4 °C.
Seed HEK 293T cells at a density of 1 × 106 cells in 2 mL of complete DMEM per well (i.e., suspend the cells at a concentration of 0.5 × 106 cells/mL and use 2 mL of cell suspension per well) and incubate at 37 °C in 5% CO2 for 2 h until the cells attach firmly. More than 7.2 ×107 cells are required for twelve 6-well plates.
Transfection into HEK 293T cells
To prepare the mixture of three transfection plasmids, combine plasmids (0.9 μg of lentivirus vector, 0.6 μg of psPAX2, and 0.3 μg of pMD2.G for each well) and then achieve a total volume of 10 μL by adding deionized water. Adjust volumes accordingly depending on the number of wells (72 wells). The amount and ratio of each plasmid may need to be further optimized to suit an individual researcher’s needs.
Carefully add 50 μL of 1× PBS and 5 μL of the diluted PEI MAX (1.0 mg/mL) to the plasmid mixture for each well and incubate for 15 min at room temperature. Next, add 1 mL of complete DMEM to the mixture for each well.
Note: PEI MAX can be stored at 4 °C after reconstitution.
Aspirate the media from the 6-well plates, add 1 mL of the plasmid mixture prepared above, and incubate at 37 °C in 5% CO2 for 3 h.
Replace the media with 2 mL of fresh, complete DMEM and incubate at 37 °C in 5% CO2 for 24 h.
The following day, add 1 mL of fresh complete DMEM and incubate at 37 °C in 5% CO2 for an additional 24 h (total incubation time is 48 h).
Lentivirus purification
Transfer the culture supernatant to 50 mL Falcon tubes and centrifuge at 3,000 × g for 15 min to remove any free-floating cells.
Using a syringe, filter the supernatant through a 0.45 μm filter into new 50 mL Falcon tubes and transfer the filtrate to new polypropylene ultracentrifuge tubes.
Ultracentrifuge at 72,100× g at rmax for 3 h at 4 °C.
Carefully aspirate the supernatant after ultracentrifugation, leaving behind the white pellet. It is ideal to aspirate as much supernatant as possible to minimize residual liquid. However, be careful to not dry the pellets.
Add 200 μL of serum-free hematopoietic cell expansion medium to the pellet and incubate on ice for 1 h. Following this, resuspend pellets without aeration. Avoid bubble formation as much as possible. The volume of final solution is typically greater than 200 μL.
Acquire a 10 μL aliquot for measuring the viral titer and store the aliquot and all remaining viral suspension at -80 °C until needed. Snap freezing is not required.
Thaw a 10 μL aliquot (prepared in step A3f) and titrate the virus with a qPCR-based assay according to the manufacturer’s instructions.
Note: The procedures outlined above should be carried out in a biosafety class II cabinet. Unless otherwise specified, all procedures can be carried out at room temperature.
Isolation of lineage-negative cells from Cas9-knockin mice
Harvest of bone marrow cells (Sano et al., 2019)
Euthanize 8–10 male CRISPR/Cas9 knock-in mice (8–10 weeks of age) by 5% isoflurane followed by cervical dislocation; then, disinfect their skin with 70% ethanol.
Using dissecting scissors, make a small incision in the skin of the abdomen and peel the skin distally in both directions to expose the legs and arms.
Carefully separate the lower limbs from the hip bone by dislocating the hip joint. Cut along the femur head to remove the femur completely from the hip. Dislocate the knee and cut at the joint to separate the femur and tibia, while keeping the bone epiphysis intact. Dislocate the ankle joint and peel away the foot and extra muscle.
Using dissecting scissors, cut over the shoulder to detach the upper limbs. Dislocate the shoulder, then cut at the elbow joint to harvest the humerus bone.
Manually dislocate hip joints on both sides and harvest hip bones.
Use cellulose-fiber wipes to carefully remove muscles from the femurs, tibias, humeri, and hip bones. Take extra precaution to ensure that the bones do not break during this process.
Place the isolated bones into a 50 mL conical tube containing RPMI-1640 medium and place on ice.
Grasp the bone with blunt forceps and, using dissecting scissors, carefully cut both epiphyses (1–2 mm). Both ends of hip bones will be opened by this procedure.
Note: An insufficient cutting of bone will lead to an incomplete harvest of bone marrow, while overly aggressive cutting will result in cell loss.
Preparation of collecting tubes: make a small hole in the bottom of sterile 0.5 mL microcentrifuge tubes using an 18 G needle and place them in sterile 1.5 mL tubes containing 200 μL of ice-cold sterile PBS.
Place the cut bones inside the 0.5 mL microcentrifuge tubes and close the lids.
Place tubes in a centrifuge and spin at 10,000× g for 30 s at 4 °C (Figure 1).
Figure 1. Isolation of bone marrow cells
After centrifugation, all bone marrow cells should have passed through the hole in the 0.5 mL microcentrifuge tubes, and the cells should have been translocated to the bottom of the 1.5 mL tubes.
Note: Bones will become white and translucent if the bone cavity has been well flushed. If not, you can re-cut the bone ends and centrifuge again.
Discard the 0.5 mL microcentrifuge tubes that contain empty bones.
After the bone marrow has been collected in 1.5 mL tubes, transfer bone marrow to a 50 mL conical tube.
Add 10 mL of RPMI-1640 medium.
Make a single-cell suspension by drawing the bone marrow into a 10 mL syringe loaded with an 18 G needle. Do this 8–10 times.
Filter cell suspension through a 70 μm cell strainer into a 50 mL Falcon tube.
Centrifuge at 310× g for 10 min at 4 °C.
Aspirate the supernatant and resuspend the cell pellets in an appropriate volume of optimized separation buffer for the following lineage-negative cell isolation process.
Purification and lentivirus transduction of lineage-negative cells
Typically, lineage-negative cells account for 2%–5% of whole bone marrow nucleated cells. The purity is usually greater than 90% following isolation.
Isolate lineage-negative cells, using the Lineage Cell Depletion kit according to the manufacturer’s instructions.
Note: The purity of lineage-negative cells can be measured by flow cytometry. Lineages can be defined using specific makers such as CD3, CD90.2, CD19, B220, NK1.1, Ter119, Gr1 (Ly6G/Ly6C), and CD11b.
Resuspend the lineage-negative cells in 1 mL of serum-free hematopoietic cell expansion medium.
Seed the cells into an ultra-low attachment 6-well plate at a density of 1.5 × 106 cells/mL.
Add recombinant murine TPO and SCF into wells at final concentrations of 20 and 50 ng/mL, respectively.
Note: We routinely prepare small aliquots of recombinant proteins after reconstitution with water at 1,000× concentration (20 μg/mL for TPO and 50 μg/mL for SCF). Aliquots are stored at -80 °C and thawed as needed.
Pre-incubate cells at 37 °C in 5% CO2 for 2 h.
Gently add lentivirus at multiplicity of infection (MOI) = 100, 4 μg/mL polybrene infection/transfection reagent, and penicillin/streptomycin to the wells and incubate at 37 °C in 5% CO2 for 16–20 h.
Transplantation of transduced cells into lethally irradiated mice
Prepare recipient mice for transplantation by placing them in an eight-slice pie cage and exposing them to two doses of total body irradiation (550 Rad/dose, total dose = 1,100 Rad), with approximately 4 h between each irradiation session.
Note: The recipient mice should be male wild type.
Prepare the cells for injection by transferring all cells to a 15 mL conical tube. Add RPMI to 15 mL and centrifuge at 300× g for 10 min.
Suspend 5 × 105 lineage-negative cells in 200 μL of RPMI medium (5 × 105 cells/200 μL) and aspirate into an insulin syringe. Keep the cells at room temperature for 1–2 h until transplantation into lethally-irradiated recipient mice.
Note: Each mouse receives approximately 5 × 105 lineage-negative cells. These lineage-negative bone marrow cells were infected with a lentivirus vector at an MOI of 100, and then cultured for 16–20 h before collecting and washing as detailed previously in this section. All cells collected at this stage are injected into each recipient mouse.
Inject cells retro-orbitally into anesthetized lethally-irradiated recipient mice.
After the cells are injected, house the mice in sterilized cages and provide them with a soft diet and drinking water supplemented with antibiotics for 14 days.
Verification of Y chromosome deletion in hematopoietic cells
Fluorescent in situ hybridization (FISH)
FISH was performed in collaboration with Cell Line Genetics, Inc. and protocols utilized were provided by them.
Verify that the internal temperature of the 2× SSC pH 7.2 is 73 ± 1 °C and then add slides and incubate for 2 min.
Pepsin digest
i. Add 200 μL of Pepsin stock solution to the Coplin jar containing 40 mL of 0.01 N HCl in a 37 °C water bath and shake to mix well.
ii. Incubate slides for 13 min.
Wash the slides in 1× PBS for 5 min at room temperature.
Fix the slides in 1% formaldehyde for 5 min at room temperature.
Wash the slides in 1× PBS for 5 min at room temperature.
Dehydrate slides in 70%, 85%, and 100% ethanol for 2 min each and allow to air dry.
Notes:
i. Slides can be held at this point indefinitely. However, if left to stand for more than 2 h, repeat step f before probing.
ii. Perform the following steps under low light conditions.
Prepare the commercial probes for X chromosome (XqF4 region) and Y chromosome (YqA1 region).
Using a micro-pipettor, apply 3 μL of probe to the target area, cover with a circular coverslip, and seal with a bead of rubber cement.
Place the slides on the ThermoBrite instrument. Turn the instrument on and select the correct program to run (Program: 9, Denaturation: Temp 68, Time 2, Hybridization: Temp 37, Time 48 h).
Place slide in a moist hybridization chamber at 37 °C overnight (i.e., Tupperware-style container lined with a moist paper towel).
At the end of the day, add 40 mL of 0.4× SSC/0.3% NP-40 (pH = 7.2) into a plastic Coplin jar and place in a 74 °C water bath overnight. Make sure the lid is on tight to prevent evaporation.
Note: On the next day, perform the following steps under low light conditions.
Remove the rubber cement from the slides and slide the coverslips off gently with forceps.
Immediately transfer the slides to the Coplin jar of 0.4× SSC/0.3% NP-40 (pH = 7.2) stringency wash pre-warmed to 73 °C ± 0.5 °C for 2 min.
Drain the slides and wash in PBD in a Coplin jar for at least 30 s.
Drain the slide briefly and wipe the back of the slide.
Add 10–15 mL of 1:20 VectaShield with DAPI to each hybridization spot using the dropper, add the coverslip (22 mm × 50 mm), and blot the edges. Remove the excess DAPI stain by gently pressing on the slide sandwiched between a single layer of paper towels.
Store slides in the dark at -20 °C until ready to screen.
Karyotyping
We first immortalized lineage-negative cells by transducing them with lentivirus containing Hoxb8 (LV.T11.Hoxb8.Puro), before conducting karyotyping. In this system, the expression of Hoxb8 is induced by tetracycline, while the reverse transactivator M2 and the puromycin resistance genes are constitutively expressed in a bisistronic manner, driven by the human phosphoglycerate kinase promoter.
Karyotyping was performed in collaboration with KaryoLogic, Inc. Some information regarding the protocols used was provided by KaryoLogic, Inc. However, due to proprietary reasons, complete details of the protocols cannot be disclosed.
Immortalization of lineage-negative cells:
Prepare Hoxb8 lentivirus as described in section A.
Follow the procedure outlined in B.2 to isolate lineage-negative cells.
Culture the isolated lineage-negative cells in StemSpan SFEM medium supplemented with 50 ng/mL murine SCF (Stem Cell Factor), 100 ng/mL human IL-11 (Interleukin-11), 100 ng/mL human Flt3-ligand, and 20 ng/mL murine IL-3 (Interleukin-3).
Gently introduce HoxB8 lentivirus to the wells at a multiplicity of infection (MOI) = 100, with 4 μg/mL polybrene and 1% penicillin/streptomycin.
Incubate the cells at 37 °C in 5% CO2 for 16 h.
Collect the cells and resuspend them in IMDM supplemented with 10% heat-inactivated FBS, 1% penicillin/streptomycin, and cytokines (100 ng/mL murine SCF, 100 ng/mL human IL-11, 100 ng/mL human Flt3-Ligand, and 20 ng/mL murine IL-3) along with 2.4 μg/mL of doxycycline hyclate.
Note: It is recommended to prepare frozen aliquots in small amounts of cytokine solutions at a concentration of 1,000×.
Treat cells with puromycin at a concentration of 0.5 μg/mL for 48 h.
Sort the fraction of cells that are RFP-positive and culture them for an additional seven days before conducting karyotype analysis.
Karyotyping of immortalized lineage-negative cells:
Add Colcemid solution to the cell cultures to a final concentration of 0.5 μg/mL.
Note: In metaphase spreads, the cell is first treated with a chemical (e.g., colcemid) to arrest it at metaphase and then the chromosomes are spread out on a microscope slide and stained to allow for their visualization under a microscope.
Incubate the cells at 37 °C in 5% CO2 for 10 min.
Transfer the cells to centrifuge tubes. Centrifuge the cells at 500× g for 7 min.
Resuspend the cells in 0.075 M KCl hypotonic solution and incubate the cells at room temperature (approximately 22 °C) for 6 min. Centrifuge the cells at 500× g for 7 min.
Resuspend the cells in 3:1 methanol:acetic acid fixative and incubate at room temperature for 30 min. Centrifuge the cells at 500× g for 7 min.
Drop a single drop of cells in 0.5 mL of fixative onto each wet microscope slide to make the slides.
Bake the slides at 65 °C for 20 h.
Treat the slides with 0.1% trypsin-EDTA and stain them with Giemsa in Gurr’s buffer at pH 6.8.
Analyze metaphase spreads using a Leica DM2500 brightfield microscope at 1,000× and Leica Biosystems CytoVision software, version 7.4.
Data analysis
The Y chromosome ablation in hematopoietic cells from mLOY mice was confirmed by fluorescent in situ hybridization (FISH) analysis of X and Y chromosomes (Figure 2A: peripheral blood) and karyotype analysis (Figure 2B: bone marrow hematopoietic stem/progenitor cells). These data and the methods were derived from Figure 1 of the original article (Sano et al., 2022).
Figure 2. Depletion of the mouse Y chromosome with CRISPR/Cas9 system
In the original article, the control condition used a lentivirus encoding a gRNA designed not to target any region of the genome as a control (gRNA-NT). However, it was possible that cleavage of the genome itself may have an effect on the function of the transduced cells, regardless of the target genomic region. For example, studies have revealed that CRISPR/Cas9-mediated genome editing in normal cells can elicit a p53-induced DNA damage response (DDR), which induces the expression of downstream effectors, such as cell cycle inhibitor p21 (Haapaniemi et al., 2018; Janic et al., 2018). Thus, the expression of DDR-induced genes was assessed eight weeks after transplantation, using bone marrow lineage-negative cells from mice transplanted with gRNA that targets the centromere of the Y chromosome or gRNA-NT (Control). There was no difference between groups in the expression of p53, p21, and Bax (Figure 3).
Figure 3. Expression of DNA damage response (DDR)-related genes in lineage-negative bone marrow cells harvested from mice transplanted cells treated with Y chromosome centromere-targeting gRNA or non-targeting gRNA
Next, to further rule out the possible confounding effects described above, we compared the action of gRNA-NT with that of a gRNA that targets the fourth intron of the alpha-actin gene (gRNA-Actin-Int4). In this experiment, we examined whether either version of control gRNA could influence heart function in response to stress in mice that had undergone BMT with transduced lineage-negative cells. For this purpose, mice undergoing BMT with hematopoietic cells treated with gRNA-NT or gRNA-Actin-Int4 were subjected to pressure overload by performing the transverse aortic constriction procedure. Compared to the data from the mLOY test condition (Sano et al., 2022), echocardiographic analyses revealed no differences in cardiac function in mice treated with gRNA-NT compared with those treated with gRNA-Actin-Int4 (Figure 4). These results indicate that CRISPR/Cas9-mediated genome cleavage in hematopoietic cells has no effect on long-term cellular function, at least in terms of cardiac function, possibly due to the disappearance of gRNA target sites following the initial editing of the genome.
Figure 4. The impact of control gRNAs used in hematopoietic cells on the cardiac phenotype in response to pressure overload. FS; fractional shortening, PWTd; left ventricular posterior wall thickness at end-diastole, LVDs; left ventricular diameter at end-systole, LVDd; left ventricular diameter at end-diastole.
Recipes
Cell culture media
DMEM (high glucose) supplemented with 10% heat-inactivated FBS, 1% penicillin and streptomycin, warmed at 37 °C
Wash buffer for cells
PBS, warmed at 37 °C
MACS buffer
PBS supplemented with 0.5% BSA, 2 mM EDTA, chilled on ice
Acknowledgments
This work was supported by the National Institutes of Health (NIH) grants AG073249, AG072095, HL141256, HL139819, and HL142650 to K.W.; NIH grant HL152174 to S.S. and K.W.; and Grant-in-Aid for Research Activity Start-up 21K20879, Grant-in-Aid for Scientific Research C 22K08162, and grants from the Grant for Basic Research of the Japanese Circulation Society, the MSD Life Science Foundation, the Cardiovascular Research Fund, and the Japanese Heart Failure Society to S.S.
This protocol was derived from our previously published paper (Sano et al., 2022) with some modifications.
Competing interests
The authors declare no competing interests.
Ethical considerations
Animals were used in accordance with NIH guidelines and approved by the Institutional Animal Care and Use Committee at the University of Virginia.
References
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Haapaniemi, E., Botla, S., Persson, J., Schmierer, B. and Taipale, J. (2018). CRISPR-Cas9 genome editing induces a p53-mediated DNA damage response.Nat Med 24(7): 927-930.
Janic, A., Valente, L. J., Wakefield, M. J., Di Stefano, L., Milla, L., Wilcox, S., Yang, H., Tai, L., Vandenberg, C. J., Kueh, A. J., et al. (2018). DNA repair processes are critical mediators of p53-dependent tumor suppression.Nat Med 24(7): 947-953.
Sano, S., Horitani, K., Ogawa, H., Halvardson, J., Chavkin, N. W., Wang, Y., Sano, M., Mattisson, J., Hata, A., Danielsson, M., et al. (2022). Hematopoietic loss of Y chromosome leads to cardiac fibrosis and heart failure mortality. Science 377(6603): 292-297.
Sano, S., Wang, Y., Evans, M. A., Yura, Y., Sano, M., Ogawa, H., Horitani, K., Doviak, H. and Walsh, K. (2019). Lentiviral CRISPR/Cas9-Mediated Genome Editing for the Study of Hematopoietic Cells in Disease Models. J Vis Exp (152). doi: 10.3791/59977.
Thompson, D. J., Genovese, G., Halvardson, J., Ulirsch, J. C., Wright, D. J., Terao, C., Davidsson, O. B., Day, F. R., Sulem, P., Jiang, Y., et al. (2019). Genetic predisposition to mosaic Y chromosome loss in blood. Nature 575(7784): 652-657.
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Developmental Biology > Cell growth and fate > Ageing
Developmental Biology > Genome editing
Molecular Biology > DNA > Chromosome engineering
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4,730 | https://bio-protocol.org/en/bpdetail?id=4730&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Construction of Activity-based Anorexia Mouse Models
MM Maria Consolata Miletta *
TH Tamas L. Horvath *
(*contributed equally to this work)
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4730 Views: 461
Reviewed by: Pilar Villacampa Alcubierre Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Metabolism Oct 2020
Abstract
Anorexia nervosa (AN) is a psychiatric disorder mainly characterized by extreme hypophagia, severe body weight loss, hyperactivity, and hypothermia. Currently, AN has the highest mortality rate among psychiatric illnesses. Despite decades of research, there is no effective cure for AN nor is there a clear understanding of its etiology. Since a complex interaction between genetic, environmental, social, and cultural factors underlines this disorder, the development of a suitable animal model has been difficult so far. Here, we present our protocol that couples a loss-of-function mouse model to the activity-based anorexia model (ABA), which involves self-imposed starvation in response to exposure to food restriction and exercise. We provide insights into a neural circuit that drives survival in AN and, in contrast to previous protocols, propose a model that mimics the conditions that mainly promote AN in humans, such as increased incidence during adolescence, onset preceded by negative energy balance, and increased compulsive exercise. This protocol will be useful for future studies that aim to identify neuronal populations or brain circuits that promote the onset or long-term maintenance of this devastating eating disorder.
Keywords: Anorexia nervosa (AN) Activity-based anorexia model (ABA) AN animal model Hyperactivity Food restriction
Background
Anorexia nervosa (AN) is a severe psychiatric disorder characterized by extreme hypophagia, severe body weight loss, distorted self-image, and maladaptive food choices (Mitchell and Peterson, 2020). The onset of AN usually occurs in adolescence or young adulthood (Favaro et al., 2009) and it mainly affects females (roughly 92%) (Udo and Grilo, 2018). It has the highest mortality rate among psychiatric disorders (Treasure et al., 2015), 12 times higher than the death rate associated with all causes of death for females 15–24 years old (Smink et al., 2013; Fichter and Quadflieg, 2016). Currently, there are no effective pharmacological cures for AN (Crow, 2019). To inform new therapies and to identify individuals at risk for the disorder, a deeper understanding of the neurobiology underlying AN is urgently needed. The etiology of AN is complex since it involves interactions between genetic, environmental, social, and cultural factors. Such complexity has made it difficult to develop an appropriate animal model so far (Siegfried et al., 2003). Previous studies have used approaches involving genetic, environmental, and/or dietary manipulations to study AN in animal models. Genetically modified mouse models have been used to ablate or activate specific neuronal populations, with immediate reduction of food intake (Luquet et al., 2005; Calvez et al., 2011). While most of these studies might provide insights into neural circuits causing anorexia (simply seen as the loss of appetite for food), their translatability to humans is minimal since the nature and timing of the stressors are different from those that increase the risk of developing AN in humans (François et al., 2022).
The best characterized animal model of AN is the activity-based anorexia model (ABA) (Klenotich and Dulawa, 2012). This is a bio-behavioral phenomenon described in rodents that models the key symptoms of AN (self-induced starvation and compulsive running, to list a few of them) and is based on the principle that mice will prefer to run instead of eating even when food is available, if previously exposed to food restriction. The ABA, however, cannot capture the sociocultural factors that might promote AN.
Here, we propose a method to couple a genetically modified mouse model to a specific ABA protocol designed for adolescent female and male mice. We chose to investigate the involvement of the hypothalamic agouti-related peptide and neuropeptide Y (AgRP/NPY)-expressing neurons. These neurons are physiologically active during fasting, but also modulate complex non-feeding behaviors (Dietrich et al., 2015). To impair AgRP neuron function, we studied AgRP neuron-specific diphtheria toxin receptor-expressing mice (AgRP-DTR), which allow ablation of AgRP neurons early postnatally by administering diphtheria toxin at P5 (Luquet et al., 2005). By combining ABA with a transgenic mouse model and further choosing a critical time window (adolescence), it is feasible to get insights into genetic × environmental factors (such as early life stress) that contribute to the onset or propagation of AN. This protocol will be useful for future studies that aim to identify the contribution of specific neuronal populations to AN symptomatology. Combining the two approaches in adolescent mice will also allow the unraveling of mechanisms that contribute to resilience or vulnerability to AN.
Materials and reagents
Mouse cages (Tecniplast, catalog number: GM500)
Wire bar lids (Tecniplast, catalog number: GM500)
Bedding, Teklab corncob bedding (Envigo, catalog number: 7092)
Standard rodent diet, Teklab global 18% protein rodent diet (Envigo, catalog number: 2018S)
Paper towels
4-week-old female and male mice, Agouti-related protein mice (AgRP-DTR mice)
70% ethanol (VWR Chemicals, catalog number: 20842.298, to be dissolved in water to final concentration)
NaCl powder (Sigma, catalog number: 7647-14-5)
Diphtheria toxin (List Biological, catalog number: 150)
Equipment
In-cage running wheel (Starr Life Science Corporation, USA)
Infrared sensor for running wheel (Starr Life Science Corporation, USA)
Standard Windows PC (operating system Windows 7 or later)
DataPort1224 (DP1224) (Starr Life Science, USA)
Powered USB hub (Conrad Electronic AG, Digitus DA-70229 10 Port USB 2.0-Hub/catalog number:1027267-5k) (to provide sufficient USB ports and equipment spacing flexibility, one USB port support for each DP1224s)
Analytical balance (1 mg) (MettlerToledo, Balance XPE206DR, catalog number: 30132913)
Compact scale (0.1 g) (Haslab, HCB 302 Highland portable precision, catalog number: 105654302)
Red headlamp (Energizer, online vendor)
Software
VitalView Data Acquisition System software version 5.1 (Starr Life Science, USA https://www.starrlifesciences.com/product/activity-software/)
GraphPad Prism version 9 (GraphPad software, San Diego, CA, https://www.graphpad.com)
Note: Do not run other software on your Windows PC while VitalView is acquiring data.
Procedure
AgRP ablation
Weigh the mice at P5.
Dissolve the diphtheria toxin powder in water or saline (0.90% w/v of NaCl) to a final concentration of 5 μg/mL (stock solution).
Inject subcutaneously a dose of 5 μg/kg in heterozygous and homozygous mice for the selected mutation (expression of the diphtheria toxin receptor in AgRP neurons) (Figure 1). The preferable site for injection is over the shoulders, into the loose skin over the neck. A new needle should be used for each animal to reduce the risk of site infection. Allow full absorption of the substance before putting back the mouse into the cage.
Figure 1. Ablation of AgRP neurons by diphtheria toxin injections on P5 in the arcuate nucleus. Both control and AgRPDTR/+ mice were injected as pups with DT (50 μg/kg) on P5. Representative immunostaining of ARC neurons of control (WT) and AgRPDTR/+ mice are shown. The asterisks indicate third ventricle. Scale bar, 50 μm. AgRP: Agouti related peptide. DTR: diphtheria toxin receptor. ARC: arcuate.
Weigh the mice and their food intake daily up to day 21 to assess standard growth.
Activity-based anorexia model
Place cages in a temperature-controlled room dedicated only to ABA testing. Set the light timers to the 12:12 h light/dark cycle.
Cage setup: clean the cage and the running wheel with 70% ethanol and afterward dry with a paper towel. Add the bedding (up to 4 cm) and the in-cage running wheel. Verify that the wheel can freely turn without touching the bedding.
Insert the infrared sensor on the side of the running wheel and connect it to the DP1224.
Boot the computer and start the running wheel software by double-clicking on the icon.
Label every cage and the corresponding sensor in alphabetic or numerical order. Set up the configuration of the data acquisition (a detailed data acquisition software manual is available online from Starr Life website).
Single-house 36-day-old mice in a cage containing bedding, a water bottle, and ad lib food.
Allocate the mice in two groups:
Running group have ad lib access to food and water and access to the running wheel.
Sedentary group have ad lib access to food and water without access to the running wheel.
Acclimation: allow animals to acclimate for four days.
For the following four days, measure weight and food intake every morning.
Use a dedicated container for the food, to avoid food contamination with feces and urine. Minimize the time that each cage is moved, weighing first the food and afterward the mouse from the same cage. When removing the mouse from the cage, manually stop the running wheel to prevent recording biased running wheels counts.
Food restriction (72 h/three days).
Approximately at noon on the fourth day of acclimation, remove the food from both groups (sedentary and running).
Before the onset of the dark phase, pre-weigh 50 g of pellet. Store food at room temperature in cups labeled with the mouse ID and cover with aluminum foil. Use a fresh pellet for every day of food restriction to avoid contamination.
At 7:00 pm, when the lights are usually turned off, place the pre-weighed pellets in the overhead food bin of each cage for 2 h.
At 9:00 pm, carefully remove the leftover pellets, place it in the labeled containers, and cover with aluminum foil. Clean the food bin with a dry paper. Wear a red headlamp while you are in the room during the dark phase.
In the morning after food restriction, weigh the mice and the food intake from the leftover pellet. Ensure that there is no food left in the bedding of the cage.
To determine weight loss, calculate the % baseline body weight dividing the current weight by the body weight recorded on the last day of acclimation. Any mouse that loses more than 25% of its body weight is at risk of death and must be removed from the experiment. Mice removed from the experiment are placed in a new cage without the running wheel and with ad lib food and water.
Recovery: approximately at noon of the third day of food restriction, remove all remaining mice from the experiment and place them in a new cage without the running wheel and with ad lib food and water. Allow the animals to recover for at least one week before undergoing additional behavioral testing.
Under the File menu in the VitalView program window, click “stop acquisition.”
Data analysis
Body weight data
Perform a survival analysis (e.g., Kaplan-Meier) of both running and sedentary groups across the three days of food restriction. Percentage survival (on the y-axis) indicates the percentage of animals within each group that were not removed from the experiment. Mice removed earlier from the experiment because they lost more than 25% of their initial body weight show higher vulnerability to ABA. The survival analysis allows evaluation of any intervention (drug and/or diet) that might improve adaptation to ABA (Figure 2, right).
Both during acclimation and food restriction, calculate the body weight separately for females and males and express it as grams (Figure 2, left).
Figure 2. Body weight and survival curve data. Error bars indicate SEM; running(R)-control, n = 16; sedentary (S)-control, n = 12; R-AgRPDTR, n = 16; S-AgRPDTR, n = 14.
Food intake data
Calculate the food intake as kcal/day. For standard diet (7% simple sugars, 3% fat, 50% polysaccharide, 15% protein) energy = 3.5 kcal/g. During acclimation, food intake is indicated as kcal/day, while during food restriction is indicated as kcal/2 h (Figure 3).
Make separate calculations for males and females.
After calculating the average, compare running and sedentary groups using a two-way ANOVA, followed by multiple comparisons test.
Figure 3. Food intake. Food intake shown as kcal in 24 h during acclimation (left) or kcal in 2 h during food restriction (right). Error bars indicate SEM; running(R)-control, n = 16; sedentary (S)-control, n = 12; R-AgRPDTR, n = 16; S-AgRPDTR, n = 14.
Running wheel data
Use the VitalView Data Acquisition System software to extract the running wheel data. The program converts the data to an Excel file by selecting the Export option in the File menu. The running wheel count is reported by day. Calculate the average total running across the acclimation and the food restriction. Use a two-way ANOVA test for group comparisons followed by multiple comparisons test.
Additionally, and separately, calculate the average light cycle (12 h) and dark cycle (12 h) running for each group and use two-way ANOVA test for group comparisons followed by multiple comparisons test. In our as well as in different experimental paradigms, an abrupt increase of running during the light phase is associated with poor adaptation to the ABA and often precedes discharge from the experiment for most mice. Running that precedes presentation of food is indicated as food anticipatory activity.
Optional: Calculate the running wheel count over the 2-h period of food restriction and put them in correlation with the food intake. One of the features of ABA is that mice prefer to run instead of eating even if food is available.
Make separate calculations for males and females (Figure 4).
Figure 4. Running wheel data. Total running wheel count averaged across acclimation (days 1–4) and food restriction (days 5–7). Error bars indicate SEM; running(R)-control, n = 16; sedentary (S)-control, n = 12; R-AgRPDTR, n = 16; S-AgRPDTR, n = 14. The units for the cumulative (24 h) wheel count are revolutions per minute (RPM).
Notes
Notes on the Procedure
Carefully assess the weight of the mice before injection with the diphtheria toxin. In our experience, mice that weigh less than 2.5 g at P5 are more prone to die after the injection.
The temperature in the room dedicated to ABA should be strictly controlled and range from 22 to 25 °C. Room temperature has been shown to affect vulnerability to ABA (Gutiérrez et al., 2002).
In this study, we used littermate mice in the experiments. This choice ensures that both the genetic background and environment are comparable through several experiments.
Handling of animals should be kept to a minimum; if possible, one operator should handle the mice throughout the experiment. Food intake and weight should be measured at the same time each day. Scents and perfumes should be avoided. All the above precautions minimize animal stress exposure by reducing unpredictability.
Adolescent mice are more vulnerable to weight loss and death induced by starvation compared to adult mice or rats. Therefore, the current protocol applies some strategies to improve their survival during ABA. These include providing food for 2 h instead of 1 h during food restriction and limiting food restriction to 72 h. Using a different age group from that specified in this protocol may require changes in food restriction duration.
Give food-restricted mice a large pellet to reduce the possibility that food will fall into the cage and mice continue to eat beyond the allowed time window.
To prevent mouse dehydration, food can be provided wet during food restriction.
To prevent data loss, power the computer through a backup power supply. Even a short interruption of power will force the computer to restart, and data acquisition will stop.
Notes on Data analysis
It is always advisable to plot the data for female and male mice separately, since their food intake and body weight differ at the baseline and after food restriction.
All data shown in Figures have been previously published (Miletta et al., 2020).
Notes on validation
Technically, it is challenging to get a consistent level of AgRP ablation through several mice generations. However, being consistent about the time of the diphtheria toxin injection can minimize it.
Acknowledgments
This work was partly funded by the Swiss National Science Foundation (Early Postdoc.Mobility P2BEP3_172252 to M.C.M.), National Institutes of Health grants AG052005 and DK111178, and a Klarman Family Foundation grant (to T.L.H.). T.L.H. was also supported by grant NKFI-126998 from the Hungarian National Research, Development and Innovation Office. The described model was adapted from previous studies in which activity-based anorexia model was tested with different conditions (Klenotich and Dulawa, 2012). The mouse strain was first described by Luquet et al. (2005) but not in correlation with the ABA.
Competing interests
The authors have no financial or no-financial competing interests to report.
Ethics considerations
This animal protocol was approved by the Institutional Animal Care and Use Committee (IACUC) at Yale University (2022-07942).
References
Calvez, J., Fromentin, G., Nadkarni, N., Darcel, N., Even, P., Tome, D., Ballet, N. and Chaumontet, C. (2011). Inhibition of food intake induced by acute stress in rats is due to satiation effects. Physiol Behav 104(5): 675-683.
Crow, S. J. (2019). Pharmacologic Treatment of Eating Disorders. Psychiatr Clin North Am 42(2): 253-262.
Dietrich, M. O., Zimmer, M. R., Bober, J. and Horvath, T. L. (2015). Hypothalamic Agrp neurons drive stereotypic behaviors beyond feeding. Cell 160(6): 1222-1232.
Favaro, A., Caregaro, L., Tenconi, E., Bosello, R. and Santonastaso, P. (2009). Time trends in age at onset of anorexia nervosa and bulimia nervosa. J Clin Psychiatry 70(12): 1715-1721.
Fichter, M. M. and Quadflieg, N. (2016). Mortality in eating disorders - results of a large prospective clinical longitudinal study. Int J Eat Disord 49(4): 391-401.
François, M., Fernandez-Gayol, O. and Zeltser, L. M. (2022). A Framework for Developing Translationally Relevant Animal Models of Stress-Induced Changes in Eating Behavior. Biol Psychiatry 91(10): 888-897.
Gutiérrez, E., Vazquez, R. and Boakes, R. A. (2002). Activity-based anorexia: ambient temperature has been a neglected factor. Psychon Bull Rev 9(2): 239-249.
Klenotich, S. J. and Dulawa, S. C. (2012). The activity-based anorexia mouse model. Methods Mol Biol 829: 377-393.
Luquet, S., Perez, F. A., Hnasko, T. S. and Palmiter, R. D. (2005). NPY/AgRP neurons are essential for feeding in adult mice but can be ablated in neonates. Science 310(5748): 683-685.
Miletta, M. C., Iyilikci, O., Shanabrough, M., Sestan-Pesa, M., Cammisa, A., Zeiss, C. J., Dietrich, M. O. and Horvath, T. L. (2020). AgRP neurons control compulsive exercise and survival in an activity-based anorexia model. Nat Metab 2(11): 1204-1211.
Mitchell, J. E. and Peterson, C. B. (2020). Anorexia Nervosa. N Engl J Med 382(14): 1343-1351.
Siegfried, Z., Berry, E. M., Hao, S. and Avraham, Y. (2003). Animal models in the investigation of anorexia. Physiol Behav 79(1): 39-45.
Smink, F. R., van Hoeken, D. and Hoek, H. W. (2013). Epidemiology, course, and outcome of eating disorders. Curr Opin Psychiatry 26(6): 543-548.
Treasure, J., Zipfel, S., Micali, N., Wade, T., Stice, E., Claudino, A., Schmidt, U., Frank, G. K., Bulik, C. M. and Wentz, E. (2015). Anorexia nervosa. Nat Rev Dis Primers 1: 15074.
Udo, T. and Grilo, C. M. (2018). Prevalence and Correlates of DSM-5-Defined Eating Disorders in a Nationally Representative Sample of U.S. Adults. Biol Psychiatry 84(5): 345-354.
Article Information
Copyright
© 2023 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 > Animal model
Medicine > Anorexia nervosa > Mouse model
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4,731 | https://bio-protocol.org/en/bpdetail?id=4731&type=0 | # Bio-Protocol Content
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Peer-reviewed
Ex vivo Drug Sensitivity Imaging-based Platform for Primary Acute Lymphoblastic Leukemia Cells
LR Lauren Rowland
BS Brandon Smart
AB Anthony Brown §
GD Gino M. Dettorre
YG Yoshihiro Gocho
JH Jeremy Hunt
WY Wenjian Yang
SY Satoshi Yoshimura
NR Noemi Reyes
GD Guoqing Du
AJ August John
DM Dylan Maxwell
WS Wendy Stock
SK Steven Kornblau
MR Mary V. Relling
HI Hiroto Inaba
CP Ching-Hon Pui
JB Jean-Pierre Bourquin
SK Seth E. Karol
CM Charles G. Mullighan
WE William E. Evans
JY Jun J. Yang
KC Kristine R. Crews
(§ Technical contact)
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4731 Views: 1144
Reviewed by: Rajesh RanjanSalah BoudjadiDipak Kumar Poria
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Original Research Article:
The authors used this protocol in Nature Cancer Jan 2021
Abstract
Resistance of acute lymphoblastic leukemia (ALL) cells to chemotherapy, whether present at diagnosis or acquired during treatment, is a major cause of treatment failure. Primary ALL cells are accessible for drug sensitivity testing at the time of new diagnosis or at relapse, but there are major limitations with current methods for determining drug sensitivity ex vivo. Here, we describe a functional precision medicine method using a fluorescence imaging platform to test drug sensitivity profiles of primary ALL cells. Leukemia cells are co-cultured with mesenchymal stromal cells and tested with a panel of 40 anti-leukemia drugs to determine individual patterns of drug resistance and sensitivity (“pharmacotype”). This imaging-based pharmacotyping assay addresses the limitations of prior ex vivo drug sensitivity methods by automating data analysis to produce high-throughput data while requiring fewer cells and significantly decreasing the labor-intensive time required to conduct the assay. The integration of drug sensitivity data with genomic profiling provides a basis for rational genomics-guided precision medicine.
Key features
• Analysis of primary acute lymphoblastic leukemia (ALL) blasts obtained at diagnosis from bone marrow aspirate or peripheral blood.
• Experiments are performed ex vivo with mesenchymal stromal cell co-culture and require four days to complete.
• This fluorescence imaging–based protocol enhances previous ex vivo drug sensitivity assays and improves efficiency by requiring fewer primary cells while increasing the number of drugs tested to 40.
• It takes approximately 2–3 h for sample preparation and processing and a 1.5-hour imaging time.
Graphical overview
BM: bone marrow; PB: peripheral blood; ALL: acute lymphoblastic leukemia; MNCs: mononuclear cells, which include leukemia cells when present; MSCs: mesenchymal stromal cells; LC50: drug concentration that kills 50% of the leukemia cells
Keywords: Ex vivo drug sensitivity Functional precision medicine Acute lymphoblastic leukemia Pharmacogenomics Pharmacotyping Fluorescence imaging
Background
Acute lymphoblastic leukemia (ALL) is a heterogenous disease that can be grouped into multiple B- and T-lineage genomic subtypes (Inaba and Mullighan, 2020; Inaba and Pui, 2021; Brady et al., 2022). While five-year survival rates for many pediatric B-cell ALL (B-ALL) subtypes can exceed 90%, survival rates are lower for adults who have more treatment-resistant B-cell ALL, particularly those with Ph-like ALL (Foà et al., 2011). Similarly, survival for certain subtypes of T-cell ALL (T-ALL) remains below 80% (Karrman and Johansson, 2017) and outcomes following relapse for both pediatric and adult patients with ALL remain dismal (Pocock et al., 2021; Laukkanen et al., 2022; Wudhikarn et al., 2022). Because of its high incidence, ALL remains the most common cause of death from childhood cancer. Treatment efficacy in ALL varies by molecular subtype and secondary genomic alterations (Pui et al., 2012; Brady et al., 2022). Progress in understanding the heterogeneity of the genomics of ALL has provided insight into actionable drug targets (Meyer et al., 2013; Irving et al., 2014; Khaw et al., 2016; Gocho et al., 2021; Brady et al., 2022). Molecularly targeted agents have shown efficacy in some subtypes of high-risk B-ALL disease. There is a need for further development of molecularly targeted therapy optimized based on characteristics of an individual patient’s cancer cells (Carroll et al., 2003; Brady et al., 2022; Lee et al., 2023).
Functional precision medicine approaches combine direct drug sensitivity profiling (i.e., pharmacotyping) with genomic testing to elucidate the biological basis of variability in treatment response (Frismantas et al., 2017; Kornauth et al., 2022; Lee et al., 2023). Defining inter-patient variability in leukemia cell drug sensitivity is a starting point for observing patterns of sensitivity to antileukemia agents. Drug sensitivity profile groups have been associated with treatment outcome, including event-free survival and cumulative risk of relapse (Holleman et al., 2004; Pieters et al., 2007; Lee et al., 2023). ALL drug sensitivity phenotype has likewise been associated with clinical characteristics and somatic genomic features (Pieters et al., 1993; Ramakers-van Woerden et al., 2002; Holleman et al., 2004; Ramakers-van Woerden et al., 2004; Lugthart et al., 2005; Pui et al., 2019). Pharmacotyping profiles have the potential to drive research to uncover new subtype-specific therapeutic opportunities for ALL.
Both flow cytometry–based and the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazoliumbromide (MTT)–based drug-resistance assays have served as reliable methods in our laboratory for quantification of B- and T-ALL leukemia cell viability upon exposure to antineoplastic agents (Holleman et al., 2004; Paugh et al., 2015; Autry et al., 2020; Gocho et al., 2021; Lee et al., 2023). We have used flow cytometry–based methods and MTT-based methods for sensitivity testing of leukemia cell lines, patient-derived xenografts (PDX), and primary patient samples; however, in our experience, these assays are markedly limited by requirements for high cell count at plating, prolonged time requirements to complete the assay and data analyses, and lack of high-throughput potential for screening of antineoplastic drugs against samples (Gocho et al., 2021). Ex vivo models have used human bone marrow mesenchymal stromal cells (MSC) to support leukemia cells in culture for functional experiments (Boutter et al., 2014). Other groups have published functional precision medicine methods using automated microscopic imaging to test drug sensitivity of PDX cells from clinically relevant ALL subgroups (Frismantas et al., 2017). Here, our group has established an ex vivo pharmacotyping platform to directly profile leukemia cells from individual patients for drug sensitivity using high-content imaging. Coupling data derived from this platform with patient genomic profiles has the potential to catapult the development of new ALL therapy using actionable target and biomarker discovery.
Here, we present our methodology for a high-throughput imaging-based assay for leukemia cell viability to identify effective cytotoxic and targeted anti-leukemia agents. Using a biologically realistic leukemia cell and MSC co-culture as described by other groups (Frismantas et al., 2017), our imaging system has produced high-throughput viability data with potential for further image analyses. This method is used to assess inter-patient differences in drug sensitivity related to patient age, sex, molecular subtypes of leukemia, and genomics, and to discover mechanisms of drug resistance in patients with ALL (Autry et al., 2020; Gocho et al., 2021; Lee et al., 2023). This knowledge may in turn be used to build more robust precision medicine approaches to improve cure rates, and to create future avenues to facilitate the rational use of molecularly targeted therapy. This protocol includes methods for this imaging-based functional precision medicine assay.
Materials and reagents
Biological materials
hTERT immortalized human bone marrow mesenchymal stromal cells (MSCs) (Applied Biological Materials, Inc., catalog number: T0523)
Reagents
AIM-V medium, 1,000 mL (Gibco, catalog number: 12055-083)
Hydrocortisone-water soluble (Sigma, catalog number: H0396)
RPMI 1640 medium, 500 mL (Gibco, catalog number: 11875-093)
Fetal bovine serum, certified, heat inactivated (Gibco, catalog number: 10082-147)
DMSO (Sigma, catalog number: D2660-100ML)
PBS, pH 7.4, 500 mL (Gibco, catalog number: 10010-023)
CyQUANTTM Direct Cell Proliferation Assay Green (Invitrogen, catalog number: C35012)
ViaStain AOPI staining solution (Nexcelom, catalog number: CS2-0106-25mL)
Trypsin-EDTA 0.25% (Gibco, catalog number: 25200-056)
Antibiotic antimycotic 100× (Gibco, catalog number: 15240-062)
RPMI 1640 without phenol red & L-glutamine (Lonza, catalog number: 12-918F)
0.5 M EDTA pH 8.0 (Gibco, catalog number: 15575-038)
0.9% sodium chloride (Baxter, catalog number: 2F7123)
Insulin-transferrin-selenium 100× (Gibco, catalog number: 41400045)
L-Glutamine 200 mM (Gibco, catalog number: A2916801)
0.1 N sodium hydroxide (Fisher Scientific, catalog number: AC124190010)
AmbionTM nuclease-free water (Invitrogen, catalog number: AM9937)
Ultrapure (Type 1) water
Drug stock solutions (Table 1)
Table 1. Drug stock solutions
# Name Vendor Catalog number Stock concentration Storage Diluent Concentration range Dilution factor
1 Trametinib MedChemExpress HY-10999 10 mM -80 °C DMSO 0.01–1,000 nM 10
2 Gilteritinib MedChemExpress HY-12432 10 mM -80 °C DMSO 0.1–10,000 nM 10
3 Venetoclax MedChemExpress HY-15531 10 mM -80 °C DMSO 0.001–100 nM 10
4 Inotuzumab Pfizer NA 0.25 mg/mL -80 °C Nuclease-free water 0.0025–250 ng/mL 10
5 Palbociclib MedChemExpress HY-50767A 10 mM -80 °C DMSO 0.1–10,000 nM 10
6 CHZ868 MedChemExpress HY-18960 10 mM -80 °C DMSO 0.1–10,000 nM 10
7 Dasatinib Selleckchem S1021 10 mM -80 °C DMSO 0.1–10,000 nM 10
8 Ibrutinib MedChemExpress HY-10997 50 mM -80 °C DMSO 1.5625–50 mM 2
9 6-mercaptopurine Acros Organics 226520050 5 mg/mL -20 °C 0.1 N NaOH 91.8–2,938 mM 2
10 Prednisolone Pfizer 00009004722 12.5 mg/mL -20 °C 0.9% NaCl 0.015–503.5 mM 8
11 L-asparaginase BioVendor RP1792872500 500 IU/mL -80 °C Type 1 water 0.0032–10 IU/mL 5
12 Vincristine Hospira 61703030906 0.25 mg/mL -20 °C Type 1 water 0.0017–54.169 mM 8
13 Nelarabine Sigma-Aldrich SML1736 200 mM -20 °C DMSO 1.03–250 mM 3
14 Bortezomib St. Jude Compound Management SJ000518968 10 mM -80 °C DMSO 0.98–1,000 nM 4
15 Vorinostat Sellekchem S1047 100 mM -80 °C DMSO 102.88–25,000 nM 3
16 Daunorubicin Teva 00703523313 0.1 mg/mL -20 °C Type 1 water 0.004–3.55 mM 4
17 Luminespib MedChemExpress HY-10215 10 mM -80 °C DMSO 0.01–1,000 nM 10
18 Clofarabine MedChemExpress HY-A0005 10 mM -80 °C DMSO 0.1–10,000 nM 10
19 Olaparib MedChemExpress HY-10162 10 mM -80 °C DMSO 0.1–10,000 nM 10
20 Selinexor Selleckchem S7252 10 mM -80 °C DMSO 0.1–10,000 nM 10
21 Ponatinib MedChemExpress HY-12047 10 mM -80 °C DMSO 0.1–10,000 nM 10
22 Saracatinib MedChemExpress HY-10234 10 mM -80 °C DMSO 0.1–10,000 nM 10
23 Acalabrutinib MedChemExpress HY-17600 10 mM -80 °C DMSO 0.1–10,000 nM 10
24 Momelotinib MedChemExpress HY-10961 10 mM -80 °C DMSO 0.1–10,000 nM 10
25 Midostaurin MedChemExpress HY-10230 10 mM -80 °C DMSO 0.1–10,000 nM 10
26 Fostamatinib MedChemExpress HY-13038A 10 mM -80 °C DMSO 0.1–10,000 nM 10
27 Larotrectinib MedChemExpress HY-12866 10 mM -80 °C DMSO 0.1–10,000 nM 10
28 Repotrectinib MedChemExpress HY-103022 10 mM -80 °C DMSO 0.1–10,000 nM 10
29 Sorafenib MedChemExpress HY-10201 10 mM -80 °C DMSO 0.1–10,000 nM 10
30 Everolimus MedChemExpress HY-10218 10 mM -80 °C DMSO 0.1–10,000 nM 10
31 Sotorasib (AMG-510) MedChemExpress HY-114277 10 mM -80 °C DMSO 0.1–10,000 nM 10
32 Birinapant MedChemExpress HY-16591 10 mM -80 °C DMSO 0.1–10,000 nM 10
33 SNDX-5613 MedChemExpress HY-136175 10 mM -80 °C DMSO 0.1–10,000 nM 10
34 Iadademstat MedChemExpress HY-12782T 10 mM -80 °C DMSO 0.1–10,000 nM 10
35 EPZ4777 (EPZ004777) MedChemExpress HY-15227 10 mM -80 °C DMSO 0.1–10,000 nM 10
36 JQ1 MedChemExpress HY-13030 10 mM -80 °C DMSO 0.1–10,000 nM 10
37 Eprenetapopt (APR-246) MedChemExpress HY-19980 10 mM -80 °C DMSO 0.1–10,000 nM 10
38 S63845 MedChemExpress HY-100741 10 mM -80 °C DMSO 0.001–100 nM 10
39 Navitoclax MedChemExpress HY-10087 10 mM -80 °C DMSO 0.01–1,000 nM 10
40 Carfilzomib MedChemExpress HY-10455 10 mM -80 °C DMSO 0.0001–10 nM 10
Solutions
1 mM hydrocortisone stock solution (see Recipes)
Cell medium (see Recipes)
MSC medium (see Recipes)
Recipes
1 mM hydrocortisone stock solution
Reagent Final concentration Amount
Hydrocortisone-water soluble 1 mM 100 mg
Type 1 water n/a 23.45 mL
Total n/a 23.45 mL
Cell medium
Reagent Final concentration Amount
RPMI w/o phenol red & L-glutamine n/a 500 mL
Antibiotic antimycotic 1× 6.25 mL
L-Glutamine 2 mM 6.25 mL
Insulin-transferrin-selenium 1× 6.25 mL
Heat-inactivated fetal bovine serum 20% 125 mL
Total n/a 643.75 mL
MSC medium
Reagent Final concentration Amount
1 mM hydrocortisone-water soluble 1 μM 0.625 mL
RPMI-1640 n/a 500 mL
Heat-inactivated fetal bovine serum 20% 125 mL
Total n/a 625.625 mL
Laboratory supplies
384-well plates (PerkinElmer, Phenoplate, catalog number: 6057302)
10 mL pipettes (Corning, catalog number: 4101)
25 mL pipettes (Corning, catalog number: 4237)
15 mL conical tubes (Falcon, catalog number: 352097)
50 mL conical tubes (Falcon, catalog number: 352098)
Microtubes 1.5 mL (Axygen, catalog number: MCT-150-L-C)
Microtubes 0.6 mL (Axygen, catalog number: MCT-060-L-C)
Stericup quick release Millipore express 0.22 μM PES 500 mL (Millipore, catalog number: 6346)
10 cm dishes (Corning, catalog number: 353003)
20 μL filter tips (Rainin, catalog number: 30389225)
200 μL filter tips (Rainin, catalog number: 30389239)
1,000 μL filter tips (Rainin, catalog number: 30389212)
125 μL tips XYZ rack of 384 griptips (Integra, catalog number: 6465)
125 μL tips ECO rack of 384 griptips (Integra, catalog number: 3425)
150 mL reagent reservoirs (Integra, catalog number: 6318)
Equipment
Operetta CLSTM high-content analysis system (PerkinElmer, HH1600000)
STR4 plus series centrifuge (Thermo Scientific, catalog number: 75016037)
BX41 microscope (Olympus, catalog number: BX41-PH-B)
Tuberoller (Benchmark, catalog number: R3010)
Nikon Eclipse TS100 microscope (Nikon, catalog number: NI-TS100)
IsoTemp digital controlled water bath (Fisher Scientific, model: 2320)
TC10 automated cell counter (Bio-Rad, catalog number: 145-0010)
Sorvall Legend X1R centrifuge (Thermo Scientific, catalog number: 75004260)
Heracell 150i CO2 incubator (Thermo Scientific, catalog number: 51032719)
VIAFLO 384 automated pipette (Integra, catalog number: 6031)
VIAFLO 384-well 5–125 μL pipette head (Integra, catalog number: 6132)
VOYAGER 12-channel 5–125 μL pipette (Integra, catalog number: 4732)
1300 Series Class II A2 biological safety cabinet (Thermo Scientific, catalog number: 1323TS)
Software
Harmony software (4.9, 2019)
Prism v9.3 (GraphPad, 11/15/2021)
Microsoft office suite
Procedure
Procedure overview
Note: AOPI staining in the drug treatment wells as well as in the quality control wells would allow direct determination of cell death; however, the method described herein uses the more cost-efficient CyQUANT stain in the drug treatment wells to keep the assay costs down.
Drug response of primary human leukemia cells is evaluated using a co-culture system and a fluorescence imaging–based assay, building on a previously published assay (Boutter et al., 2014) with adaptations as described herein. hTERT immortalized MSCs are first seeded in a 384-well plate. After 24 h, leukemia cells are added to the stromal cell layer in AIM-V medium at a ratio of 1:10 MSCs to leukemia cells, along with a drug solution prepared in the same medium at six drug concentrations in duplicate. After a 96-hour incubation, images are acquired with a 20× objective, and the total number of live leukemia cells is evaluated. The leukemia cells and MSCs are analyzed in the same fluorescence channel based on the size and intensity of the nuclei. Drug-induced inhibition at each concentration is estimated by comparing to leukemia cells treated with the vehicle alone. The drug concentration that kills 50% of the leukemia cells (LC50) is determined using a dose-response model and is used as a measurement of drug sensitivity. Quality control is performed to remove cases with low viability (defined as less than 500 viable blast cells in each well in the absence of drugs on day 4).
Study approval
This study was approved by the respective institutional review boards at St. Jude Children’s Research Hospital, MD Anderson Cancer Center, and University of Chicago, and written informed consent was obtained from parents, guardians, and/or patients, as appropriate, before enrollment, in accordance with the Declaration of Helsinki.
Items to prepare in advance
To help with the following procedure, it is advised to prepare MSC medium and cell medium before starting. It is also recommended to prepare drug stocks to the concentrations stated in Table 1 or other concentrations desired.
MSC cell culture procedure
Grow and expand the MSC cells. MSC cells are cultured in MSC medium as described in Recipes.
Store MSC cells in incubator at 37 °C.
Twenty-four hours before receipt of a fresh primary sample, prepare MSC plate as below.
Remove a 10 cm dish of MSC from the incubator. Aspirate old medium.
Add 5 mL of PBS to the MSC dish. Rinse the monolayer by gently rocking the dish.
Aspirate all PBS from dish.
Slowly add 1 mL of trypsin-EDTA to the dish. Spread evenly by gently rocking the dish.
Incubate dish for 3 min in incubator (37 °C) until cells release from plate.
Microscopically verify that cells have released from the plate. Cells should be rounded and move when the dish is gently tapped.
Add 9 mL of MSC medium to the dish. Resuspend cells and medium 3–5 times, then transfer the suspension to a 15 mL (or 50 mL) Falcon tube.
Centrifuge at 300× g for 5 min at 4 °C.
While cells are spinning, prepare a 384-well plate by adding 50 μL of PBS per well to outside border wells.
Remove cells from centrifuge and aspirate the supernatant.
Resuspend cells in 5–10 mL of MSC medium.
Aliquot 10 μL of cells and count cells using an automated cell counter.
Calculate volume of MSC medium needed to dilute to a 1 × 105 cells/mL cell density.
Add appropriate volume of MSC medium to the Falcon tube containing cells.
Transfer 25 μL of the MSC suspension per well (= 2.5 × 103 cells/well) using a multichannel pipette.
Place plate in incubator and incubate overnight (37 °C, 5% CO2).
Notes before starting
AIM-V media that will be used for MSC plate washing and sample preparation should be warmed to 37 °C prior to use.
Primary sample procedure
Note: This method is for fresh cells processed within 24 h of collection. If the sample will not be plated on day of collection, keep cells rotating at 4 °C until ready to do so.
Leukemia blasts are obtained from either bone marrow or peripheral blood samples after undergoing Ficoll gradient centrifugation. The readers are referred to the Methods of the following references: Gocho et al. (2021), Lee et al. (2023).
To proceed to testing with the pharmacotyping assay, the sample must meet the following parameters: viability above 80%, red blood cell (RBC) contamination below 30%, and a blast percentage above 85%.
If the sample has more than 30% RBC contamination but above 85% blasts, the RBCs are removed from the sample through RBC depletion using an institutional procedure (see Supplemental Methods).
If the sample has a blast percentage of less than 85%, the sample is enriched for leukemia by using magnetic cell separation (Miltenyi Biotec) with a negative selection for leukemia cells using an institutional procedure (see Supplemental Methods).
Keep primary sample suspended in cell medium and rotating at 4 °C until ready to proceed.
Day 0 cell culture procedure
Note: While this protocol is designed for primary patient samples, it can be adapted for experiments with cell lines.
Prepare drug dilutions in AIM-V medium to achieve the final concentration ranges listed in Table 1.
Note: AIM-V media does not need to be warmed to 37 °C for drug plate preparation.
Remove the 384-well imaging plate containing MSCs from the incubator.
Aspirate the 25 μL of MSC medium from each well using the VIAFLO 384 automated pipette with a 384-channel head on a speed setting of 4 or lower to prevent MSC detachment.
Note: Automated pipettes are not required for aspiration steps described in this procedure; however, if done manually, make sure to remove media slowly and without touching the bottom of the well to prevent MSC removal.
Wash each well by adding 40 μL of AIM-V medium.
Centrifuge primary sample at 300× g for 5 min at 4 °C.
Aspirate cell medium of primary cells and flick pellet to resuspend cells.
Resuspend primary patient samples at 6.25 × 105 cells/mL in AIM-V medium for plating in the 384-well MSC plate.
Aspirate the AIM-V medium from each well using the VIAFLO 384 automated pipette on a speed setting of 4 or lower.
Add 40 μL of AIM-V medium to the MSC-only control wells.
Add 40 μL of the cell suspension to every well except the MSC-only control wells.
Add 10 μL of the drug dilutions to the designated drug wells (Figure 1).
Figure 1. Example plate layout
Add 10 μL of AIM-V medium to wells in Columns 2 and 3 in Figure 1.
Incubate the plate at 37 °C (5% CO2) for 96 h for primary patient samples.
Day 4 procedure
Note: Stain each plate individually so that incubation times are ~15 min, to reduce the chance of MSCs detaching from the plate; longer (> 1 h) incubation times are associated with detachment of MSCs, which may cause an experiment to fail.
Dilute CyQUANT Direct nucleic acid stain 1:26 and the CyQUANT Direct background suppressor 1:5.2 in PBS.
Note: Add the suppressor to the PBS first, because the nucleic acid stain will separate out of solution if added to PBS without the suppressor.
Add 5 μL of the CyQUANT staining solution to each well except AOPI control well.
Note: Verify visually that CyQuant was added to every well before moving to the next step. This can be done by checking the color of the well.
Add 50 μL of AOPI staining solution to the AOPI control well for viability analysis.
If desired, measure blast percentage of samples in Blast QC wells using Cytospin or institutional procedure.
Incubate the plate at 37 °C (5% CO2) for 15 min.
Critical step: Prolonged incubation with CyQuant can lead to cell apoptosis, which can influence the results of the assay.
The CyQUANT imaging procedure
Note: The imaging procedure described here applies specifically to the Operetta CLS; however, imaging can be performed with any imager that can acquire images in the green channel from a 384-well plate.
Remove imaging plate from incubator.
Spray a Kimwipe with 70% alcohol and wipe the bottom of the plate, as dust particles on the bottom of the plate may cause autofocusing errors during imaging.
Click the Eject button in Harmony software.
Properly position the plate in the plate holder of the imager.
Click the Load Plate button in the Harmony software.
Click the Setup tab in the Harmony software.
Click New in Experiment window to create a new experiment.
Use the following settings for the experiment:
Plate Type: 384 PerkinElmer CellCarrier Ultra
Autofocus: Two Peak (Default)
Objective: 20× Air, NA 0.4
Opt. Mode: Confocal
Binning: 2
In the Channel Selection window, add the Brightfield and CyQuant Green channels with the following settings:
Note: CyQuant Green may not be a pre-provided channel. To create the CyQuant Green channel, use wavelengths 460–490 for excitation and 500–550 for emission. Green Fluorescent Protein (eGFP) has similar excitation and emission wavelengths.
Brightfield:
i. Time = 20 ms
ii. Power = 60%
iii. Height = 8.0 μM
CyQuant Green:
i. Time = 60 ms
ii. Power = 100%
iii. Height = 8.0 μM
In the Navigation window, under the define layout tab, highlight all the wells that will be imaged and click Select. Within the Well window, highlight the nine inner squares, which correspond to the number and position of images being taken per well, and click Select. If the well/field of view has only been highlighted, it will be orange, but if it has been selected, it will be grey.
To take a snapshot: highlight one of the control wells and a single field of view. Then, within the Channel Selection window, click Snapshot in any of the channels to take one image to check if the leukemia cells are in focus. If cells are not in focus, change the heights for CyQuant Green and Brightfield channels until the cells are in focus (Figure 2).
Figure 2. Fluorescence imaging using a 20× objective of a primary patient leukemia sample overlaid with mesenchymal stromal cells (MSCs) stained with CyQUANT Green. White arrows indicate MSCs, and red arrows indicate leukemia cells.
Save the experiment.
Note: Once experiments have been saved, they can be reused by clicking the “…” icon to the right of the experiment name.
Click the Run Experiment tab in the Harmony software.
Fill out Plate Name with desired file name as this is what the measurement will be titled.
Click Start to run measurement.
AOPI imaging procedure
Click the Setup tab in the Harmony software.
Click New in experiment window to create a new experiment.
Use the following settings for the experiment:
Plate type: 384 PerkinElmer CellCarrier Ultra
Autofocus: Two Peak (Default)
Objective: 20× Air, NA 0.4
Opt. Mode: Confocal
Binning: 2
In the Channel Selection window, add the Acridine Orange, Propidium iodide, and Brightfield channels with the following settings:
AO:
i. Time = 40 ms
ii. Power = 80%
iii. Height = 8.0 μM
PI:
i. Time = 20 ms
ii. Power = 60%
iii. Height = 8.0 μM
Brightfield
i. Time = 20 ms
ii. Power = 60%
iii. Height = 8.0 μM
In the Navigation window, under the Define Layout tab, highlight all the wells that will be imaged and click Select. Within the Well window, highlight the nine inner squares, which correspond to the number and position of images being taken per well, and click Select. If the well/field of view has only been highlighted, it will be orange, but if it has been selected, it will be grey.
To take a snapshot: highlight one of the control wells and a single field of view. Then, within the Channel Selection window, click Snapshot in any of the channels to take one image to check if the leukemia cells are in focus. If cells are not in focus, change the heights for AO, PI, and Brightfield channels until the cells are in focus (Figure 3).
Figure 3. Overlay of primary patient leukemia cells and mesenchymal stromal cells (MSCs) stained with acridine orange/propidium iodide (AO/PI) for calculating percent cell viability. White arrows indicate MSCs, red arrows indicate dying leukemia cells, and yellow arrows indicate live leukemia cells.
Save the experiment.
Click the Run Experiment tab in the Harmony software.
Fill out Plate Name with desired file name as this is what the measurement will be titled.
Click Start to run measurement.
Data analysis
Data evaluation
The Harmony software for the PerkinElmer Operetta CLSTM high-content analysis system is used for the data analysis of the primary samples. The analysis pipeline distinguishes viable leukemia nuclei from MSC nuclei by setting nuclei area and CyQUANT fluorescence intensity cutoffs. The pipeline identifies and draws an outline of the nuclei in an image. Any nuclei that touch the border of the image are removed. Next, the CyQUANT intensity and area of the nuclei are measured. Manually select the viable leukemia nuclei with intensity values above 2,000 and an area less than 70 μm2 (Figure 4) (Note: Intensity cutoff values and/or area cutoff values may be adjusted for individual samples). Nine fields of view per well at a 20× magnification are evaluated for both the control and drug treatment wells. The data output is the number of viable leukemia cells per well. This output is copied into a Microsoft Excel analysis template, in which calculations are made for the relative viabilities of each drug treatment in comparison to the average of the untreated control wells.
Figure 4. Representative scatter plot of an analysis pipeline. Perpendicular lines represent cutoffs for nuclei size and nuclei intensity. Green dots indicate cells gated as leukemia cells and red dots indicate cells gated as mesenchymal stromal cells (MSCs).
Data analysis procedure for CyQUANT
Note: Attach an assay layout to the measurement you are analyzing before starting data analysis.
Create an assay layout by clicking the Settings button in the Harmony software and then select the Assay Layout Editor.
Complete the Compound, Concentration, Cell Type, and Cell Count entries that match your experiment plate layout.
Save the assay layout.
Attach assay layout to desired experiment.
Click the Image Analysis tab in the Harmony software.
Click the button to the right of Analysis and load the analysis pipeline you would like to use or, if setting up a new pipeline, press the New button under Measurement.
For setting a new pipeline:
Press the “+” button at the bottom of the tab to add a building block to your pipeline.
Click Find Nuclei.
Under Find Nuclei, select the channel you would like to use and the method that best labels the cells in your image.
Label the output population as Nuclei.
Press the “+” button again and click Select Population.
Select the Remove Border Object box and use these settings:
i. Population = Nuclei
ii. Method = Common Filters
iii. Output Population = Nuclei NB
Press the “+” button again and click calculate intensity properties.
Use these settings:
i. Channel = CyQuant Green
ii. Population = Nuclei NB
iii. Region = Nucleus
iv. Method = Standard
v. Property Prefix = Intensity Nucleus NB
Press the “+” button again and click calculate morphology properties, select all boxes, and use these settings:
i. Population = Nuclei NB
ii. Region = Nucleus
iii. Method = Standard
iv. Property Prefix = Nucleus NB
Press the “+” button again, click select population, and use these settings:
i. Population = Nuclei NB
ii. Method = Filter by Property
iii. Intensity Nucleus NB Mean > 2,000
iv. Nucleus NB Area [μm2] < 70
v. Output Population = Nuclei NB Leukemia Viable
Press the “+” button again, click select population, and use these settings:
i. Population = Nuclei NB
ii. Method = Filter by Property
iii. Nucleus NB ≥ 75
iv. Output population = Nuclei NB MSC
Press the “+” button again, click define results, and use these settings:
i. Under Population: Nuclei NB Leukemia Viable select the number of objects box
ii. Apply to All = Individual selection
iii. Intensity Nucleus = Mean
iv. Intensity Area [μm2] = Mean
v. Under Population: Nuclei NB MSC select the number of objects box
vi. Apply to all = Individual selection
vii. Intensity Nucleus = Mean
Click the save button and name the pipeline accordingly.
Click the Evaluation tab in the Harmony software.
Check that you have the correct analysis pipeline and measurement files.
Select all the wells and fields of view that you want to analyze.
Click the Start button to analyze the plate.
Data analysis procedure for AOPI
Verify that the correct assay layout is attached to your plate.
Click the Image Analysis tab in the Harmony software.
Click the button to the right of Analysis and load the analysis pipeline you would like to use or, if setting up a new pipeline, press the new button under measurement.
For setting a new pipeline:
Press the “+” button at the bottom of the tab to add a building block to your pipeline.
Click Find Nuclei.
Under Find Nuclei, select the channel you would like to use and the method that best labels the cells in your image.
Label the output population as AO Nuclei.
Press the “+” button again and click select population.
Select the remove border object box and use these settings:
i. Population = AO Nuclei
ii. Method = Common Filters
iii. Output Population = AO Nuclei NB
Press the “+” button again and click calculate intensity properties.
Use these settings:
i. Channel = Brightfield
ii. Population = AO Nuclei NB
iii. Region = Nucleus
iv. Method = Standard
v. Property Prefix = AO Intensity
Press the “+” button again and click calculate morphology properties, select area box, and use these settings:
i. Population = AO Nuclei NB
ii. Region = Nucleus
iii. Method = Standard
iv. Property Prefix = AO Nucleus
Press the “+” button again, click select population, and use these settings:
i. Population = AO Nuclei NB
ii. Method = Filter by Property
iii. AO Intensity Mean > 200
iv. AO Intensity Mean < 100
v. Output Population = AO Viable Nuclei
Press the “+” button again, click find nuclei, and use these settings:
i. Channel = Brightfield
ii. ROI = None
iii. Method = C
iv. Output population – PI Nuclei
Press the “+” button again, click select population, and use these settings:
i. Population = PI Nuclei
ii. Method = Common Filters
iii. Click the remove border object box
iv. Output population = PI Nuclei NB
Press the “+” button again and click calculate intensity properties.
Use these settings:
i. Channel = Propidium Iodide
ii. Population = PI Nuclei NB
iii. Region = Nucleus
iv. Method = Standard
v. Property Prefix = PI Intensity
Press the “+” button again and click calculate morphology properties, select the area box, and use these settings:
i. Population = PI Nuclei NB
ii. Region = Nucleus
iii. Method = Standard
iv. Property Prefix = PI Nucleus
Press the “+” button again, click select population, and use these settings:
i. Population = PI Nuclei NB
ii. Method = Filter by Property
iii. PI Intensity Mean > 1,000
iv. PI Nucleus Area [μm2] < 100
v. Output Population = PI Dead Nuclei
Press the “+” button again, click define results, and use these settings:
i. Under Population: AO Viable Nuclei select the number of objects box
ii. Apply to All = Individual selection
iii. AO Intensity = Mean
iv. AO Nucleus Area [μm2] = Mean
v. Under Population: PI Dead Nuclei select the number of objects box
vi. Apply to all = Individual selection
vii. PI Intensity = Mean
viii. Method = Formula Output
ix. Formula = [a/(a+b)]*100
x. Variable a = AO Viable Nuclei – Number of Objects
xi. Variable b = PI Dead Nuclei – Number of Objects
xii. Output name = Viability %
Click the save button and name the pipeline accordingly.
Click the Evaluation tab in the Harmony software.
Check that you have the correct analysis pipeline and measurement files.
Select all the wells and fields of view that you want to analyze.
Click the Start button to analyze the plate.
Data evaluation
Click the Evaluation tab in the Harmony software.
Load the correct evaluation using the button next to the Evaluation line.
The evaluation results will populate in the middle of the screen.
Data from the CyQuant Green wells will populate as number of viable leukemia cells per well.
Data from the AOPI well will populate as Viability % per well and can be used as an internal control for the sample.
Copy data into a Microsoft Excel analysis template to calculate the relative viabilities of each drug treatment compared to the average of the untreated control wells. Dose response curves are generated from the raw data using software of choice.
Validation of protocol
Our group has measured and reported ex vivo drug sensitivity of eight drugs used in the current method (i.e., L-asparaginase, daunorubicin, bortezomib, mercaptopurine, nelarabine, prednisolone, vincristine, and vorinostat) via an MTT-based method as previously described (Holleman et al., 2004; Lugthart et al., 2005; Autry et al., 2020). Likewise, our group has measured and reported ex vivo drug sensitivity of eight drugs used in the current method (i.e., CHZ868, dasatinib, gilteritinib, ibrutinib, inotuzumab ozogamicin, palbociclib, trametinib, and venetoclax) via a flow cytometry–based method as previously described (Gocho et al., 2021; Lee et al., 2023). To validate the current imaging-based method for these 16 drugs, a minimum of 15 patient samples (either primary or obtained from patient-derived xenograft models) were studied in parallel by the current imaging assay and either the MTT-based or flow cytometry–based method (Figure 5). Cell viability was calculated using each method. For each individual concentration data point, percent cell viability was charted and R2 values were generated. Correlation of the cell viability results was reproducibly achieved, with all R2 values above 0.5 (except for two drugs: 6-mercaptopurine, which has historically been tested via an MTT-based method, for which the observed R2 value was 0.4780, and CHZ868, which has historically been tested via a flow cytometry–based method, for which the observed R2 value was 0.4412).
Figure 5. Comparison of imaging assay with previous methods. (A–H) Correlation plots showing the correlation between the imaging method and the gold standard flow cytometry–based mesenchymal stromal cell (MSC) co-culture method. (I–P) Correlation plots showing the correlation between the imaging method and the gold standard MTT-based metabolic activity method. Leukemia cells were treated for 96 h in the presence of increasing concentrations of each agent and cell viability was determined. For each individual plot, viability determined by the current imaging assay is shown on the x-axis, and viability determined by the gold-standard assay method is shown on the y-axis.
In addition to the 16 drugs for which we already had a validated method established, we introduced ex vivo sensitivity testing of an additional 24 drugs with the current imaging-based method. We determined drug sensitivity of MSCs alone to each single agent after a 96-hour incubation for all 40 drugs used with the proposed assay (Figure S1). Optimal concentration ranges were chosen based on clinically achievable concentrations in vivo and on the frequency distribution of LC50 values for each drug.
We measured ex vivo drug sensitivity to our drug panel of 40 antineoplastic agents with the current imaging-based method in 30 primary adult and pediatric ALL samples. Samples were assayed as received and reflect the variability of the population to be studied with this method. We observed varying patterns of drug sensitivity across patients (Figure 6). Leukemia cells with certain molecular markers had a high probability of showing a predictable sensitivity pattern (e.g., samples positive for the ETV6::RUNX1 fusion are usually sensitive to asparaginase; samples positive for the TCF3::PBX1 fusion are usually sensitive to dasatinib) (Frismantas et al., 2017; Lee et al., 2023). In this pilot dataset, we observed a lower median asparaginase LC50 value in cases with the ETV6::RUNX1 fusion (n = 4) vs. other cases (p = 0.02; Figure 6B), and a lower median dasatinib LC50 value in cases with the TCF3::PBX1 fusion (n = 3) vs. other cases (p = 0.009; Figure 6C).
Figure 6. Validation of imaging assay with primary patient samples. (A) Heatmap indicates the sensitivity of 30 primary leukemia samples to 40 anti-leukemia agents. LC50 values for each drug were ranked, with red indicating most resistant cases and violet indicating most sensitive cases. Grey squares indicate no data. (B) LC50 values of asparaginase in 30 primary ALL cases displayed as cases with the ETV6::RUNX1 fusion (n = 4) vs. other cases (n = 26). Median LC50 value for each group is shown as a white circle (p = 0.02 by Wilcoxon rank-sum test). (C) LC50 values of dasatinib in 30 primary ALL cases displayed as cases with the TCF3::PBX1 fusion (n = 3) vs. other cases (n = 27). Median LC50 value for each group is shown as a white circle (p = 0.009 by Wilcoxon rank-sum test). LC50, the concentration required to kill 50% of cells.
General limitations and potential problems
This protocol has potential limitations. This method is designed for analyzing fresh primary samples, one at a time. If a laboratory adopting this protocol cannot receive fresh primary samples, adjustments would potentially need to be made to the method. The imager used in this protocol may not be easily acquired by every institute. While this protocol may be adapted for other high-throughput imaging platforms, this adaptation would need to be validated beforehand. The ability to separate two cell populations is essential and is not exclusive to the imaging platform. There are options for free open-source image analysis software that could be used to analyze images taken from other platforms, which have the ability to perform batch analysis.
Due to the heterogeneity of primary samples, occasionally, samples may contain visible clumps upon receipt. If this occurs, samples can be strained using a 40 μm cell strainer to remove cell clumps before processing. Finally, due to the amount of manual pipetting, either with a single-channel or multichannel pipette, there is potential for variability among samples, and great care needs to be taken to minimize error.
Acknowledgments
This work was supported by the American Lebanese Syrian Associated Charities; NIH grants P30 CA021765 (SJCRH Cancer Center Support Grant) and P50 GM115279 (to MVR, CGM, WEE, and JJY); and Hyundai Hope on Wheels (to SEK). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. CGM is the William E. Evans Endowed Chair and JJY is the Endowed Chair of Pharmacogenomics at SJCRH.
Competing interests
All authors declare no competing interests.
References
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Wudhikarn, K., King, A. C., Geyer, M. B., Roshal, M., Bernal, Y., Gyurkocza, B., Perales, M.-A. and Park, J. H. (2022). Outcomes of relapsed B-cell acute lymphoblastic leukemia after sequential treatment with blinatumomab and inotuzumab. Blood Adv 6(5): 1432-1443.
Supplementary information
The following supporting information can be downloaded here:
Supplemental Methods
Figure S1. MSC Drug Sensitivity Curves for the 40 Therapeutic Agents Tested
Article Information
Copyright
© 2023 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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Peer-reviewed
Fluorescent PCR–based Screening Methods for Precise Knock-in of Small DNA Fragments and Point Mutations in Zebrafish
BC Blake Carrington
RS Raman Sood
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4732 Views: 619
Reviewed by: Alberto RissoneQin TangGülçin ÇAKAN AKDOĞAN
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Original Research Article:
The authors used this protocol in BMC Genomics Dec 2022
Abstract
Generation of zebrafish (Danio rerio) models with targeted insertion of epitope tags and point mutations is highly desirable for functional genomics and disease modeling studies. Currently, CRISPR/Cas9-mediated knock-in is the method of choice for insertion of exogeneous sequences by providing a repair template for homology-directed repair (HDR). A major hurdle in generating knock-in models is the labor and cost involved in screening of injected fish to identify the precise knock-in events due to low efficiency of the HDR pathway in zebrafish. Thus, we developed fluorescent PCR–based high-throughput screening methods for precise knock-in of epitope tags and point mutations in zebrafish. Here, we provide a step-by-step guide that describes selection of an active sgRNA near the intended knock-in site, design of single-stranded oligonucleotide (ssODN) templates for HDR, quick validation of somatic knock-in using injected embryos, and screening for germline transmission of precise knock-in events to establish stable lines. Our screening method relies on the size-based separation of all fragments in an amplicon by fluorescent PCR and capillary electrophoresis, thus providing a robust and cost-effective strategy. Although we present the use of this protocol for insertion of epitope tags and point mutations, it can be used for insertion of any small DNA fragments (e.g., LoxP sites, in-frame codons). Furthermore, the screening strategy described here can be used to screen for precise knock-in of small DNA sequences in any model system, as PCR amplification of the target region is its only requirement.
Key features
• This protocol expands the use of fluorescent PCR and CRISPR-STAT for screening of precise knock-in of small insertions and point mutations in zebrafish.
• Allows validation of selected sgRNA and HDR template within two weeks by somatic knock-in screening.
• Allows robust screening of point mutations by combining restriction digest with CRISPR-STAT.
Graphical overview
Overview of the three-phase knock-in pipeline in zebrafish (created with BioRender.com)
Keywords: Zebrafish CRISPR/Cas9 Knock-in Epitope tags Point mutations ssODN ssDNA CRISPR-STAT Screening method
Background
The use of zebrafish for functional genomics and disease modeling has gained popularity in recent years due to the availability of CRISPR/Cas9-based methods for generating desired gene knockout and knock-in models. These methods are based on the introduction of a double-strand break (DSB) at a desired site in the genome by CRISPR/Cas9 followed by non-homologous end joining (NHEJ)-mediated indels causing gene knockouts or homology-directed repair (HDR)-mediated knock-in of exogenous sequences by providing a donor DNA template. Several studies have shown that single-stranded oligonucleotides (ssODNs) with asymmetric homology arms based on Richardson et al., (2016) can be used to generate fish with targeted knock-in of in-frame epitope tags for proteomics analysis, LoxP sites for conditional knockout, and stop codons or patient-specific point mutations for in vivo disease modeling (Burg et al., 2018; Prykhozhij et al., 2018; de Vrieze et al., 2021; Sieliwonczyk et al., 2023). However, this is a challenging process due to the low efficiency of HDR in zebrafish, requiring extensive founder screening by either labor-intensive gel-based methods—such as allele-specific PCR and restriction fragment length polymorphism, cloning, and sequencing—or highly sophisticated computational analysis of next generation sequencing data (Bedell et al., 2012; Armstrong et al., 2016; Prykhozhij et al., 2018; Zhang et al., 2018; de Vrieze et al., 2021; Sieliwonczyk et al., 2023). Thus, easy, scalable, and robust screening methods are required to identify the rare precise knock-in events from the common NHEJ-mediated indels or recombination errors.
Here, we describe our recently demonstrated fluorescent PCR–based screening methods for successful generation of zebrafish knock-in models with epitope tags, point mutations, or any other small insertions using ssODNs (Carrington et al., 2022). Fluorescent PCR is highly sensitive and allows accurate sizing of all fragments in a PCR product even from mosaic samples using CRISPR-STAT (Carrington et al., 2015). For insertion of epitope tags or any other sequences of known size, we can distinguish knock-in alleles from the wildtype (WT) or CRISPR/Cas9-induced random indels by size of the expected peak. For knock-in of point mutations, we combined fluorescent PCR with a restriction digest so that exact sizes of digested products with or without the knock-in can be determined. The restriction site can be introduced in the repair template by the desired point mutation, change of the PAM site, or a silent change. Our screening method cannot be used if the same restriction site is also present in the amplicon upstream of the intended point mutation, or a restriction site cannot be introduced by the above methods. Overall, our screening methods allow the user to determine within a week if knock-in is occurring in injected embryos or if the design of sgRNA/repair template needs to be reevaluated. Our methods are easy to implement as many zebrafish researchers are already using fluorescent PCR for genotyping and CRISPR-STAT for sgRNA evaluations (Varshney et al., 2016; Ramanagoudr-Bhojappa et al., 2018; Hoshijima et al., 2019; Pillay et al., 2022). This method can also be adopted by researchers using other model systems, as it is a PCR-based screening method.
Materials and reagents
Biological materials
Zebrafish (Danio rerio): zebrafish strains can be obtained from collaborators or repositories, such as the Zebrafish International Resource Center (https://zebrafish.org/home/guide.php) or the European Zebrafish Resource Center (https://www.ezrc.kit.edu/).
General reagents
Instant ocean aquarium salt (Petco, catalog number: 77763)
Reverse osmosis water
TE, pH 8.0 (Quality Biological, catalog number: 351-011-131)
Ultrapure water (KD medical, catalog number: RGF-3410)
Project-specific restriction enzymes and associated buffers (New England Biolabs)
Primers and HDR template
M13F-FAM Primer-/56-FAM/TGTAAAACGACGGCCAGT (IDT)
Note: Primer should be resuspended to 100 μM using TE, pH 8.0.
Project-specific primers for fluorescent PCR designed as described in section A1, step 2: forward M13F tailed primer and reverse PIG-tailed primer (IDT)
Note: Primers should be resuspended to 100 μM using TE, pH 8.0.
Project-specific primers for fluorescent knock-in screening designed as described in section A4, step 2: knock-in screening forward M13F tailed primer and knock-in screening reverse PIG-tailed primer (IDT)
Note: Primers should be resuspended to 100 μM using TE, pH 8.0.
Project-specific top strand oligos (IDT) designed as described in section A1, step 1c.
Note: Top strand oligos should be resuspended to 100 μM using TE, pH 8.0. After resuspension, store indefinitely at -20 °C.
Universal bottom strand ultramer (5′-AAAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGC CTTATTTTAACTTGCTATTTCTAGCTCTAAAAC-3′) (IDT)
Critical: Aliquots should not go through multiple freeze-thaw cycles. Resuspension in water is not recommended.
Note: Bottom strand ultramer should be resuspended to 100 μM using TE, pH 8.0. After resuspension, store indefinitely in 10–15 μL aliquots at -80 °C.
Project-specific HDR templates ordered as ultramers (IDT) or ssDNA (Azenta Life Sciences)
sgRNA and Cas9 synthesis reagents
Phusion High Fidelity DNA Polymerase and associated 5× buffer (NEB, catalog number: M0530S)
Advantage UltraPure PCR dNTP mix (10 mM each) (Takara, catalog number: 639125)
HiScribe T7 Quick High Yield RNA Synthesis kit (NEB, catalog number: E2050S)
RNA Clean and Concentrator-5 kit (Zymo Research, catalog number: R1016) Critical: Other RNA purification kits may not work as well due to the small size of the RNA being purified.
Ethyl alcohol, pure (EtOH) (Sigma, catalog number: 459844)
pT3TS-nls-zCas9-nls plasmid (Addgene, catalog number: 46757)
XbaI and CutSmart buffer (New England Biolabs, catalog number: R0145S)
QIAquick PCR Purification kit (Qiagen, catalog number: 28104)
mMessage mMachine T3 Transcription kit (Thermo Fisher, catalog number: AM1348)
Gel electrophoresis reagents
Note: Equivalent reagents can be used as substitutes.
GeneRuler 50 bp DNA ladder (Thermo Fisher, catalog number: SM0371)
GeneRuler 1 kb DNA ladder (Thermo Fisher, catalog number: SM0311)
5× DNA gel loading solution (Quality Biological, catalog number: 351-028-661)
10× TAE buffer (Quality Biological, catalog number: 351-009-131)
UltraPure agarose (Thermo Fisher, catalog number: 16500500)
UltraPure ethidium bromide (Thermo Fisher, catalog number: 15585-011)
Caution: Carcinogenic. Wear gloves when handling.
DNA extraction reagents
Extraction solution (Sigma, catalog number: E7526)
Tissue preparation solution (Sigma, catalog number: T3073)
Neutralization solution B (Sigma, catalog number: N3910)
PCR reagents
AmpliTaq Gold DNA Polymerase with Gold Buffer and MgCl2 (Thermo Fisher, catalog number: 4311820)
GeneAmp dNTP blend (100 mM) (Thermo Fisher, catalog number: N8080261)
Capillary electrophoresis reagents
GeneScan 400HD ROX dye size standard (Thermo Fisher, catalog number: 402985)
Hi-Di formamide (Thermo Fisher, catalog number: 4440753)
Caution: Carcinogenic and target organ and reproductive toxicity. Wear gloves when handling.
Note: Store at -20 °C until ready to use. Avoid freeze thawing. Once thawed, store at 4 °C and use contents within four weeks.
POP-7 polymer for 3730/3730xl DNA Analyzer (Thermo Fisher, catalog number: 4335615)
Note: Store at 4 °C. Do not freeze.
3730 running buffer (10×) (Thermo Fisher, catalog number: 4335613)
Note: Cheaper alternative from VWR (catalog number: 97063-502) also works well.
TOPO cloning and sequencing reagents
MinElute Reaction Cleanup kit (Qiagen, catalog number: 28204)
TOPO TA Cloning kit for Sequencing, with One Shot TOP10 Electrocomp E. Coli (Thermo Fisher, catalog number: K458001)
LB agar plates with 100 μg/mL ampicillin (Thermo Fisher, catalog number: J63197)
Note: These can be prepared in-house using standard techniques.
ExoSAP-IT PCR Product Cleanup (Thermo Fisher, catalog number: 78201)
BigDye Terminator v3.1 Cycle Sequencing kit (Thermo Fisher, catalog number: 4337455)
DyeEx 96 kit (Qiagen, catalog number: 63183)
Solutions
Egg water (see Recipes)
Fluorescent PCR primer mix (see Recipes)
Fluorescent PCR master mix (see Recipes)
Recipes
Egg water
Reagent Final concentration Amount
Instant ocean aquarium salt n/a 1.92 g
Reverse osmosis water n/a 32 L
Total n/a 32 L
Store solution at room temperature (RT) for up to six months.
Fluorescent PCR primer mix
Reagent Final concentration Amount
TE, pH 8.0 n/a 485 μL
Forward M13F tailed primer (100 μM) 1 μM 5 μL
Reverse PIG-tailed primer (100 μM) 1 μM 5 μL
M13F-FAM primer (100 μM) 1 μM 5 μL
Total n/a 500 μL
Store primer mix at 4 °C in a covered box to avoid exposure to light.
Fluorescent knock-in screening primer mix
Reagent Final concentration Amount
TE, pH 8.0 n/a 485 μL
Knock-in screening forward M13F tailed primer (100 μM) 1 μM 5 μL
Knock-in screening reverse PIG-tailed primer (100 μM) 1 μM 5 μL
M13F-FAM primer (100 μM) 1 μM 5 μL
Total n/a 500 μL
Store primer mix at 4 °C in a covered box to avoid exposure to light.
Fluorescent PCR master mix
Reagent Final concentration Amount
10× Gold Buffer 1.5× 7.94 mL
25 mM MgCl2 solution 3.97 mM 7.94 mL
GeneAmp 100 mM dNTP blend 1.59 mM 795 μL
AmpliTaq Gold DNA Polymerase 1.25 U 635 μL
Ultrapure water n/a 32.69 mL
Total n/a 50 mL
Gold Buffer and MgCl2 come with the AmpliTaq Gold DNA Polymerase. Aliquot PCR master mix into 500 μL and 1 mL aliquots and store indefinitely at -20 °C.
Laboratory supplies
Note: Any equivalent supplies can be used.
Microcentrifuge tubes, 1.7 mL (VWR, GeneMate, catalog number: 490003-230)
Pipettes [Rainin, catalog numbers: 17014393 (0.1–2 μL), 17014388 (0.5–10 μL), 17014392 (2–20 μL), 17014391 (20–200 μL), 17014382 (100–1,000 μL), 17013808 (2–20 μL multichannel)]
Needle loading tips (Eppendorf, catalog number: 930001007)
Glass capillaries (World Precision Instruments, catalog number: TW100F-4)
96-well PCR plates (SSIbio, catalog number: 3400-00)
Plate seals (Azenta, catalog number: 4ti00500)
96-well semi-skirted plates for 3730xl DNA analyzer (SSIbio, catalog number: 3450-00)
Equipment
Note: Any equivalent device can be used.
Microcentrifuge (MidSci, catalog number: C2400)
Vortex (Diagger, catalog number: SI-0236)
Block heater at 37 °C (Thermo Fisher, catalog number: 13687720)
Owl D2 wide-gel electrophoresis system (Thermo Fisher, catalog number: D2)
NanoDrop spectrophotometer 1000 (Thermo Fisher)
PCR plate spinner (VWR, catalog number: 89184-608)
Veriti 96-well thermal cycler 0.2 mL (Thermo Fisher, catalog number: 4479071)
Microinjection apparatus (World Precision Instruments)
Needle puller (Kopf Instruments model 710)
-20 °C freezer (VWR, catalog number: 10819-850)
-80 °C freezer (Thermo Fisher, catalog number: TSX40086A)
3730xl DNA Analyzer equipped with a 36 cm capillary (Thermo Fisher, catalog number: A41046)
Incubator at 28.5 °C (Thermo Fisher, catalog number: 51028068)
Incubator at 37 °C (Thermo Fisher, catalog number: 50125590)
Software and datasets
UCSC genome browser (Zebrafish Assembly GRCz11/danRer11) https://genome.ucsc.edu/cgi-bin/hgGateway
Sequencher version 5.3 (Gene Codes)
Note: Any equivalent sequence analysis software can be used.
3730xl Data Collection Software 5 (Thermo Fisher).
Note: CRISPR-STAT module settings can be found in Varshney et al. (2016).
GeneMapper Software version 6 (Thermo Fisher, catalog number: 4475073)
Note: Panel manager and analysis method editor settings can be found in Varshney et al. (2016).
Procedure
Design phase
A1. sgRNA/Cas9 design and synthesis
sgRNA target selection
Access the “ZebrafishGenomics” (Varshney et al., 2016) and/or “CRISPRscan” (Moreno-Mateos et al., 2015) tracks on the UCSC Genome Browser (Figure 1).
Select ~30 bp of sequence flanking each site of the intended integration site (~60 bp total sequence) to search for target sequences. Select 2–3 targets that generate the DSB as close to the integration site as possible and obtain their CRISPR target sequences (Figure 1).
Note: See General notes for information about alternate sgRNA design.
Figure 1. Screenshot of the UCSC Genome Browser with CRISPRscan and ZebrafishGenomics hubs enabled to identify sgRNA target sites. Use tracks (marked in yellow) to search for targets that can be synthesized with a T7 promoter (guide sequence begins with a “GG”) as close as possible to the intended nucleotide for point mutation or site of insertion. In this example, both CRISPRscan and ZebrafishGenomics predicted two overlapping targets (sgRNA-T1 and sgRNA-T2) close to the target nucleotide (marked in red box) and were selected for sgRNA synthesis.
Add the T7 promoter sequence (TAATACGACTCACTATA) to the 5′ side of the target sequence (obtained from the genome browser) and the gRNA overlapping sequence (GTTTTAGAGCTAGAAATAGC) to the 3′ side of the target sequence to obtain the final sequence of the top strand oligo. The oligo sequence for a 20 bp sgRNA will be as follows:
5′-TAATACGACTCACTATAggn(18)GTTTTAGAGCTAGAAATAGC-3′. Critical: Do not include the PAM sequence in the oligo.
Order the top strand oligos as standard desalted oligos and universal bottom strand oligo as an ultramer (see Reagents).
Fluorescent PCR primer design
Design primers to amplify a 200–350 bp product surrounding the target region with the sgRNA binding sites approximately in the middle of the amplicon. Critical: Target sequences should be a minimum of 25 bp from the ends of the amplicon to prevent NHEJ from removing any of the primer binding sequence. If possible, design primers to avoid repetitive sequences or stretches of the same nucleotide, as it is important that the PCR from a WT sample gives a single peak to avoid false positive data. A primer design software can also be used.
Add the M13F sequence (5′-TGTAAAACGACGGCCAGT -3′) to the 5′ side of the forward primer and a PIG-tail sequence (5′-GTGTCTT-3′) to the 5′ side of the reverse primer. Tailed primers will be as follows:
Forward: 5′-TGTAAAACGACGGCCAGT (n)18–22 -3′
Reverse: 5′-GTGTCTT (n)18–22 -3′
Order tailed primers and the M13-FAM primer (see Reagents) as standard desalted primers. Critical: It is recommended that each primer set be run with a minimum of eight WT DNA samples to ensure the amplicon produces a single peak (see section A3 for PCR setup).
sgRNA synthesis
Set up the oligo annealing reactions as follows:
5× Phusion buffer 10 μL
Advantage UltraPure PCR dNTP mix (10 mM each) 1 μL
Top-strand oligo (10 μM) 1 μL
Bottom strand ultramer (10 μM) 1 μL
Phusion high fidelity DNA polymerase 0.5 μL
Ultrapure water 36.5 μL
Total 50 μL
Anneal and extend the oligos in a thermocycler using the following conditions:
98 °C, 2 min
50 °C, 10 min
72 °C, 10 min
4 °C, hold
Run 4 μL of the assembled oligo reaction along with a 50 bp DNA ladder on a 2.5% (w/v) agarose gel using standard conditions. A single band is expected at 120 bp (Figure 2).
Following gel validation, set up the sgRNA synthesis reaction using the HiScribe T7 Quick High Yield RNA Synthesis kit as follows:
NTP buffer mix 5 μL
Assembled oligo 4 μL
T7 RNA polymerase 1 μL
Total 10 μL
Incubate the reaction for 4 h at 37 °C followed by a 4 °C hold.
Add 1 μL of DNAse (included with HiScribe T7 Quick High Yield RNA Synthesis kit) to the reaction and incubate for 20 min at 37 °C.
Purify the RNA synthesis reaction using the RNA Clean and Concentrator-5 kit according to the manufacturer’s instructions and elute with 20 μL of ultrapure water.
Quantify the sgRNA using a NanoDrop and run ~300 ng along with a 50 bp DNA ladder on a 2.5% (w/v) agarose gel using standard conditions to confirm correct synthesis. A band between 100 and 200 bp is expected (Figure 2). Occasionally, a slightly larger band is also seen due to secondary structure. Dilute the remaining sgRNA to a final concentration between 100 and 300 ng/μL with an A260/280 ratio of 2.0–2.3. Pause point: sgRNA can be stored at -80 °C for several months.
Figure 2. Agarose gel electrophoresis showing products of oligo assembly and sgRNA synthesis reactions. Lane M: marker (50 bp ladder); lane 1: correctly assembled oligo runs at ~120 bp; lane 2: successful sgRNA synthesis product runs between 100 and 200 bp.
SpCas9 mRNA synthesis
Note: See General notes for information about alternate Cas options.
Linearize 5 μg of the pT3Ts-nls-zCas9-nls plasmid by digesting with XbaI in a total volume of 50 μL at 37 °C for 2 h.
Purify the linearized DNA with the QIAquick PCR Purification kit according to the manufacturer’s instructions.
Verify digestion on a 1% (w/v) agarose gel using standard protocols. A single band is expected at 7.3 kb. Example of a gel with expected band can be found in Varshney et al. (2016).
Quantify the digested DNA with a NanoDrop.
Set up the in vitro transcription reaction using the mMessage mMachine T3 Transcription kit:
2× NTP/CAP 10 μL
10× reaction buffer 2 μL
Linearized DNA template 1 μg
Enzyme mix 2 μL
Ultrapure water up to 20 μL
Total 20 μL
Incubate at 37 °C for 2 h.
Add 1 μL of TURBO DNase (included in the mMessage mMachine T3 Transcription kit) and incubate for 15 min and 37 °C.
Purify the RNA transcription reaction using LiCl (included in the mMessage mMachine T3 Transcription kit) according to the manufacturer’s instructions. Resuspend the RNA pellet with 20 μL of ultrapure water.
Quantify the mRNA using a NanoDrop. The concentration should be > 500 ng/μL with an A260/280 ratio of 2.0–2.3.
Run ~500 ng on an 1% (w/v) agarose gel to confirm quality using standard protocols. The expected band will run at ~1.6 kb.
Aliquot the Cas9 mRNA into 5 μL aliquots with a concentration of 500–1,000 ng/μL. Pause point: Cas9 aliquots can be stored at -80 °C for months. Multiple freeze-thaw cycles are not recommended.
A2. Injections of sgRNA/Cas9 mRNA for evaluation of sgRNA activity
Prepare injection apparatus and pull needles.
Prepare an injection mixture for each sgRNA from section A1, step 3 with SpCas9 mRNA from section A1, step 4 as follows:
sgRNA (354 ng) variable volume
Cas9 mRNA (2.1 μg) variable volume
Ultrapure water up to 10 μL
Total 10 μL
Load the injection mixture into a capillary injection needle and inject 1.4 nL per embryo into ~100 embryos. Repeat this step for each injection mixture.
Note: If a different volume is injected, adjust amounts of sgRNA and Cas9 mRNA in injection mixture such that each embryo receives 50 pg sgRNA and 300 pg Cas9 mRNA.
Incubate the injected embryos as well as uninjected controls from the same clutch in a 28.5 °C incubator in egg water. Remove any unfertilized and injected damaged embryos at 3–6 h post fertilization (hpf).
At 1 day post fertilization (dpf), remove any deformed and dead embryos.
A3. Determine activity of each sgRNA using CRISPR-STAT
At 1 dpf, collect and euthanize eight embryos from each injection group and uninjected WT as controls.
Place one embryo per well in a 96-well plate. Each injection group will use one column on the plate. Critical: If embryos are still in chorions, place the plate into a -20 or -80 °C freezer for 20–30 min to completely freeze and burst the chorion, allowing extraction reagents in following steps to access the tissue.
Mix 25 μL of extraction solution and 7 μL of tissue preparation solution for each embryo (i.e., 10 samples: 250 μL extraction solution and 70 μL tissue preparation solution). Add 32 μL of the extraction/tissue preparation solution mixture to each sample. Critical: Embryos should be submerged in the solution. Do not vortex.
Incubate the plate at RT for 10 min followed by 5 min at 95 °C.
Add 25 μL of neutralization solution B to each well and mix by vortexing. Pause point: DNA can be stored at -20 °C for up to four weeks.
Note: See General notes for information about alternate DNA extraction methods.
Dilute the extracted DNA 1:10 with ultrapure water for use in PCR reactions.
Add 30 μL of fluorescent PCR primer mix (see Recipes) to a 500 μL aliquot of fluorescent PCR master mix (see Recipes) and aliquot 10 μL/well of a new 96-well plate.
Note: These volumes are for 48 samples and can be adjusted depending on the number of samples being processed.
Add 3 μL of diluted DNA from section A3, step 6 to each well. Vortex the plate and spin down.
Run the fluorescent PCR reaction with the following conditions:
94 °C, 12 min
40 cycles
94 °C, 30 s
57 °C, 30 s
72 °C, 30 s
72 °C, 10 min
4 °C, hold
Pause point: Fluorescent PCR products can be stored at -20 °C for up to one month.
Make 1:50 dilution of GeneScan 400HD ROX using Hi-Di formamide.
Add 10 μL of the ROX/Hi-Di mixture to a plate that will fit onto a 3730xl DNA analyzer.
Add 3 μL of the fluorescent PCR product to the plate containing the ROX/Hi-Di mixture. Vortex the plate and spin down.
Incubate plate at 95 °C for 5 min using a thermocycler.
Load the plate into the 3730xl DNA analyzer and run using CRISPR-STAT settings.
Analyze data using the GeneMapper Software package to determine sgRNA activity and select the best target (Figure 3).
Note: Active sgRNAs show multiple peaks corresponding to the NHEJ-mediated indels. If all tested sgRNAs show good activity, choose the target that generates the DSB closest to the intended integration site.
Figure 3. Examples of CRISPR-STAT plots showing sgRNA activity. In this example, both sgRNAs selected in Figure 1 are active. sgRNA-T1 would be selected as the best target and used in subsequent steps, as it generates the DSB closer to the intended nucleotide change. The x-axis shows the size of the peaks (bp), the y-axis shows fluorescent intensity, and the green arrow denotes the WT allele.
A4. Design of HDR template and screening primers
HDR template design
Note: The design described below is for a sgRNA in the reverse orientation (CCN PAM site) with respect to the open reading frame (ORF). If the sgRNA is in the forward orientation (NGG PAM site), it is recommended to align the guide in the reverse compliment orientation and follow the steps below.
Align the selected sgRNA target sequence to the reference sequence in the orientation that displays PAM as CCN.
Determine the sequence of homology arms by identifying the site of DSB. Select 91 bp of the sequence on the left side of the DSB site for the left homology arm. Select 36 bp of sequence on the right side of the DSB site for the right homology arm.
Note: For SpCas9, the DSB occurs 5 bp upstream of the PAM site [i.e., N18(DSB)NN NGG].
For insertion of DNA fragments: insert the desired sequence between these homology arms (Figure 4). Critical: If the inserted sequence separates the sgRNA binding and PAM sites, modification of the PAM site is not required to prevent recutting after HDR (example in Figure 4A). Otherwise, make a silent change in the PAM site. If a silent change cannot be made to the PAM site, 3–4 silent changes in the sgRNA binding site are typically sufficient to disrupt binding and prevent recutting from the sgRNA (example in Figure 4B).
Note: Depending on where the DSB occurs, it might be necessary to add additional sequences to maintain the ORF (example in Figure 4C).
For point mutations: modify the desired nucleotide in the homology arm sequence. If the desired point mutation disrupts the PAM site and generates a restriction site, additional nucleotide changes are not required (example in Figure 5A). If the desired point mutation does not disrupt the PAM site, silent changes are required to either destroy the PAM site (example in Figure 5B) or sgRNA binding site (example in Figure 5C). Any combination of these bp changes (desired change, PAM site, and/or sgRNA binding site) will need to generate a new restriction site that can be used for screening. Critical: It is important to generate a new restriction site instead of destroying a restriction site. Using loss of a restriction site for screening can lead to false positives, as this site can be destroyed by random indels instead of the integration of the DNA fragment.
Figure 4. Examples of HDR template design to knock-in desired DNA sequences. (A) The DNA insertion of an in-frame codon TTA occurs directly at the DSB (denoted by red arrowhead). This insertion disrupts the sgRNA binding site and no additional modifications are required. (B) The DNA insertions occur 5 bp upstream of the DSB (denoted by red arrowhead). The insertion does not disrupt the sgRNA binding or the PAM site and a silent change cannot be made to the PAM site. Therefore, several silent changes (shown in magenta) are required to prevent recutting after HDR repair. (C) The HA epitope tag is inserted at the 3′ end of a gene’s ORF. The DSB (denoted by red arrowhead) occurs 6 bp upstream of the stop codon (shown in red font). To maintain the gene’s ORF, these 6 bp (shown in blue font) are added into the HDR template with one of the nucleotides (shown in magenta font) changed to a silent change to modify PAM site and prevent recutting after HDR repair. A new stop codon (shown in purple font) is added after the HA tag to terminate translation. In all examples, the sgRNA is in bold and the PAM site is underlined.
Figure 5. Examples of HDR template design to knock-in a point mutation. (A) The point mutation G>A (shown in blue font) modifies the PAM site as well as generates an AluI restriction site that can be used for screening. (B) The point mutation G>C (shown in blue font) is 17 bp downstream of the DSB site. A silent change is made to disrupt the PAM site (G>C, shown in magenta font) and this change also generates a SalI restriction site that can be used for screening. (C) The point mutation C>A (shown in blue font) is 6 bp upstream of the DSB site. In this example, a silent change cannot be made to the PAM site, and, therefore, four silent changes are required (shown in magenta font) to destroy the sgRNA binding site. One of these changes also generates a SfcI restriction site that can be used for screening. In all examples, the DSB site is denoted by a red arrowhead, sgRNA is shown in bold font, and the PAM site is underlined.
Identify potential polymorphisms in the homology arms by fin clipping 24 male and 24 female adult fish followed by PCR and sequencing of the target region.
Note: The previously designed primers may be used for this step, depending on their binding sites.
Analyze the sequence data to identify any polymorphisms. If polymorphisms are identified, fish should be grouped into a cohort with 100% homology to the reference sequence used for design of HDR template. Alternately, modify the homology arms to match the sequence of your fish cohort.
To obtain the final sequence of the HDR template, orient the sequence with the 91 bp homology arm on the 5′ side and the 36 bp homology arm on the 3′ side. Order templates smaller than 200 bp as an ultramer (IDT) and larger than 200 bp as a ssDNA fragment (Azenta Life Sciences).
Note: Depending on the vendor used to synthesize the HDR template, the amount delivered varies widely. The HDR template should be resuspended in ultrapure water to a concentration that can be accurately pipetted to prepare the injection mix in section B3.
Design of knock-in screening primers
For knock-in of DNA fragments: design primers that amplify the target region but do not overlap with the HDR template. The primers should amplify a region of up to 350 bp (including the insertion sequence). Add the M13F sequence (5′-TGTAAAACGACGGCCAGT-3′) to the 5′ side of the forward primer and a PIG-tail sequence (5′-GTGTCTT-3′) to the 5′ side of the reverse primer (Figure 6).
Note: The previously designed fluorescent PCR primers may be used for screening if they meet these criteria.
Figure 6. Schematic showing design of screening primers for insertion of a DNA fragment. Primers should not overlap with the HDR template to allow for screening of precise integration. Homology arms (left HA and right HA) are marked in red in both genomic DNA and HDR template. Inserted sequence is marked in blue on the HDR template. M13F sequence is added to the forward primer and the PIG-tail sequence is added to the reverse primer.
For point mutations: design primers that amplify the target region but do not overlap with the HDR template. The primers should amplify a region up to 350 bp. Add the M13F sequence (5′-TGTAAAACGACGGCCAGT-3′) to the 5′ side of the forward primer and a PIG-tail sequence (5′-GTGTCTT plus restriction site sequence-3′) to the 5′ side of the reverse primer (Figure 7). Critical: This screening strategy will not work if the same restriction site is also present between the new knock-in restriction site and the forward primer. The digested PCR fragment must be > 100 bp for the fluorescent signal to be detected.
Note: The previously designed forward primer may be used for screening if it meets these criteria. The reverse primer will need to have the new restriction site added between the PIG-tail and binding sequence. A schematic of the fluorescent PCR digestion strategy can be found in Carrington et al. (2022) (Figure 3B).
Figure 7. Schematic showing design of screening primers for knock-in of a point mutation. Primers should not overlap with the HDR template (shown with homology arms in red, restriction enzyme recognition sequence in blue, and point mutation by black line) to allow for screening of precise integration. M13F sequence is added to the 5′ end of forward primer. PIG-tail and the restriction enzyme recognition sequences (RS) are added to the 5′ end of reverse primer.
Somatic screening phase
B1. Injections to deliver sgRNA/Cas9 and HDR template
Prepare injection apparatus and pull needles.
Prepare injection mixture for sgRNA/Cas9 as described in section A2, step 2.
Prepare injection mixture for sgRNA/Cas9 plus HDR template as follows:
sgRNA (354 ng) variable volume
Cas9 mRNA (2.1 μg) variable volume
HDR template (175 ng) variable volume
Ultrapure water up to 10 μL
Total 10 μL
Note: These amounts are for injections of 1.4 nL per embryo. If a different volume is injected, adjust amounts of sgRNA, Cas9 mRNA, and HDR template in injection mixture such that each embryo receives 50 pg sgRNA, 300 pg Cas9 mRNA, and 25 pg of HDR template.
Load the injection mixture into a capillary injection needle and inject 1.4 nL per embryo. Repeat this step for each injection mixture. Inject ~100 embryos for the sgRNA/Cas9 mixture and ~200 embryos for the sgRNA/Cas9 plus HDR template mixture.
Incubate the injected embryos as well as uninjected controls in a 28.5 °C incubator in egg water. Remove any unfertilized and injected damaged embryos at 3–6 hpf.
At 1 dpf, remove any deformed or dead embryos.
B2. Evaluation of somatic knock-in using CRISPR-STAT
At 1 dpf, collect 24 embryos from each injection group (sgRNA/Cas9 and sgRNA/Cas9 plus HDR template) and uninjected controls in a 96-well plate (one embryo per well). Keep the remaining embryos in the incubator to be raised after knock-in is confirmed. Critical: If embryos are still in chorions, place the plate into a -20 or -80 °C freezer for 20–30 min to completely freeze and burst the chorion, allowing extraction reagents in following steps to access the tissue.
Extract DNA, perform fluorescent PCR using primers designed for knock-in screening (section A4, step 2), and run samples as described in section A3. For DNA insertion projects, proceed to step 7 and skip digestion steps 3–6 that are used for point mutation screening.
Note: Store extracted DNA at -20°C, as it will be needed for confirmation by TOPO cloning.
Prepare digestion mixture as follows:
Enzyme (200 U/μL) 15 μL
10× digestion buffer 75 μL
Ultrapure water 660 μL
Total 750 μL
Note: This is an example using SalI-HF that should be modified based on the enzyme that is appropriate for the designed HDR template.
Add 5 μL of the digestion mixture to a new 96-well plate.
Add 5 μL of the fluorescent PCR products to the plate containing the digestion mixture.
Incubate with the following conditions using a thermocycler:
37 °C, 1.5 h (digestion)
60 °C, 20 min (heat inactivation of restriction enzyme)
4 °C, hold
Critical: It is highly recommended that restriction digestion conditions are modified based on the restriction enzyme being used and optimized using WT samples before using injected samples. WT sample digestion is assessed by removal of the PIG-tail, resulting in a product that is 7–10 bp shorter than undigested PCR products (see Figure 8 panel B to compare undigested and digested samples to see the shift in WT peak size after digestion).
Mix 3 μL of fluorescent PCR product (DNA insertion) or digested fluorescent PCR product (point mutation) with 10 μL of ROX-Hi-Di mixture and run plates on 3730xl DNA analyzer as described in section A3, steps 10–14.
Analyze data using the GeneMapper Software package to determine the number of samples in each group that have peaks at the expected knock-in allele size (Figure 8).
Note: For point mutations, it is important to confirm complete digestion in the uninjected WT samples, since incomplete digestion can result in false negatives.
Figure 8. Examples of CRISPR-STAT plots from injected embryos to detect somatic knock-in. CRISPR-STAT plots from uninjected, sgRNA/Cas9-injected, and sgRNA/Cas9 plus HDR template–injected embryos are analyzed to determine if an enrichment of the peak corresponding to the knock-in allele size is seen in the presence of HDR template. The x-axis shows the size of the peaks (bp), the y-axis shows fluorescent intensity, and the red arrow denotes the expected knock-in allele. (A) Example of a DNA insertion with the knock-in peak being larger than the WT allele. (B) Example of a point mutation before and after digestion. The undigested samples serve as controls, with peak for WT allele at 7–10 bp larger than in the digested samples. The peak for knock-in allele should only be observed after digestion in sgRNA/Cas9 plus HDR template samples.
Compare the number of samples with knock-in peaks for each treatment and determine if there is an enrichment of the desired knock-in peak in the sgRNA/Cas9 plus HDR template samples compared to the sgRNA/Cas9 alone. If an enrichment is not observed, repeat injections to rule out technical errors or design a new sgRNA/HDR template combination.
B3. Somatic knock-in confirmation by TOPO cloning and sequencing
Select 2–3 samples that are positive for the expected size peak from the sgRNA/Cas9 plus HDR template group.
Note: Samples with a more robust peak at the expected knock-in size are preferred for this step.
Set up a PCR for each sample as follows:
10× Gold buffer 5 μL
25 mM MgCl2 3 μL
Advantage UltraPure PCR dNTP mix (10 mM each) 1 μL
Knock-in screening forward M13F tailed primer (20 μM) 3 μL
Knock-in screening reverse PIG-tailed primer (20 μM) 3 μL
AmpliTaq Gold DNA Polymerase 0.5 μL
Ultrapure water 31.5 μL
Diluted DNA (section B2, step 2) 3 μL
Total 50 μL
Run PCR with the following conditions:
94 °C, 12 min
35 cycles
94 °C, 30 s
57 °C, 30 s
72 °C, 30 s
72 °C, 10 min
4 °C, hold
Clean up PCR products with the MinElute Reaction Cleanup Kit following the manufacturer’s instructions and elute with 10 μL of ultrapure water.
Set up the TOPO cloning reaction using 4 μL of purified PCR product according to the manufacturer’s instructions. Critical: PCR product should be fresh.
Incubate reaction at RT for 1–2 h. Critical: Shorter incubation times may result in insufficient number of clones.
Transform 4 μL of the TOPO reaction using TOP10 cells according to manufacturer’s instructions. Plate 50 and 250 μL onto two separate agar plates containing ampicillin.
Incubate agar plates overnight at 37 °C.
Prepare PCR master mix for colony PCR as follows:
10× Gold buffer 250 μL
25 mM MgCl2 150 μL
Advantage UltraPure PCR dNTP mix (10 mM each) 50 μL
Knock-in screening forward M13F tailed primer (20 μM) 150 μL
Knock-in screening reverse PIG-tailed primer (20 μM) 150 μL
AmpliTaq Gold DNA Polymerase 25 μL
Ultrapure water 1,725 μL
Total 2,500 μL
Add 25 μL of the PCR master mix into each well of a 96-well plate.
Using clean pipette tips, touch the tip to individual clones followed by dipping the tip into a single well containing the PCR master mix. Repeat this to select 48 clones for each cloned sample. This allows for two cloned samples to be screened on a 96-well plate.
Run the colony PCR reaction with the following conditions:
94 °C, 12 min
35 cycles
94 °C, 30 s
57 °C, 30 s
72 °C, 30 s
72 °C, 10 min
4 °C, hold
Critical: If an alternative polymerase is used, the first denaturation should be a minimum of 10 min at 94 °C to lyse the bacteria.
Transfer 4 μL of the PCR product to a fresh 96-well plate and add 1 μL of ExoSAP-IT and 1 μL of ultrapure water to each reaction.
Using a thermocycler, run the following reaction:
37 °C, 15 min
80 °C, 15 min
4 °C, hold
Prepare BigDye Terminator master mix as follows:
Sequencing primer (5 μM) 100 μL
BigDye sequencing buffer (5×) 200 μL
BigDye terminator v3.1 100 μL
Total 400 μL
Note: Volumes are for 100 reactions and can be scaled as needed based on the number of samples to be processed. The sequencing primer is either the forward or reverse primer that was used for the colony PCR and is farthest away from the HDR template. The resulting sequence will need to cover both ends of the HDR template to confirm precise integration.
Add 4 μL of the BigDye Terminator master mix to each ExoSAP-IT-treated PCR reaction. Vortex the plate and spin down.
Run the BigDye Terminator reaction using the following conditions:
96 °C, 1 min
25 cycles
96 °C, 10 s
50 °C, 5 s
60 °C, 4 min
4 °C, hold
Clean up the BigDye Terminator reactions using DyeEx 96 kit plates according to the manufacturer’s instructions.
Run the purified products on a 3730xl DNA analyzer using standard sequencing programs.
Analyze clone sequences by aligning reads to reference sequence of the target region with the knock-in DNA fragment (see Data analysis section).
Grow remaining injected embryos or repeat injections to grow founder fish if precise knock-in is confirmed by cloning.
Germline screening phase
C1. Prioritization of founders to screen
Fin clip all adult founder fish and place each fin in a well of a 96-well plate.
Mix 50 μL of extraction solution and 12.5 μL of tissue preparation solution for each sample (i.e., 100 samples: 5.0 mL of extraction solution and 1.25 mL of tissue preparation solution).
Add 62.5 μL of the extraction/tissue preparation solution mixture to each sample. Critical: Fin tissue should be submerged in the solution. Do not vortex.
Incubate the plate at RT for 10 min followed by 5 min at 95 °C.
Add 50 μL of neutralization solution B to each well and mix by vortexing. Pause point: DNA can be stored at -20°C for up to four weeks.
Dilute DNA 1:10 with ultrapure water for use in PCR reactions.
Add 60 μL of fluorescent knock-in screening primer mix (see Recipes) to a 1 mL aliquot of fluorescent PCR master mix (see Recipes) and aliquot 10 μL/well of a new 96-well plate.
Note: These volumes are for 100 samples and can be adjusted depending on the number of samples being processed.
Using the multichannel pipette, transfer 3 μL of diluted DNA from step 6 to the plate with fluorescent knock-in screening primer mixture/fluorescent PCR master mix. Vortex the plate and spin down.
Run the fluorescent PCR reaction with the following conditions:
94 °C, 12 min
40 cycles
94 °C, 30 s
57 °C, 30 s
72 °C, 30 s
72 °C, 10 min
4 °C, hold
Pause point: Fluorescent PCR products can be stored at -20 °C for up to one month.
If screening for point mutations, perform restriction digest as described in section B2, steps 3–7.
Make 1:50 dilution of GeneScan 400HD ROX using Hi-Di formamide.
Add 10 μL of the ROX/Hi-Di mixture to a plate that will fit onto a 3730xl DNA analyzer.
Transfer 3 μL of the fluorescent PCR product from step 9 for DNA insertions and step 10 for point mutations to the plate containing the ROX/Hi-Di mixture. Vortex the plate and spin down.
Incubate plate at 95 °C for 5 min using a thermocycler.
Load the plate into the 3730xl DNA analyzer and run using CRISPR-STAT settings.
Analyze data using the GeneMapper Software package and identify founder fish that have an expected knock-in peak present in the fin clip. Pool these fish into a tank and use for screening (described below), as these fish are most likely to show germline transmission to their progeny. Founder fish that are negative for the knock-in peak in their fins should be kept separated until a positive germline transmitting fish is identified.
C2. Screening prioritized founder fish by pooled embryos
Set up pairwise crosses of each prioritized founder fish with a WT fish and collect embryos for screening.
Incubate embryos at 28.5 °C overnight.
At 1 dpf, collect three embryos per well (up to 24 wells per founder depending upon the clutch size) in a 96-well plate.
Note: This strategy allows for screening of 72 embryos per founder and four founders per plate.
Extract DNA, perform fluorescent PCR, and run samples as described in section C1, steps 2–15. For point mutations, perform restriction digest steps described in section B2, steps 3–7 after fluorescent PCR.
Analyze data using the GeneMapper software package to determine if any samples show the expected knock-in allele and identify the corresponding founder fish (Figure 9).
Figure 9. Examples of fluorescent PCR plots from pooled embryo screening to identify germline transmitting founders. Each embryo has a WT allele from the WT parent and an unknown allele (could be WT, indel, or knock-in) from the founder fish. Thus, a robust WT peak (denoted by green arrow) is detected in all pools with up to three additional peaks of unknown sizes. (A) Example of a positive pool with one embryo heterozygous for the knock-in allele (denoted by red arrow). Other peaks in this sample (denoted by the blue arrows) are random indels generated by NHEJ. (B) Example of a negative sample where all embryos are heterozygous for random indels (denoted by blue arrows). The x-axis shows the size of the peaks (bp), and the y-axis shows fluorescent intensity.
C3. Confirmation by screening individual embryos
At 1–5 dpf, collect 48 embryos (one embryo per well) in a 96-well plate from founders positive for germline transmission of knock-in peak in pooled embryo screening. This may require crossing the fish again and collecting a new clutch of embryos.
Extract DNA, perform fluorescent PCR, and run samples as described in section C1, steps 2–15. For point mutations, also perform restriction digest as described in section B2, steps 3–7.
Analyze data using the GeneMapper Software package to determine if any samples have the expected knock-in allele.
Note: Positive knock-in plots will have two peaks that represent the WT and knock-in allele. Examples can be seen in Carrington et al. (2022) in Figures 4, S2, and S5.
Verify precise knock-in by Sanger sequencing of positive samples and aligning reads to a reference knock-in DNA fragment.
C3. Establish stable lines
Outcross germline transmitting founders with WT fish and grow progeny to adulthood.
Fin clip F1 fish as adults and extract DNA, perform fluorescent PCR, and run samples as described in section C1, steps 2–15. For point mutations, also perform restriction digest as described in section B2, steps 3–7.
Analyze data in GeneMapper software and determine samples with the knock-in allele.
Verify precise knock-in by Sanger sequencing and aligning reads to a reference knock-in DNA fragment.
Note: It is recommended that all F1 fish are sequence confirmed to ensure all fish have precise knock-in at both ends.
F1 fish can be incrossed for subsequent experiments or phenotyping.
Data analysis
CRISPR-STAT plots and sequence chromatograms are the two main types of data generated during this procedure that require further analysis. CRISPR-STAT plots are generated and analyzed using 3730xl Data Collection software or its equivalent if a different capillary electrophoresis machine is used. The plots show peaks corresponding to all fragments present in an amplicon by their size (bp) along the x-axis and relative amount as fluorescence intensity along the y-axis. For sgRNA evaluation, peaks are analyzed for a decrease in the intensity of the WT peak with simultaneous presence of additional peaks indicating the sgRNA is active (Figure 3). For somatic and pooled embryo screening, plots are analyzed for the presence of a peak at the expected size of the knock-in allele (Figures 8 and 9). Using these methods, we screen embryos injected with CRISPR/Cas9 alone as control to determine the likelihood of random insertions leading to the same size peak as the expected peak with precise insertion of the repair template. An enrichment of samples with the expected size peak in the presence of repair template indicates successful knock-in. Sequence chromatograms are generated during confirmation of somatic knock-in by Sanger sequencing of TOPO clones from the positive sample, and germline knock-in by Sanger sequencing of heterozygous F1 embryos or adults for establishing stable lines. Sequencher or any equivalent software can be used to align sequence chromatograms with the reference sequence of the amplicon. For sequence data from somatic knock-in samples, three types of clones can be identified: 1) clones with an exact match to the WT sequence; 2) clones with random indels; 3) clones with precise knock-in. Determine the number of each type of clones to estimate the knock-in efficiency. Sequence chromatograms from heterozygous F1 samples can be analyzed either manually using Sequencher or by online tools, such as Poly Peak Parser (Hill et al., 2014) or CRISP-ID (Dehairs et al., 2016).
Validation of protocol
We and others have successfully used the protocols described here for sgRNA and SpCas9 synthesis, injections, evaluation of sgRNA activity by CRISPR-STAT, and genotyping of CRISPR/Cas9-mediated indels by fluorescent PCR for generation of gene knockouts (Varshney et al., 2016; Ramanagoudr-Bhojappa et al., 2018; Hoshijima et al., 2019; Pillay et al., 2022). The screening strategy for somatic knock-in in injected embryos using CRISPR-STAT with or without restriction digest has been validated by generation of fish lines with in-frame epitope tag insertions in two genes and a single nucleotide substitution leading to the patient-specific point mutation in one gene (Carrington et al., 2022).
General notes and troubleshooting
General notes
This protocol is designed and written for use with sgRNAs that can be synthesized with a T7 promoter (guide sequence begins with a “GG”). Alternate protocols for sgRNA synthesis, commercially available guides, and alternatives to Cas9 mRNA such as commercially available recombinant protein are also acceptable for use, as the only requirement is that the sgRNA/Cas9 should efficiently generate a DSB near the intended knock-in site. In addition, this protocol describes the use of the Sigma Extract-N-Amp kit for DNA extraction from zebrafish embryos and fin biopsies. However, cheaper and alternative DNA extraction methods, such as the HotSHOT method (Meeker et al., 2007), can be used as long as the dilution factor is adjusted to ensure robust PCR amplification of the target loci.
As with all genome editing using CRISPR/Cas9-based methods, selection of active guides that generate a DSB as close to the integration site as possible may be limited due to its sequence constraints. This limitation can be overcome by use of alternate Cas proteins with different PAM requirements, thus expanding the repertoire of sgRNAs. When using alternative Cas proteins such as Cas12a, different size homology arms may be needed (Fernandez et al., 2018); however, the screening can still be performed as described here. Another limitation that is not specific to this protocol is when a targeted gene is necessary for survival. Use of highly active guides to induce DSB often generates biallelic indel mutations, resulting in death of founder fish before they can be screened for germline transmission. This limitation can be overcome by reducing the amount of sgRNA in injection mixtures but may also reduce the knock-in efficiency. Alternatively, Cas9 expression can be targeted to germ cells using Cas9-nanos-3′UTR (Moreno-Mateos et al., 2015; Vejnar et al., 2016) to reduce somatic mutations in the body for increased survival of the injected fish.
The timing for each section in this protocol is given below:
Design phase
A1. sgRNA/Cas9 design and synthesis (2–3 days)
A2. Injections of sgRNA/Cas9 mRNA for evaluation of sgRNA activity (1 day)
A3. Determine activity of each sgRNA using CRISPR-STAT (1 day)
A4. Design of HDR template and screening primers (1 day)
Somatic screening phase
B1. Injections to deliver sgRNA/Cas9 and HDR template (1 day)
B2. Evaluation of somatic knock-in using CRISPR-STAT (1 day)
B3. Somatic knock-in confirmation by TOPO cloning and sequencing (3–4 days)
Germline screening phase
C1. Prioritization of founders to screen (1 day*)
C2. Screening prioritized founder fish by pooled embryos (4–5 days)
C3. Confirmation by screening individual embryos (4–5 days)
C3. Establish stable lines (4–5 days*)
*Does not include three months to grow embryos to adults.
Troubleshooting
For successful use of our screening methods, robust amplification of the target region by fluorescent PCR is very important. Therefore, always test fluorescent PCR primers on a few WT samples. If no peak is observed, run PCR products on a gel to ensure amplification. If there is no amplification, test alternate PCR conditions depending upon the annealing temperature of the primers or redesign primers. If multiple peaks are observed, sequence PCR products to identify endogenous indels and design new primers to avoid indels or use a cohort of fish with identical sequences of the target region for all experiments.
Acknowledgments
This research was funded by the Intramural Research Program of the National Human Genome Research Institute, National Institutes of Health (ZIC HG200351). This protocol is associated with Carrington et al. (2022).
Competing interests
The authors declare no competing interests.
Ethical considerations
All zebrafish experiments were performed in compliance with the National Institutes of Health guidelines for animal handling and research under NHGRI Animal Care and Use Committee (ACUC) approved protocol G-05-5.
References
Armstrong, G. A., Liao, M., You, Z., Lissouba, A., Chen, B. E. and Drapeau, P. (2016). Homology Directed Knockin of Point Mutations in the Zebrafish tardbp and fus Genes in ALS Using the CRISPR/Cas9 System. PLoS One 11(3): e0150188.
Bedell, V. M., Wang, Y., Campbell, J. M., Poshusta, T. L., Starker, C. G., Krug, R. G., 2nd, Tan, W., Penheiter, S. G., Ma, A. C., Leung, A. Y., et al. (2012). In vivo genome editing using a high-efficiency TALEN system. Nature 491(7422): 114-118.
Burg, L., Palmer, N., Kikhi, K., Miroshnik, E. S., Rueckert, H., Gaddy, E., MacPherson Cunningham, C., Mattonet, K., Lai, S. L., Marín-Juez, R., et al. (2018). Conditional mutagenesis by oligonucleotide-mediated integration of loxP sites in zebrafish. PLoS Genet 14(11): e1007754.
Carrington, B., Ramanagoudr-Bhojappa, R., Bresciani, E., Han, T. U. and Sood, R. (2022). A robust pipeline for efficient knock-in of point mutations and epitope tags in zebrafish using fluorescent PCR based screening. BMC Genomics 23(1): 810.
Carrington, B., Varshney, G. K., Burgess, S. M. and Sood, R. (2015). CRISPR-STAT: an easy and reliable PCR-based method to evaluate target-specific sgRNA activity. Nucleic Acids Res 43(22): e157.
de Vrieze, E., de Bruijn, S. E., Reurink, J., Broekman, S., van de Riet, V., Aben, M., Kremer, H. and van Wijk, E. (2021). Efficient Generation of Knock-In Zebrafish Models for Inherited Disorders Using CRISPR-Cas9 Ribonucleoprotein Complexes. Int J Mol Sci 22(17): 9429.
Dehairs, J., Talebi, A., Cherifi, Y. and Swinnen, J. V. (2016). CRISP-ID: decoding CRISPR mediated indels by Sanger sequencing. Sci Rep 6: 28973.
Fernandez, J. P., Vejnar, C. E., Giraldez, A. J., Rouet, R. and Moreno-Mateos, M. A. (2018). Optimized CRISPR-Cpf1 system for genome editing in zebrafish. Methods 150: 11-18.
Hill, J. T., Demarest, B. L., Bisgrove, B. W., Su, Y. C., Smith, M. and Yost, H. J. (2014). Poly peak parser: Method and software for identification of unknown indels using sanger sequencing of polymerase chain reaction products. Dev Dyn 243(12): 1632-1636.
Hoshijima, K., Jurynec, M. J., Klatt Shaw, D., Jacobi, A. M., Behlke, M. A. and Grunwald, D. J. (2019). Highly Efficient CRISPR-Cas9-Based Methods for Generating Deletion Mutations and F0 Embryos that Lack Gene Function in Zebrafish. Dev Cell 51(5): 645-657.e4.
Meeker, N. D., Hutchinson, S. A., Ho, L. and Trede, N. S. (2007). Method for isolation of PCR-ready genomic DNA from zebrafish tissues. Biotechniques 43(5): 610, 612, 614.
Moreno-Mateos, M. A., Vejnar, C. E., Beaudoin, J. D., Fernandez, J. P., Mis, E. K., Khokha, M. K. and Giraldez, A. J. (2015). CRISPRscan: designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nat Methods 12(10): 982-988.
Pillay, L. M., Yano, J. J., Davis, A. E., Butler, M. G., Ezeude, M. O., Park, J. S., Barnes, K. A., Reyes, V. L., Castranova, D., Gore, A. V., et al. (2022). In vivo dissection of Rhoa function in vascular development using zebrafish. Angiogenesis 25(3): 411-434.
Prykhozhij, S. V., Fuller, C., Steele, S. L., Veinotte, C. J., Razaghi, B., Robitaille, J. M., McMaster, C. R., Shlien, A., Malkin, D. and Berman, J. N. (2018). Optimized knock-in of point mutations in zebrafish using CRISPR/Cas9. Nucleic Acids Res 46(17): e102.
Ramanagoudr-Bhojappa, R., Carrington, B., Ramaswami, M., Bishop, K., Robbins, G. M., Jones, M., Harper, U., Frederickson, S. C., Kimble, D. C., Sood, R., et al. (2018). Multiplexed CRISPR/Cas9-mediated knockout of 19 Fanconi anemia pathway genes in zebrafish revealed their roles in growth, sexual development and fertility. PLoS Genet 14(12): e1007821.
Richardson, C. D., Ray, G. J., DeWitt, M. A., Curie, G. L. and Corn, J. E. (2016). Enhancing homology-directed genome editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA. Nat Biotechnol 34(3): 339-344.
Sieliwonczyk, E., Vandendriessche, B., Claes, C., Mayeur, E., Alaerts, M., Holmgren, P., Canter Cremers, T., Snyders, D., Loeys, B. and Schepers, D. (2023). Improved selection of zebrafish CRISPR editing by early next-generation sequencing based genotyping. Sci Rep 13(1): 1491.
Varshney, G. K., Carrington, B., Pei, W., Bishop, K., Chen, Z., Fan, C., Xu, L., Jones, M., LaFave, M. C., Ledin, J., et al. (2016). A high-throughput functional genomics workflow based on CRISPR/Cas9-mediated targeted mutagenesis in zebrafish. Nat Protoc 11(12): 2357-2375.
Vejnar, C. E., Moreno-Mateos, M. A., Cifuentes, D., Bazzini, A. A. and Giraldez, A. J. (2016). Optimized CRISPR-Cas9 System for Genome Editing in Zebrafish. Cold Spring Harb Protoc 2016(10). doi: 10.1101/pdb.prot086850.
Zhang, Y., Zhang, Z. and Ge, W. (2018). An efficient platform for generating somatic point mutations with germline transmission in the zebrafish by CRISPR/Cas9-mediated gene editing. J Biol Chem 293(17): 6611-6622.
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4,733 | https://bio-protocol.org/en/bpdetail?id=4733&type=0 | # Bio-Protocol Content
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Peer-reviewed
Simple Growth Complementation Assay in Yeast
RF Robert Fuhrmeister
JS Jana Streubel
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4733 Views: 491
Reviewed by: Anu P. Minhas Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular Plant Pathology Feb 2022
Abstract
The study of genes and their products is an essential prerequisite for fundamental research. Characterization can be achieved by analyzing mutants or overexpression lines or by studying the localization and substrate specificities of the resulting proteins. However, functional analysis of specific proteins in complex eukaryotic organisms can be challenging. To overcome this, the use of heterologous systems to express genes and analyze the resulting proteins can save time and effort. Yeast is a preferred heterologous model organism: it is easy to transform, and tools for genomics, engineering, and metabolomics are already available. Here, we describe a well-established and simple method to analyze the activity of plant monosaccharide transporters in the baker’s yeast, Saccharomyces cerevisiae, using a simple growth complementation assay. We used the famous hexose-transport-deficient yeast strain EBY.VW4000 to express candidate plant monosaccharide transporters and analyzed their transport activity. This assay does not require any radioactive labeling of substrates and can be easily extended for quantitative analysis using growth curves or by analyzing the transport rates of fluorescent substrates like the glucose analog 2-NBDG. Finally, to further simplify the cloning of potential candidate transporters, we provide level 0 modular cloning (MoClo) modules for efficient and simple Golden Gate cloning. This approach provides a convenient tool for the functional analysis of plant monosaccharide transporters in yeast.
Key features
• Comprehensive, simple protocol for analysis of plant monosaccharide transporters in yeast
• Includes optional MoClo parts for cloning with Golden Gate method
• Includes protocol for the production and transformation of competent yeast cells
Does not require hazardous solutions, radiolabeled substrates, or specialized equipment
Keywords: Yeast Heterologous expression Plant monosaccharide transporters EBY.VW4000 Drop-out assay
Background
In the past decade, the availability of full genome sequences, powerful genome engineering tools, and innovations in transcriptomics and metabolomics have revolutionized plant research (Rai et al., 2019; Purugganan and Jackson, 2021; Wang and Doudna, 2023). Plant sugar transporters are an important area of research, as they play a crucial role in delivering sugars as energy sources for developmental processes, contribute to disease resistance mechanisms, and are key determinants of yield (Julius et al., 2017; Bezrutczyk et al., 2018; Breia et al., 2021). However, the analysis of individual transporter candidates or transporter mutants is challenging because plant genomes are complex and sugar transporters are often organized in large gene families. To solve this bottleneck, heterologous systems, such as the baker’s yeast Saccharomyces cerevisiae, are powerful tools for analyzing plant sugar transporter functions on a single scale (Boles, 2003). Yeast is easy to transform, and many different strains with various phenotypes exist, which are suitable for analyzing specific sugar-transport abilities. In this protocol, we used the well-known hexose-transport-deficient strain EBY.VW4000 to analyze the activity of plant monosaccharide transporters and their mutants (Wieczorke et al., 1999). Instead of measuring the uptake of radioactively labeled substrates, as described in other protocols (Milne et al., 2017), this protocol describes a simple and cost-effective drop-out assay. Therefore, the hexose-transport-deficient strain EBY.VW4000 is transformed with expression constructs for the monosaccharide transporter of interest. A dilution series of the complemented EBY.VW4000 strains is spotted on selective plates to determine their growth on different carbon sources like maltose or glucose. The described protocol has already been extensively used in many studies dealing with sugar transporters (e.g., Chen et al., 2010; Moore et al., 2015; Skoppek et al., 2022; Tamayo et al., 2022; Yue et al., 2023). To provide a comprehensive workflow, we also implemented a protocol for the production of competent yeast cells and for transformation. This approach is based on the protocols described by Gietz and Schiestl (2007a and 2007b) and has been optimized for EBY.VW4000. Finally, we provide optional level 0 modular cloning (MoClo) modules for the yeast PMA1 promoter (plasma membrane H+ ATPase 1) and the ADH2 (alcohol dehydrogenase 2) terminator, which are commonly used for the expression of transporters in yeast (Meyer et al., 2006; Skoppek et al., 2022). These modules allow an easy and cost-effective cloning of new constructs using the Golden Gate cloning method (Engler et al., 2009; Werner et al., 2012).
The described drop-out assay is a simple method to rapidly screen for the transport activities of transporter candidates. However, if quantitative analysis is needed, we suggest extending this analysis to a growth curve or by analyzing the transport rate of fluorescent sugar analogs [for example esculin (sucrose analog) or 2-NBDG (glucose analog)] (Gora et al., 2012; Roy et al., 2015).
Materials and reagents
Biological materials
Saccharomyces cerevisiae strain EBY.VW4000 (hexose transport-deficient)
Genotype: CEN.PK2-1C hxt13Δ::loxP hxt15Δ::loxP hxt16Δ::loxP hxt14Δ::loxP hxt12Δ::loxP hxt9Δ::loxP hxt11Δ::loxP hxt10Δ::loxP hxt8Δ::loxP hxt4-1-5Δ::loxP hxt2Δ::loxP hxt3-6-7Δ::loxP gal2Δ::(ura3/FOA) stl1Δ::loxP agt1Δ::loxP mph2(ydl247w)Δ::loxP mph3(yjr160c)Δ::loxP
(Wieczorke et al., 1999)
For more information about the choice of suitable yeast strains see General Note 1.
Reagents
Agar-agar, Kobe I (Carl Roth, catalog number: 5210.5)
Peptone from meat (Carl Roth, catalog number: 2366.1)
Yeast synthetic drop-out medium supplements, without uracil (amino acid mix -Ura) Merck KGaA, catalog number: Y1501-20G)
Yeast extract (Carl Roth, catalog number: 2363.4)
Yeast nitrogen base without amino acids (YNB) (Merck KGaA, catalog number: Y0626-250G)
D-Glucose monohydrate (Duchefa B.V., catalog number: G0802.1000)
D(+)-maltose monohydrate (Carl Roth, catalog number: 8951.4)
Salmon sperm DNA sodium salt (Carl Roth, catalog number: 5434.1)
Lithium acetate dihydrate (LiAc) (LiCH3COO·2H2O) (Carl Roth, catalog number: 6713.1)
Poly(ethylene glycol) 3350, PEG3350, H(OCH2CH2)nOH (Merck KGaA, catalog number: P4338)
Dimethyl sulfoxide (DMSO), (CH3)2SO (Merck KGaA, catalog number: 276855-1L)
Glycerol (C3H8O3) 86% (Carl Roth, catalog number: 4043.3)
Sodium chloride (NaCl) (Carl Roth, catalog number: 3957.2)
Potassium chloride (KCl) (AppliChem GmbH, catalog number: A1039.1000)
Tris (C4H11NO3) (Carl Roth, catalog number: AE15.3)
Ethylenediamine tetra acetic acid disodium salt dihydrate (EDTA), C10H14N2Na2O8·2H2O (Carl Roth, catalog number: 8043.2)
1 N hydrochloric acid (HCl) or any other non-fuming HCl (Carl Roth, catalog number: 6792.1)
1 N sodium hydroxide (NaOH) or similar (Carl Roth, catalog number: 6785.1)
ddH2O (sterile)
Optional: MoClo plasmids level 0 and ready-to-use level 1
pJS506 [PMA1 promoter (Pro5U module), pICH41295, GGAG-AATG (BsaI)] (Addgene, #200712)
pJS237 [ADH2 terminator (3′U+Ter module), pUCGent, GCTT-CGCT (BsaI)] (Addgene, #200713)
pJS288 [Hxt1 transcription unit (Level 1 TU, P1), pAGT572, TGCC-GCAA (BpiI)] (Addgene, #200714)
Acceptor vectors: pAGT572 (selection marker for uracil) was a kind gift from Sylvestre Marillonnet and Alain Tissier (Scheler et al., 2016). Alternative yeast-compatible vectors is, for example, pJOG417 (Addgene #105341; selection marker for leucine) (Gantner et al., 2018)
Solutions
20% glucose solution (sterile filtered)
20% maltose solution (sterile filtered)
14% glycerol solution
YPM medium (see Recipe 1)
Frozen competent cell solution (FCC) (see Recipe 2)
10× yeast synthetic drop-out medium supplements, without uracil, (amino acid mix -Ura) (see Recipe 3)
YNB medium with amino acid mix -Ura and carbon source (see Recipe 4)
50% PEG 3350 (autoclaved) (see Recipe 5)
1 M LiAc (sterile filtered) (see Recipe 6)
Salmon sperm carrier DNA (2 mg/mL) (see Recipe 7)
1 M Tris-HCl (pH 8.0) (see Recipe 8)
0.2 M EDTA (pH 8.0) (see Recipe 9)
10 mM TE buffer (pH 8.0) (see Recipe 10)
Recipes
YPM medium (full medium with maltose)
Reagent Final concentration Amount
Yeast extract 1% (w/v) 5 g
Peptone 2% (w/v) 10 g
Maltose 2% (w/v) 10 g
Agar-agar (optional for plates) 2% (w/v) 10 g
ddH2O n/a Ad 500 mL
Autoclave. Pour plates with solid medium.
Note: If needed for a certain yeast strain, the carbon source can be replaced by glucose, galactose, or glycerol. Maltose was most suitable to grow EBY.VW4000.
Frozen competent cell solution (FCC)
Reagent Final concentration Amount
Glycerol 5% (v/v) 5 mL
DMSO 10% (v/v) 10 mL
Total n/a Ad 100 mL
Store FCC at room temperature (RT).
10× yeast synthetic drop-out medium supplements, without uracil (amino acid mix -Ura)
Reagent Final concentration Amount
Yeast synthetic drop-out medium supplements, without uracil (amino acid mix -Ura) 10× 1.92 g
ddH2O n/a Ad 100 mL
Filter sterilize. Store at 4 °C and protect from light.
Note: To dissolve the supplements in water, stir the solution on a magnetic stirrer. Carefully warm the solution (maximum 50 °C) if the supplements do not dissolve well. Always tightly close the stock bottle with the supplements after usage, to prevent the powder from absorbing water. For additional information on how to choose the yeast synthetic drop-out medium, see General Note 2.
YNB medium with amino acid mix -Ura and carbon source
To prepare 500 mL of YNB medium with amino acid mix -Ura and carbon source, first weigh the required amount of YNB and dissolve in 400 mL of water. For solid plates, add 2% agar-agar. Autoclave and store until usage.
Reagent Final concentration Amount
YNB n/a 3.35 g
Agar-agar (optional for plates) 2% (w/v) 10 g
ddH2O n/a Ad 400 mL
For solid medium: Carefully boil up the medium in a microwave. Secondly, add the following supplements before use:
Reagent Final concentration Amount
20% carbon source1 2% (v/v) 50 mL
10× amino acid mix -Ura (Recipe 3) 1% (v/v) 50 mL
For solid medium: pour plates.
1The carbon source is chosen according to the intended use of the medium. For the required carbon source, e.g., glucose, maltose, or similar, a 20% stock solution is prepared in water, filter sterilized, and stored at RT until use. For selection and cultivation of transformed EBY.VW4000 strains, add maltose as carbon source. For performing the drop-out assay on different carbon sources, add the required sugar, e.g., glucose.
50% PEG 3350
Reagent Final concentration Amount
PEG 3350 50 % 50 g
ddH2O n/a Ad 100 mL
Autoclave, store at RT.
1 M LiAc
Reagent Final concentration Amount
PEG 3350 1 M 10.2 g
ddH2O n/a Ad 100 mL
Autoclave, store at RT.
Salmon sperm carrier DNA (2 mg/mL)
Reagent Final concentration Amount
Salmon sperm carrier DNA 2 mg /mL (w/v) 8 g
10 mM TE buffer (Recipe 8) 10 mM Ad 4 mL
Dissolve the salmon sperm carrier DNA in 10 mM TE buffer on ice for 3–4 h with gentle shaking. Prepare aliquots, for example with 500 μL each. Incubate the aliquots for 10 min at 95 °C and immediately transfer to ice. Store at -20 °C.
1 M Tris-HCl (pH 8.0)
Reagent Final concentration Amount
Tris 1 M 12.114 g
ddH2O n/a Ad 100 mL
Dissolve Tris in 80 mL of ddH2O and adjust the pH to 8.0 by carefully adding drops of HCl (1 N or any other non-fuming HCl). Fill up the volume to 100 mL.
0.2 M EDTA
Reagent Final concentration Amount
EDTA 200 mM 7.44 g
ddH2O n/a Add 100 mL
Dissolve EDTA in 80 mL of ddH2O and adjust the pH to 8.0 by carefully adding NaOH (1 N NaOH solution or pellets). Fill up the volume to 100 mL.
10 mM TE-buffer (pH 8.0)
Reagent Final concentration Amount
Tris-HCl pH 8.0 10 mM 1 mL
EDTA pH 8.0 1 mM 0.5 mL
ddH2O n/a 98.5 mL
Laboratory supplies
Squared Petri dishes for drop-out (Carl Roth, catalog number: EL50.1)
Petri dish (Sarstedt, catalog number: 82.1473.001)
2 mL reaction tube (Sarstedt, catalog number: 72.695.500)
Parafilm
Disposable gloves
50 mL reaction tubes (Sarstedt, catalog number: 62.547.254)
Equipment
Biosafety cabinet or bench with flame
Pipettes
Photometer (e.g., IMPLEN, OD600 DiluPhotometer; or similar)
Tabletop centrifuge (e.g., Eppendorf Micro Centrifuge 5425, maximum 21,300 rcf or similar)
Centrifuge for 50 mL reaction tubes (e.g., Eppendorf 5810R, maximum 20,913 rcf or similar)
Mobile phone or any other camera device
Optional: INTEGRA multichannel pipet Voyager (INTEGRA Biosciences GmbH, catalog number: 4721)
Optional: INTEGRA tips (INTEGRA Biosciences GmbH, e.g., as box 3416 or as bag 4411)
Procedure
Optional: available modular cloning (MoClo) modules for construct design
A helpful protocol on how to design and clone constructs by using the Golden Gate cloning and the MoClo system can be found in Engler et al. (2014).
If cloning is planned with this method, the provided level 0 PMA1 promoter and ADH2 terminator module can be used (Figure 1A).
As level 1 acceptor vector, choose a yeast-compatible one. Here, the MoClo level 1 acceptor vector pAGT572 (uracil selection) was used (Figure 1B and 1C).
Figure 1. Golden Gate cloning strategy with the provided modular cloning (MoClo) modules. A. We provide the level 0 MoClo modules for the PMA1 promoter (Pro+5U module) and the ADH2 terminator (3U+Ter module). The candidate transporter gene has to be amplified with CDS1 stop or without stop (CDS1 no stop) according to the MoClo guidelines. The latter can be used to fuse a C-terminal tag (CT). The required BsaI overhangs for cloning are indicated for each module. B. In this example, the yeast-compatible level 1 acceptor vector pAGT572 with uracil selection was used. Cloning of the level 0 modules into the level 1 acceptor vector is done in a cut-ligation with BsaI. C. The final level 1 transcription unit contains all parts required for yeast transformation and expression of the candidate transporter.
Remember to clone suitable negative and positive controls for the drop-out assay. We usually use an empty plasmid or GFP on a plasmid as negative control and the yeast monosaccharide transporter Hxt1 (NCBI 856494) as positive control. The transcription unit (pJS288) for the positive control used in this protocol (Hxt1 under control of the PMA1 promoter and ADH2 terminator cloned in pAGT572) is available via Addgene #200714.
For more background information, see General Note 3.
Preparation of competent EBY.VW4000 cells
A helpful protocol on how to prepare competent yeast cells can be found in Gietz and Schiestl (2007a).
For EBY.VW4000, use the following modifications of this protocol:
Streak out a stock of EBY.VW4000 cells on a plate with solid YPM medium (Recipe 1) (single colony needed). Incubate at 28 °C for two days.
Pick a single yeast colony and inoculate a 50 mL preculture by using liquid YPM medium. Incubate the culture overnight at 28 °C on a shaker at 140 rpm.
Take a 1 mL sample and dilute 1:10 with liquid YPM. Mix the sample before diluting and before measuring. Measure the OD600 of the 1:10 dilution.
Calculate the volume of preculture needed to inoculate a 200 mL main culture with an OD600 of 0.1 in YPM medium. Do not forget to include the dilution factor in your calculation.
Incubate the yeast cells at 28 °C on a shaker at 140 rpm until an OD600 between 0.6 and 0.8 is reached.
Note: Regularly check the OD600. Do not overgrow the culture. In our hands, it took approximately 5 h to reach an OD600 between 0.6 and 0.8.
Perform the next steps at RT.
Split the culture equally into six 50 mL reaction tubes (approximately 33.33 mL for each tube).
Note: For centrifugation, we never fill the tubes with the maximum possible volume. We usually centrifuge a maximum of 2/3 of the total tube volume. This prevents spilling during centrifugation, as tubes are not tightly closed.
Centrifuge at 3,000× g for 5 min.
Discard the supernatant.
Add 16.6 mL (0.5 V of the starting culture, divided by six tubes: 0.5 × 200 mL/6) of sterile ddH2O to each tube and resuspend the pellet.
Combine two resuspended pellets in one tube. This results in three tubes with approximately 33.2 mL each.
Centrifuge at 3,000× g for 5 min.
Discard the supernatant.
Resuspend the three pellets by adding 0.666 mL (0.01 V of the starting culture, divided by three tubes: 0.01 × 200 mL/3) of sterile ddH2O to each tube. Combine the resuspended pellets of all three tubes into one tube (total volume approximately 1.99 mL).
Centrifuge at 3,000× g for 5 min.
Discard the supernatant.
Resuspend the pellet by adding 2 mL (0.01 V of the starting volume: 0.01 × 200 mL) of FCC solution (Recipe 2) to each tube.
Prepare aliquots of 50 μL competent cells in 1.5 mL tubes for subsequent transformation.
Store at -20 °C for at least 3 h (overnight is also possible).
Note: Slowly freezing the cells is essential for survival.
Store at -80 °C for long-term storage.
Optional: To check the general viability of the competent cells, plate a 1:100 dilution (in ddH2O) of one aliquot on YPM medium and incubate overnight at 28 °C. The plate should be nicely overgrown with yeast.
Yeast EBY.VW4000 transformation
This method is based on the protocol for LiAc/SS carrier DNA/PEG transformation by Gietz and Schiestl (2007b), with the following modifications:
Thaw competent EBY.VW4000 cells on ice (50 μL per transformation).
For each plasmid, one transformation mix is prepared:
Mix the plasmid with ddH2O for a total volume of 14 μL and 600 ng plasmid in a 1.5 mL reaction tube. Vortex. Prepare one negative control with 14 μL of ddH2O only.
Add 260 μL of 50% PEG3350 (Recipe 5), 36 μL of 1 M LiAc (Recipe 6), and 50 μL of salmon sperm carrier DNA (2 mg/mL) (Recipe 7).
Mix well.
Note: PEG3350 is very viscous. Pipette slowly and ensure that the transformation mix is well mixed.
Store transformation mix at RT until use.
Centrifuge competent yeast cells at 13,000× g for 2 min at RT.
Discard the supernatant.
Add the transformation mix, including the plasmid.
Vortex or mix by pipetting up and down until the pellet is resuspended.
Incubate for 60 min at 42 °C (water bath or heat block)
Note: If not in stock, pour YNB plates with selective amino acid mix during the incubation time (Recipe 3 and Recipe 4).
Centrifuge at 13,000× g for 30 s.
Discard the supernatant.
Add 1 mL of sterile ddH2O.
Resuspend the pellet by pipetting up and down.
Plate 50–200 μL on YNB medium (with selective amino acid mix, in our setup -Ura, and maltose).
Note: We usually prepare two plates: one with 50 μL and a backup with 200 μL of transformed cells. The plasmids we are using contain a 2 μ origin of replication and typically yield between 50 and 100 clones when we plate 50 μL. The transformed cells can be stored at 4 °C and plated again one day later if needed.
Incubate the cells for 2–3 days at 28 °C.
Transfer 2–4 single colonies per transformation to a new YNB selection plate and grow overnight at 28 °C.
To generate long-term stocks: scrape off one of the clones, resuspend in sterile 14% glycerol solution, and store at -80 °C. Usually, we prepare stocks of 2–3 individual clones.
Yeast sugar uptake analysis via drop-out assay
Prepare YNB plates with selective amino acid mix and the required carbon source in squared Petri dishes for the drop-out assay. Squared Petri dishes offer more space to analyze several strains and dilutions on one plate. In our experiment, we used YNB -Ura + maltose and YNB -Ura + glucose.
Note: For yeast sugar uptake analysis, we usually include a negative control strain with an empty plasmid or GFP on a plasmid. As positive control, we use the yeast monosaccharide transporter Hxt1. We usually analyze at least two independent clones of the transporter of interest.
Plate two clones per strain (fresh transformed or from stock) onto fresh YNB selective medium with maltose two days prior to the drop-out assay.
The following steps are prepared at RT with sterile conditions, for example by working in a biosafety cabinet or next to a flame. Performing the assay under sterile conditions is recommended to avoid contamination of the yeast strains or plates.
Prepare a 1.5 mL reaction tube with 1 mL of ddH2O for each clone.
Scrape the colonies from the plates and resuspend in the tubes.
Prepare a 1:10 dilution with ddH2O and measure the OD600.
Calculate the volume needed for 1.5 mL of cell suspension with an OD600 of 0.4. Do not forget to include the dilution factor.
Prepare 1.5 mL of cell suspension with an OD600 of 0.4 in a new 2 mL reaction tube. Vortex.
Take 1 mL to check the OD600. Adjust again if needed.
Prepare a serial dilution (500 μL volume) for each transformant in a new sterile 1.5 mL tube.
Always vortex the solutions before preparing the next dilution.
Label the squared plates for the drop-out assay. We usually use grid lines to generate nicely distributed drops (Table 1 and Figure 2A).
Table 1. Example layout for drop-out grid lines
OD600 0.4 0.04 0.004 0.0004
Strain 1
Strain 2
Strain 3
Strain 4
Strain 5
Strain 6
Figure 2A shows an example of a possible labeling of the plates; the individual yeast transformants are ordered vertically and the corresponding serial dilution horizontally. At least two types of plates are required for the assay: one with YNB -Ura + maltose and one with YNB -Ura + glucose.
Organize your tubes in a rack according to the droplet scheme on the plates (e.g., yeast transformants vertically, serial dilutions horizontally).
Always vortex each tube immediately before pipetting.
We prefer using a multichannel pipette for the drops as it is faster. This might require a little bit of training to place the drops correctly. You can also use a normal single-channel pipette.
Note: The serial dilution and the spotting of the droplets can also be performed using a sterile 96-well plate and a microplate replicator (e.g., Millipore Sigma, Replica plater for 96-well plate, R2508 or similar).
Drop 3 μL of each sample on the two plates. Always ensure that the pipette tips in the multichannel pipette are loaded equally. Avoid pricking the tips into the agar (Figure 2B). After pipetting, the drops should be visible on the plate (Figure 2C).
Figure 2. Preparation and procedure of the yeast drop-out assay. A. Material needed. Prepare a vortex, pipette, and pipette tips. Prepare the plates needed with a pipetting scheme. In this case, a grid line was used, with the yeast strains ordered vertically and serial dilutions horizontally. Prepare a rack with the tubes ordered in the same scheme as the plate. B. A multichannel pipette is used to place the drops on the plate. C. Liquid drops visible on the plate after pipetting.
Dry the plates (opened lid) under sterile conditions until no liquid drops are visible.
Incubate the plates for 48 h at 28 °C. If the colonies are not clearly visible, the incubation time can be prolonged.
Document the growth of the colonies by using a normal mobile phone camera or any other device. We typically take pictures with the Bio-Rad Chemidoc using the colorimetric mode (Figure 3).
Data analysis
The colonies growing on YNB plates with maltose should all grow in a comparable range. The colonies on YNB plates containing glucose generally grow a little bit slower compared to maltose. If the expressed plant sugar transporter complements the growth defect of EBY.VW4000 on hexose-containing medium, these colonies should grow. The negative control (empty vector or other protein cloned, e.g., GFP) should not grow on glucose-containing medium. As positive control, we usually use Hxt1, a sugar transporter from yeast.
Figure 3. Example pictures of a drop-out assay result for different sugar transport protein (STP) variants (STP_V1 to STP_V6) expressed in yeast. The STP variants (STP_V1 to STP_V6) were transformed to the hexose-transport-deficient yeast strain EBY.VW4000. A serial dilution of the respective strains was dropped on selective medium either containing 2% maltose or 2% glucose as carbon source. Selective medium lacking uracil was used (-Ura). Plates were grown for two days at 28 °C and growth was documented using the Chemidoc from Bio-Rad. GFP was used as negative control and the yeast hexose transporter Hxt1 served as positive control.
General notes and troubleshooting
Yeast is a highly suitable heterologous system to express and analyze the functions of certain proteins on an individual scale. Besides the described plant monosaccharide transporters (sugar transport protein family), other transporters, for other substrates or from other organisms, can be analyzed using the described drop-out assay and growth complementation. Therefore, a suitable yeast strain has to be chosen with a deficiency in transporting a specific substrate. This deficiency can be complemented by the expression of the respective transporter, thus conferring growth on selective medium or transport of fluorescent substrates. For example, members of the barley and potato SWEET sugar transporter family were also analyzed using yeast strain EBY.VW4000 and the substrates glucose, fructose, galactose, or mannose (Tamayo et al., 2022; Yue et al., 2023). The type I sucrose transporters StSUT1 from potato and AtSUC2 from Arabidopsis were analyzed using the yeast strain SEY6210 and BY4742 and the fluorescent sucrose analog esculin (Gora et al., 2012). The Arabidopsis γ-aminobutyric acid (GABA) transporter AtGAT1 was analyzed using the yeast strain 22574d and selective medium with different nitrogen sources (citrulline, proline, or GABA) (Meyer et al., 2006).
Different yeast synthetic drop-out medium supplements are available. Always choose the required supplements fitting to your plasmid and the genotype of the used yeast strain. In this protocol, we used the yeast strain EBY.VW4000, that is auxotrophic for uracil, leucine, histidine, and tryptophan. The plasmid pAGT572 encodes the URA3 selection marker to identify transformed EBY.VW4000 clones on selective medium lacking uracil.
For our protocol, we provide plasmids that allow the Golden Gate–based cloning of yeast expression constructs. To clone the transporter candidates, a commonly used vector is pDR196 (Meyer et al., 2006; Chen et al., 2010; Gora et al., 2012; Roy et al., 2015; Tamayo et al., 2022; Yue et al., 2023). Here, the candidate transporter sequence is inserted between the PMA1 promoter (plasma membrane H+ ATPase 1) and the ADH2 (alcohol dehydrogenase 2) terminator by using a multiple cloning site and type IIP restriction enzyme (e.g., EcoRI). In our laboratory, we use the simple and efficient Golden Gate cloning with the standardized modular cloning (MoClo) syntax to flexibly generate single or multigene constructs. The Golden Gate cloning method is based on the use of type IIS restriction enzymes that cut outside of their recognition sequence and generate user-defined overhangs. These overhangs allow the pre-defined, overhang-dependent, and scarless fusion of modules by using a one-pot restriction and ligation reaction (Engler et al., 2009; Weber et al., 2011; Werner et al., 2012). In addition, the standardized MoClo syntax allows an easy and flexible exchange of modules according to the requirement of the user. To our knowledge, no modules were available for the standard MoClo syntax to flexibly clone yeast expression constructs comparable to the pDR196 setup. Therefore, we provide publicly available level 0 MoClo modules for the yeast PMA1 promoter and the ADH2 terminator, to clone a transcription unit for the candidate transporter by using Golden Gate. In addition, we also provide a ready-to-use transcription unit to express the positive control Hxt1 (Skoppek et al., 2022).
Acknowledgments
This work was funded by university core funding only. We thank Jens Boch for general support. The protocol was used in Skoppek et al. (2022).
Competing interests
The authors declare to have no competing interests.
References
Bezrutczyk, M., Yang, J., Eom, J. S., Prior, M., Sosso, David., Hartwig, T., Szurek, Boris., Oliva, R., Vera-Cruz, C., White, F. F., et al. (2018). Sugar flux and signaling in plant-microbe interactions. Plant J 93(4): 675-685.
Boles, E. (2003). Yeast as a Model System for Studying Glucose Transport. In: Sibley, D. R. and Quick, M. W. (Eds.). Transmembrane Transporters. Wiley‐Liss, Inc.
Breia, R., Conde, A., Badim, H., Fortes, A. M., Gerós, H. and Granell, A. (2021). Plant SWEETs: from sugar transport to plant–pathogen interaction and more unexpected physiological roles. Plant Physiol 186(2): 836-852.
Chen, L. Q., Hou, B. H., Lalonde, S., Takanaga, H., Hartung, M. L., Qu, X. Q., Guo, W. J., Kim, J. G., Underwood, W., Chaudhuri, B., et al. (2010). Sugar transporters for intercellular exchange and nutrition of pathogens. Nature 468(7323): 527-532.
Engler, C., Gruetzner, R., Kandzia, R. and Marillonnet, S. (2009). Golden gate shuffling: a one-pot DNA shuffling method based on type IIs restriction enzymes.PLoS One 4(5): e5553.
Engler, C., Youles, M., Gruetzner, R., Ehnert, T. M., Werner, S., Jones, J. D., Patron, N. J. and Marillonnet, S. (2014). A golden gate modular cloning toolbox for plants. ACS Synth Biol 3(11): 839-843.
Gantner, J., Ordon, J., Ilse, T., Kretschmer, C., Gruetzner, R., Lofke, C., Dagdas, Y., Burstenbinder, K., Marillonnet, S. and Stuttmann, J. (2018). Peripheral infrastructure vectors and an extended set of plant parts for the Modular Cloning system. PLoS One 13(5): e0197185.
Gietz, R. D. and Schiestl, R. H. (2007a). Frozen competent yeast cells that can be transformed with high efficiency using the LiAc/SS carrier DNA/PEG method. Nat Protoc 2(1): 1-4.
Gietz, R. D. and Schiestl, R. H. (2007b). High-efficiency yeast transformation using the LiAc/SS carrier DNA/PEG method. Nat. Protoc. 2: 31-34.
Gora, P. J., Reinders, A. and Ward, J. M. (2012). A novel fluorescent assay for sucrose transporters. Plant Methods 8: 13.
Julius, B. T., Leach, K. A., Tran, T. M., Mertz, R. A. and Braun, D. M. (2017). Sugar Transporters in Plants: New Insights and Discoveries. Plant Cell Physiol 58(9): 1442-1460.
Meyer, A., Eskandari, S., Grallath, S. and Rentsch, D. (2006). AtGAT1, a high affinity transporter for γ-aminobutyric acid in Arabidopsis thaliana. J Biol Chem 281(11): 7197-7204.
Milne, R. J., Dibley, K. E. and Lagudah, E. S. (2017). Yeast as a heterologous system to functionally characterize a multiple rust resistance gene that encodes a hexose transporter. In: Periyannan, S. (Ed.). Wheat Rust Diseases. Methods in Molecular Biology, vol 1659. Humana Press, New York, NY.
Milne, R. J., Dibley, K. E. and Lagudah, E. S. (2017). Yeast as a heterologous system to functionally characterize a multiple rust resistance gene that encodes a hexose transporter. In: Periyannan, S. (Ed.). Wheat Rust Diseases. Methods in Molecular Biology (pp. 265-274). Humana Press.
Moore, J. W., Herrera-Foessel, S., Lan, C., Schnippenkoetter, W., Ayliffe, M., Huerta-Espino, J., Lillemo, M., Viccars, L., Milne, R., Periyannan, S., et al. (2015). A recently evolved hexose transporter variant confers resistance to multiple pathogens in wheat. Nat Genet 47(12): 1494-1498.
Purugganan, M. D. and Jackson, S. A. (2021). Advancing crop genomics from lab to field. Nat Genet 53: 595-601.
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Roy, A., Dement, A. D., Cho, K. H. and Kim, J. H. (2015). Assessing glucose uptake through the yeast hexose transporter 1 (Hxt1). PLoS One 10(3): e0121985.
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Skoppek, C. I., Punt, W., Heinrichs, M., Ordon, F., Wehner, G., Boch, J. and Streubel, J. (2022). The barley HvSTP13GR mutant triggers resistance against biotrophic fungi. Mol Plant Pathol 23(2): 278-290.
Tamayo, E., Figueira-Galan, D., Manck-Gotzenberger, J. and Requena, N. (2022). Overexpression of the Potato Monosaccharide Transporter StSWEET7a Promotes Root Colonization by Symbiotic and Pathogenic Fungi by Increasing Root Sink Strength. Front Plant Sci 13: 837231.
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4,734 | https://bio-protocol.org/en/bpdetail?id=4734&type=0 | # Bio-Protocol Content
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Using Fiber Photometry in Mice to Estimate Fluorescent Biosensor Levels During Sleep
MA Mie Andersen
AT Anastasia Tsopanidou
TR Tessa Radovanovic
VC Viviane Noelani Compere
NH Natalie Hauglund
MN Maiken Nedergaard
CK Celia Kjaerby
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4734 Views: 797
Reviewed by: Nafisa M. JadavjiNoa matosevichAlejandro Osorio-Forero
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Original Research Article:
The authors used this protocol in Nature Neuroscience Jul 2022
Abstract
Sleep is not homogenous but contains a highly diverse microstructural composition influenced by neuromodulators. Prior methods used to measure neuromodulator levels in vivo have been limited by low time resolution or technical difficulties in achieving recordings in a freely moving setting, which is essential for natural sleep. In this protocol, we demonstrate the combination of electroencephalographic (EEG)/electromyographic (EMG) recordings with fiber photometric measurements of fluorescent biosensors for neuromodulators in freely moving mice. This allows for real-time assessment of extracellular neuromodulator levels during distinct phases of sleep with a high temporal resolution.
Keywords: Fluorescence Biosensors Fiber photometry EEG Sleep Noradrenaline Locus coeruleus
Background
In comparison to neurotransmitters such as glutamate and GABA, neuromodulators—norepinephrine, dopamine, serotonin, etc.—exhibit a slower and longer-ranging diffuse form of transmission, typically propagated by the activation of metabotropic G protein–coupled receptors (GPCRs). Until recently, electrochemical and microdialysis-based approaches (Shouse et al., 2000; Léna et al., 2005; Park et al., 2011) were the golden standard in the measurement of neuromodulator levels. However, technical and temporal constraints associated with these techniques prohibited the estimation of faster neuromodulator dynamics in freely moving mice. These constraints were recently eliminated by the development of novel fluorescent biosensors, which allow for specific and sensitive measurements of diverse neuromodulators with a high temporal resolution by employing the corresponding GPCRs as the sensing module. This permits inference about causal relationships between neuromodulators and behavioral and biological phenomena.
More specifically, the biosensors are based on the insertion of a perturbed fluorescent protein on the neuromodulator-specific GPCR, which, upon binding of the corresponding neuromodulator, generates an increase in fluorescence (Patriarchi et al., 2018; Sun et al., 2018; Feng et al., 2019). Fiber photometry utilizes this increase in fluorescence to detect neuromodulator levels in freely moving mice. Chronic implantation of an optical fiber in the brain region where the biosensor is expressed allows for the simultaneous delivery of excitation light and collection of fluorescence emission, thus enabling the real-time imaging of neuromodulator levels. This creates the unprecedented opportunity to study fast neuromodulator responses across natural stress-free behaviors such as sleep–wake transitions.
In addition, the combination of biosensor measurements with the use of fluorescent calcium indicators allows for the correlation of the calcium activity of the neuronal population releasing the neuromodulator of interest with the extracellular neuromodulator levels, a method that we and others recently implemented for the study of the locus coeruleus-norepinephrine (LC-NE) system during sleep (Osorio-Forero et al., 2021; Antila et al., 2022; Kjaerby et al., 2022). Furthermore, by combining electroencephalographic (EEG)/electromyographic (EMG) recording measurements with fiber photometric measurements, we were able to assess LC-NE dynamics during sleep and uncover the underlying intrinsic relationship of the LC-NE system, sleep micro-architecture, and memory performance. Finally, apart from their biological significance, these distinct features of norepinephrine signaling during sleep show unique potential as new sleep scoring markers for the refined classification of sleep into wake, micro-arousals, NREM, and REM sleep.
Biosensors not only allow us to assess the relative changes in neuromodulator dynamics during sleep and wake behaviors but also to compare upregulating or downregulating treatment effects. However, to assess the absolute levels of neuromodulators, microdialysis still constitutes the most precise technique, since the delivery and expression of biosensor constructs as well as the placement of the fibers greatly impact the measurement of raw fluorescence levels. For this reason, the raw fluorescence signal is normalized according to a control isosbestic channel, thus allowing for measurements of only relative fluctuations. Importantly, it should also be noted that fluorescent indicators can be subject to tissue artifacts such as pH changes (Bizzarri et al., 2009; Remington, 2011) and hemoglobin levels (Zhang et al., 2022) that affect fluorescence levels. This observation is particularly relevant in the study of sleep and especially during REM sleep (Kjaerby et al., 2022), when marked vasodilation occurs (Turner et al., 2020 and 2023; Bojarskaite et al., 2022) alongside predicted quenching of fluorescence. Therefore, measurement of Ca2+ independent fluorescence by the use of the isosbestic point of the fluorescent indicator or simultaneous imaging of an inert red fluorescent marker (Zhang et al., 2022) is necessary for the removal of these artifacts.
Materials and reagents
Electrode preparation for EEG and EMG
Breakaway female header pins (SWISS MACHINE PIN, Let Elektronik, Sparkfun electronics, catalog number: PRT-00743)
EEG screw electrodes (prefabricated low-impedance EEG screw electrodes from NeuroTek, ~0.8 mm OD, with a length of 2–3 mm)
EMG wire on roll, Teflon-coated stainless-steel wire (W3 Wire International, catalog number: W3 632)
Lead-free soldering tin wire (Goobay, catalog number: 40844)
Super glue (e.g., Loctite Super Glue Power Flex)
Glue accelerator, Insta-Set Super Glue Accelerator (Bob Smith Industrie, catalog number: BSI-152)
6-channel cable with open-ended connection (PlasticsOne, catalog number: 363-000)
Metal braided cable sleeving, tinned copper, 6 mm diameter (CableOrganizer, catalog number: MBN0.25-10FT)
Surgical tape, 2.5 cm (3M, Micropore tape, catalog number: B10428996)
Surgery
Adult mouse (C57BL/6J, 7–9 weeks old)
Preoperative analgesia (e.g., buprenorphine; working concentration: 0.01 mg/mL)
Local anesthetic (e.g., lidocaine; working concentration: 0.2 mg/mL)
Post operative analgesia (e.g., carprofen; working concentration: 1 mg/mL)
Isoflurane (Attane vet®, Piramal, 1,000 mg/g)
Shaver, Aesculap Isis rodent shaver (Agnthos, catalog number: GT421)
70% ethanol (EtOH)
Distilled water
10% iodine solution
1 ml syringes (Chirana T. InjectaM, catalog number: CHINS01)
30 G × 12 mm needles (Sterican®, B Braun, catalog number: 4656300)
Cotton swabs (Th. Geyer, catalog number: 6085021)
Disposable wipes KIMTECH® (VWR, catalog number: 115-2221)
Eye drops (e.g., Ophtha Neutral eye gel)
Surgical marker pen (e.g., Universal Surgical Skin Marking Pen Fine, Premier HH Ltd., catalog number: PEN002)
Biosensor virus (e.g., BrainVTA, WZ Biosciences, Addgene, titer 1012 gc/mL)
Parafilm (Sigma-Aldrich, catalog number: P7793-1EA)
EEG/EMG electrodes (see section A)
Mono optic fiber implant (400 μm core, NA = 0.48, receptacle: MF2.5, Doric Lenses)
Super-Bond C&B Dental cement (Generique International, catalog numbers: 7110, 7111-100, T060E)
Super-Bond C&B Green Activator (Prestige Dental, catalog number: 7115-100)
Recording
White bedding, ALPHA-dri Dust free (LBS Biotechnology, catalog number: 1032003)
White nesting, soft paper wool (LBS Biotechnology, catalog number: 1034007)
Wooden stick, food pellets, water bottle
Plastic cylinder, Plexiglas, H 40 cm × Ø 30 cm
Fiber-optic patch cord (400 μm core, NA = 0.48, 1.5 m, Doric Lenses)
Zirconia sleeves (Ø 2.5 mm)
EEG/EMG patch cords (see section A)
Equipment
The list below describes equipment that we use but that can be replaced with equivalent models.
Electrode preparation for EEG and EMG
Forceps (e.g., Agnthos, Fine Graefe Titanium, catalog number: 11650-10)
Side-cutting pliers (e.g., Stanley, catalog number: 84-079)
End-cutting pliers (e.g., Stanley, catalog number: 84-079)
Soldering clamp stand
Soldering iron (Weller, WE1010 Soldering station, 70 W, 100–450 °C, LCD, ESD, catalog number: H42077)
Multimeter (e.g., XL830L Digital Multimeter)
Scalpel
Microscope (e.g., Olympus, SZ61)
Fine scissors (e.g., Agnthos, catalog number: 14184-09)
Surgery
Flow table
Surgery microscope (e.g., Nikon SMZ745T)
Isoflurane high-flow vaporizer with induction chamber (Kent Scientific, catalog numbers: VetFlo-1205S and VetFlo-0530SM)
Stereotactic frame with nose cone (RWD, catalog number: 68025)
Gooseneck lights (VWR, catalog number: 631-1755)
Heating pads: small surgical pad, big recovery pad (Stoelting, catalog numbers: 53850, 53850M, and 53850C)
Surgery tools: blunt forceps (Fine Science Tools, catalog number: 11002-12), fine scissors (Fine Science Tools, catalog number: 14084-08), fine angled forceps (Fine Science Tools, catalog number: 11251-35)
Screwdriver for EEG screws
Electric drill with drill tip, size 005 (Kopf Instruments, model: 1474)
Stereotaxic drill holder (Kopf Instruments, holder included with the drill model: 1474)
Hamilton 10 μL syringe (World Precision Instruments, catalog number: NANOFIL)
NanoFil 35 G bevel needle tip (World Precision Instruments, catalog number: NF35BV-2)
Microinjection syringe pump (World Precision Instruments, catalog number: UMP3T-1)
Pipette 2 μL (Rainin, catalog number: 17014393) and tips (Gilson, catalog number: F171100)
Holder for the optic fiber implant (RWD, catalog number: 68210)
Timer
Recovery cage
Recording
Insulated recording chambers (ViewPoint Behavior Technology)
16-channel AC amplifier (National Instruments, model: 3500)
Multifunction I/O DAQ device (National Instruments, model: USB-6343)
Fiber photometry system: RZ10-X Lux-I/O Processor (Tucker Davis Technologies) with integrated LED drivers, integrated excitation LEDs (405, 465, and 560 nm, LUX LEDs, Tucker Davis Technologies, Lx405, Lx465, and Lx560 respectively) and integrated LUX photosensors (Tucker Davis Technologies, LxPS1)
Digital handheld optical power meter (Thorlabs, model: PM100D)
Four-port Minicubes [Doric, Ordering code: ilFMC4-G2_IE(400-410)_E(460-490)_F(500-550)_S] per animal
Software
Synapse (Tucker-Davis Technologies)
Sleepscore (ViewPoint Behavior Technology)
MATLAB (Mathworks)
Procedure
The procedures below describe how we perform the experiments, but researchers should feel free to adjust these according to their needs and local guidelines.
Electrode and patch cord preparation for EEG and EMG
Turn on the soldering iron and heat it up to 450 °C.
Clean the tip of the soldering iron with a brass sponge from the soldering station.
Break off the breakaway female header pins in the amount and size you need. Here, we use two pins per electrode (Figure 1A).
Figure 1. Parts for electroencephalographic (EEG) and electromyographic (EMG) fabrication. Breakaway female header pins before (A) and after (B) male pins are shortened. C. Two pieces of EMG wire cut to approximately 1 cm. D. Prefabricated EEG screws with attached wire.
Use the end cutting pliers to cut the tips off the female header pins to allow for a wider contact surface for soldering (Figure 1B).
Prepare EEG electrode:
Take out the number of prefabricated EEG screws you need for EEG electrodes and check that the wire is properly attached to each screw by pulling lightly on it before using them.
Cut the wire approximately 1 cm from the screw (Figure 1D).
Uncoat the tip (~1 mm) of the cut wire using a scalpel or a side-cutting plier to remove the insulation and check that the tip is properly exposed under the microscope.
Use a clamp to fixate an unused row of female headers. This will create a holder for the newly cut female header pins (Figure 2).
Figure 2. Electrode holder. A row of breakaway female header pins can serve as a holder for the electrode during soldering.
Stick your cut pins onto the holder.
Using forceps, hold the uncoated wire tip against the male pin and, with the other hand, use the heated soldering iron tip to transfer a small droplet of solder tin to the male pin, to solder the EEG wire to the pin.
Let the solder material harden before you let go of the wire. You can blow lightly on the solder material to make it harden faster and check that soldering was successful by pulling gently on the wire.
Repeat steps A1f–A1g for the other screw and be careful such that the soldering material does not connect to the first pin.
When both pins have EEG screws attached, remove the electrode from the holder.
Check the conductance from each screw to their respective female plug using a multimeter. The multimeter can also be used to check if the two pins have been short-circuited.
Note: If there is no conductance, redo the soldering and test it again.
When the electrode is working, use the super glue to insulate the electrode. Cover all metal parts with a thin layer of glue as shown in Figure 3.
Figure 3. Finished electroencephalographic (EEG) (left) and electromyographic (EMG) (right) electrodes
Apply the glue accelerator to dry the glue. The accelerator can be applied with a cotton swab or a plastic one-time-use pipette. Let electrodes dry before packing them away.
Prepare EMG electrode:
Cut EMG wire into approximately 1 cm long pieces (Figure 1C). You will need two pieces per electrode.
Uncoat ~1 mm in both ends of each EMG wire as described in step A5c to remove insulation.
Attach the two EMG wires to each pair of female header pins as described in steps A5d–A5i.
Check the conductance from each EMG wire tip to their respective female plugs using the multimeter, and redo if there is no connection.
When the electrode is working, cover the soldered area with glue as described in steps A5k–A5l (Figure 3).
Prepare EEG/EMG patch cord:
Shorten the 6-channel cable to a length appropriate for your EEG setup by cutting the end with no connector. The wire should be just long enough to reach all corners of the chamber.
Cut a ~15 cm piece of the metal braided sleeve and pull it over the 6-channel cable. Secure the ends of the cut sleeve with surgical tape to prevent it from unwinding.
Remove ~30 cm of outer insulation (the thick clear plastic sleeve) from the 6-channel cable at the end with no connector using a fine scissor. This will expose the six colored cords within. Be careful not to cut the colored cords in the process.
Untwine the colored cords and cut away the strengthening yarn.
Check that all the cords are of equal length, trim them if necessary, and uncoat ~2 mm of each colored cord.
Break off two pairs of breakaway female headers and leave the male pins intact, since these will serve as connectors to the electrodes.
Work out which channels will be used for EMG, EEG, and reference, and pair the cords accordingly. This will depend on the EEG setup and software settings.
Solder the pair of EMG cords to the female ends of a breakaway female header pair and do the same for the pair of EEG cords. The two unused channels can either be left open ended or soldered to another pair of breakaway female headers.
Check the conductance from each breakaway female header pin to their respective pin on the 6-channel cable connector. If there is no conductance, redo the soldering and test it again.
Cover the soldered connections with glue as described in steps A5k–A5l.
Slide the metal braided sleeve down to the electrode connectors and secure it with surgical tape ~2 cm from the electrode connectors. The unused cords can be taped against the outside of the sleeve (Figure 4).
Figure 4. Electroencephalographic (EEG) patch cord. Patch cord with two connectors for EEG and electromyographic (EMG). The excess wires (here, blue and green) are taped against the metal braided sleeve.
Surgery
Turn on all equipment (Figure 5A). Set both heating pads to 37.2 °C. Disinfect surgical table and tools with ethanol. Prepare analgesic and local anesthetic solutions (buprenorphine, lidocaine, and carprofen). Wrap the small heating pad with KIMTECH® wipes in a single layer to make sure heat can travel from the pad to the mouse.
Note: If proposed analgesic and local anesthetic solutions do not comply with your laboratory and state regulations, use the compliant ones.
Figure 5. Surgery setup. A. Equipment placed on the surgery table. Surgery microscope and gooseneck lights are positioned for optimal visibility and flexibility. B. Close-up of implant holder with optical implant clasped.
Place the mouse in the isoflurane induction chamber and anesthetize the mouse with 5% isoflurane.
Note: The mouse should be deeply anesthetized, as apparent by a deep breathing pattern.
Once the mouse is anesthetized, apply clean air to the induction chamber to remove isoflurane before opening the chamber. Transfer the mouse to the stereotactic frame and secure its head with a nose cone. Ensure the tongue is sticking out of the mouth before fastening the nose cone.
Lower the isoflurane to support anesthesia and administer preoperative analgesia subcutaneously (e.g., buprenorphine dose: 0.05 mg/kg). Make sure the stereotactic frame is placed on a surface with appropriate ventilation to reduce any risk of exposure to isoflurane for the experimenter.
Note: Monitor the breathing pattern of the mouse (~55–65 breaths per minute, no gasping) every 15th min during surgery and adjust isoflurane accordingly.
Apply eye drops and re-apply whenever necessary to keep eyes moist at all times during surgery.
Prepare surgical field.
Shave the mouse’s head (pay attention not to cut whiskers) and secure the ear bars.
Disinfect the shaved skin three times, alternating between iodine and 70% EtOH using cotton swabs. Be careful to go from the center and out.
Inject local anesthetic (e.g., lidocaine) subcutaneously at incision site and check reflexes using toe pinch.
Using blunt forceps, touch the scalp to make sure that it is properly fixed.
Remove a small piece of skin from the scalp using forceps and scissors so the scalp is exposed and ensure both bregma and lambda can be seen (Figure 6).
Soak the cotton swab in ethanol and wipe the skull to move the skin to the sides and dry up the skull.
Apply Green Activator to the skull with a cotton tip for better adherence of the subsequently used dental cement.
Align the skull for precise injection and implantation.
Mark up bregma and lambda with a surgical marker pen according to guidelines in your reference atlas (Paxinos and Franklin, 2001) (Figure 6).
Roughly align the head by eyesight.
Fix the electric drill and use the drill tip to guide the precise alignment of the skull. Bregma and lambda should display no more than 0.05 mm differences in height (dorsoventral, D/V coordinates). Mediolateral coordinates ± 2 mm from bregma should be aligned to display no more than 0.05 mm height differences (Figure 6).
Note: Always loosen the nose cone before adjusting the head and remember to fasten again when done.
Figure 6. Skull reference points. Bregma (b) is located where the sagittal suture meets the coronal suture, and lambda (l) is defined as the point of intersection of tangent lines through the sagittal and lambdoid sutures. 2 mm on either side of bregma (indicated by the two X) is used for sideways alignment of the skull.
Proceed with drilling holes for the optic fiber implant and EEG electrode.
Optic fiber implant hole: using a drill holder, move the drill tip to bregma and zero before moving to the coordinate of interest and make a hole with the drill. After drilling, carefully probe the hole with a needle to ensure the hole was made, to avoid bending the NanoFil needle upon injection.
EEG electrode hole: plan where in the skull EEG electrodes should be inserted. As a minimum, you need two screws: one for signal recording and one as reference. The reference screw is usually placed over cerebellum, which is electrically neutral. Make sure to place screws so you have room for other implants, e.g., optic fiber implants. Drill one hole for each EEG screw using a drill tip to match the diameter of the EEG screws.
Implant EEG and EMG electrodes.
Clean the EEG and EMG electrodes in ethanol and dry them off. If needed, straighten and trim the wire tips of the EMG electrode and remember to leave the tip uncoated. This will ease insertion into the muscle fascia.
Place the EEG electrode screws in designated holes (Figure 7A).
i. Hold the screw in place using sterilized forceps and screw it in with a small screwdriver without using too much force. Grip the screwdriver closer to its tip for better control. Do the same for all screws and, if necessary, untwine the wire once in a while. The screws are appropriately placed if they cannot be easily moved from their designated holes; at the same time, they should not enter the brain tissue.
ii. Place the EEG electrode connector to the side.
Place the EMG electrode in the trapezius muscle (Figure 7B).
i. Hold fine forceps in your dominant hand and blunt forceps in the other hand with the electrode within reach.
ii. Pull out the cut skin at the base of the skull using the blunt forceps, poke a hole in the muscle fascia with the fine forceps, and keep it in place as a guide for the EMG electrode.
iii. Grab one of the wires of the EMG electrode with the blunt forceps and guide it along the fine forceps through the hole in the fascia and into the muscle.
iv. Repeat this with the other wire of the EMG electrode.
Figure 7. Electrode insertion. Image showing how the surgical field looks after insertion of electroencephalographic (EEG) (A) and electromyographic (EMG) (B) electrodes.
Inject viral probe using Hamilton micro syringe.
Test if the micro syringe is not clogged by withdrawing/ejecting distilled water from it.
Mount the micro syringe on the holder, place the needle tip at bregma, and zero the coordinates. Place the needle tip over the target coordinate and test if the needle can go through the pre-drilled hole.
Set the infusion rate of the infusion pump to 100 nL/s.
Place a small rectangle of parafilm on top of the skull and put a drop (~1.2 μL) of virus on the parafilm using a pipette (assuming that 300–500 nL of virus will be injected per region).
Replace the default needle on the 10 μL syringe with a 35 G bevel needle tip.
Place the needle above the drop and withdraw 200 nL of air. Check if the plunger is moving.
Then, place the needle tip in the drop and withdraw virus (withdraw ~300 nL more than needed for the infusion itself). Using the microscope, ensure the drop on the parafilm is getting smaller during withdrawal. There should be a small volume of virus left on the parafilm after withdrawal is finalized to make sure no air was withdrawn in the end.
Note: If the withdrawal was not successful, infuse everything back onto the parafilm and try again. The needle should be replaced if problems persist.
If the withdrawal was successful, remove the parafilm and place the needle above the site of injection. Make a test drop by infusing 50 nL and look at the needle through the microscope to see whether a drop forms at the tip. Keep track of how much virus should be left in the needle.
Note: If there is still no drop formed after the volume of virus needed for injection is reached, infuse everything back onto the parafilm and start over from step B10a.
When the test infusion drop is successful, adjust the infusion rate to 100 nL/min and lower the needle to the D/V coordinate. Inject the virus.
Note: Depending on the depth of the target D/V coordinate, you can perform 1–4× injections at different depths (150–200 nL per depth location) to ensure appropriate viral expression. E.g., if you perform three injections, firstly lower the needle to 0.2 mm above the target D/V coordinate and inject one third of the total volume of virus. Repeat this at the target D/V coordinate and 0.2 mm below the target D/V coordinate.
When the last injection is finished, leave the needle in for at least 7 min to allow for diffusion of the virus. Use the timer to keep track of time.
Slowly, pull out the needle and clean the Hamilton at least 10 times with distilled water.
If several optic fiber implants are implanted, make sure all holes (step B8a) and virus injections (step B10) are done before starting any optical implant inserts. In addition, plan out the order of implantations to ensure that bregma is not covered until the last implant is in place.
Place an optic fiber implant at the target coordinate.
Replace the micro syringe holder on the stereotax with the holder for the optic fiber implant.
Place an optic fiber in the holder (Figure 5B). Be careful not to touch the fiber tip.
Make sure the skull is completely clean and dry for better adherence to dental cement.
Place the fiber tip at bregma very carefully not to scratch the tip, and zero the coordinates.
Go to the target coordinates and insert the fiber slowly into the hole at the desired depth. Make sure that it fits the hole and that no bleeding occurs at insertion.
Note: If bleeding occurs, pause insertion and clean off the blood using a KIMTECH® wipe before continuing.
Arrange EEG and EMG connectors before using dental cement for fixation. Ensure optic fiber implant and EEG/EMG connectors are appropriately distanced. See Figure 8 for final result.
Note: It is important to place optic fiber implant and EEG/EMG electrodes in a way that will ensure all of them can be easily plugged in during subsequent behavioral experiments.
Mix dental cement as instructed on the package and apply it on the skull starting from the optic fiber implant.
Note: Ensure that approximately 5 mm of the ferrule top of the optic fiber implant is free of cement as this will be used to connect the fiber-optic patch cord.
Wait for the cement to harden for approximately 10 min or until it is completely solid before loosening and removing the fiber holder from the stereotax.
Note: If several optic fiber implants are implanted, make sure not to cover bregma with dental cement if needed for zeroing of subsequent implants.
Apply more layers of dental cement as needed to cover the entire skull and EEG and EMG wires and to stabilize all connectors (Figure 8). This requires several rounds of dental cement fabrication. Avoid covering the skin with dental cement and avoid sharp edges.
Figure 8. Completed implant. Image shows an example of how the implant looks upon completion.
Administer postoperative analgesia subcutaneously (e.g., carprofen dose: 5 mg/kg) and saline to prevent dehydration and lower the isoflurane to speed up recovery.
When the cement is completely hardened, gently pull the incised skin to cover the edge of the dental cement and turn off isoflurane.
Place the mouse on a heating pad in the recovery cage until the mouse is up and moving around.
After recovery, place the mouse in its home cage. It should be group-housed to avoid stress.
Wet some food pellets and place them in the home cage of the mouse. Remove the grid permanently from the home cage to ensure implants do not get bumped or stuck in the grid. Use the filter top as lid and place food pellets on the bottom of the cage.
Administer postoperative analgesia according to local guidelines in the following days after the surgery.
Recording
Before recordings (three weeks)
Allow 2–3 weeks between surgery and fiber photometry experiments to ensure recovery and adequate virus expression.
Recommended: habituate the mouse to the experimenter. To minimize stress during the experiment, gently handle the mouse for five days before the experiment and practice light scruffing. Avoid tail-lifting mice; instead, scoop them up using hands or a plastic cylinder.
One day before recordings
Prepare fiber photometry protocol:
i. In the fiber photometry recording software, add channels for the excitation wavelength and the isosbestic point of the fluorescent biosensor of interest. The isosbestic point of the biosensor serves as an autofluorescence detection control and may differ between biosensors. Information on excitation wavelength and isosbestic control can usually be found in the original publication characterizing the biosensor. For the norepinephrine biosensor, we use the 465 nm excitation wavelength and the 405 nm isosbestic point, which are also commonly used for GCaMP constructs.
ii. Modulate each LED at different frequencies that are not multiple of each other or the power line hum (i.e., electrical noise, typically at 50 or 60 Hz). We usually use 211, 330, and 531 Hz.
iii. Using a power meter, measure the wavelength output at the tip of the patch cord and adjust the LED power to get an output within 10–30 μW.
iv. In the fiber photometry recording software, create a channel that generates TTL pulses for alignment of the fiber photometry and EEG/EMG recordings and connect the two setups accordingly.
Note: This may require a custom-made channel in the EEG setup, which the vendor can help you achieve.
Prepare EEG chambers: turn on EEG equipment and add white bedding and nesting material on chamber floor for C57Bl6 mice to increase contrast in video recording. Avoid using excessive nesting material and cut it to approximately 5 cm long strings so that the animal is not entangled or hidden while sleeping. Add food pellets to the floor of the EEG chamber and supply a water bottle through a hole in the cylinder.
Test that the virus expression is adequate:
i. Place the mouse on the metal grid of a cage and scruff it very lightly. Attach a zirconia sleeve to the fiber-optic patch cord and connect it to the metal ferrule of the optical implant by sliding the sleeve over the ferrule. Do not push the fiber-optic patch cord on the optical probe but gently place it on by using a screw-on motion instead (Video 1). Check through the slid on the sleeve that the two ferrule ends meet.
Video 1. Connecting fiber-optic patch cord
ii. Preview the signal in the fiber photometry recording software and make sure that no light is escaping the connection between the fiber-optic patch cord and the optical probe (Figure 9).
Figure 9. Connecting fiber-optic patch cord. A. Example of patch cord connected properly with the two metal ferrule ends touching one another. B. Example showing inadequate connection of patch cord revealed by light escaping from the side.
iii. Subsequently, place the animal in its home cage and allow it to rest for 5 min.
iv. Following this, perform a swift tail lift or another type of startle: most biosensor signal rises due to the acute stress of the tail lift. If no signal occurs, the animal likely has poor expression of sensor and might not be worth running.
v. Remove the optic fiber (see step C3e below).
Habituate the mouse to EEG chambers and EEG/EMG patch cords over one active phase to ensure natural sleep in the following inactive phase. Connect EEG and EMG patch cords to the corresponding implants, while scruffing the animal lightly. Ensure that the EEG reference patch cord is connected to the screw that was placed over cerebellum.
Check EEG and EMG signals and test that the custom-made TTL channel receives input from the fiber photometry setup.
Bleach the fiber-optic patch cord by turning on the excitation LEDs at maximum power for 2 h. Make sure the animal is disconnected from the patch cord during bleaching.
On the day of the recordings:
Turn on fiber photometry recording equipment, select protocol, and test light responsivity by holding the tip of the patch cord to a white material to ensure that fluorescence is detected. When testing on white material, all channels should respond. If no signal appears, check that your equipment is appropriately connected and turned on.
Turn on EEG setups and, on the amplifier, select filtering of 0.3HP and 100LP.
Connect the fiber-optic patch cords to corresponding implants as described above (step C2ci) and check that EEG and EMG patch cords are still properly connected after habituation.
Start recording in both EEG/EMG and fiber photometry setups. Remember to start the TTL pulse for the synchronization of the systems (this should be done after recordings are started for both EEG/EMG and fiber photometry). Record during the light phase to ensure natural sleep and consider circadian timing when planning experiments. If no combined EEG/EMG/fiber optic rotary joint is added to the setup, the recording should not exceed 5–6 h as the animal will have limited capacity for rotation.
After the recordings are finished, disconnect the patch cords from the animal. Place the animal on the metal grid and remove EEG and EMG patch cords first, followed by the fiber-optic patch cords. There is no need to scruff the animal for this procedure; gently remove patch cords by lightly holding on to the rest of the implants and applying minimal resistance. The fiber-optic patch cord can be removed with a screw-off motion (Video 2).
Video 2. Disconnecting fiber-optic patch cord
Place the animal in the home cage and turn off the equipment if no other recordings ensue.
Data analysis
Sleep scoring
Open the EEG and EMG raw data in a scoring software. We use Sleepscore (ViewPoint Behavior Technology), but any equivalent software can be used; preferably a software that allows for:
1-s scoring windows for higher temporal resolution around transitions.
Monitoring simultaneous video recording.
Fast-Fourier Transform (FFT) of the EEG scoring window as an extra visual aid to determine frequency band power.
Do an initial quick scoring of the entire recording using a 5 s window. Wakefulness is characterized by high EMG activity and low EEG amplitude (Figure 10). Non-rapid eye movement (NREM) sleep is characterized by low EMG activity and high EEG amplitude, especially dominated by high delta (0.5–4 Hz) power. Rapid eye movement (REM) sleep is characterized by low EMG activity and low EEG amplitude with a relative increase in theta (4–8 Hz) and a decrease in delta activity compared to NREM sleep.
Figure 10. Electroencephalographic (EEG) and electromyographic (EMG) traces across brain states. Example of an EMG trace (top) and EMG trace (bottom) during wake and NREM and REM sleep.
Change the scoring window to 1 s and go through your scoring once more, paying special attention to transitions to score them more precisely. Use the video to help distinguish EMG noise from actual movement to help score short awakenings, which will later be classified as micro-arousals.
Extract the onsets and durations for each state from the scored hypnogram data in a format that can be loaded into MATLAB.
Fiber photometry analysis and EEG analysis
Extract the fiber photometry data (465 nm channel, 405 nm channel, sampling frequency, and the TTL timestamps), load them into MATLAB, and open the script provided on GitHub (https://github.com/MieAndersen/Bio-Protocol_FibPho_NE) to perform the analysis steps below.
Cut fiber photometry traces before the first TTL timestamp for alignment with EEG recording.
Calculate dF/F of the 465 nm channel: in short, fit the 405 nm signal to the 465 nm signal using a first-degree polynomial fit to remove signal drift present in both signals due to bleaching (Figure 11, top). Calculate dF/F by subtracting the fitted 405 signal from the 465 signal and divide it by the fitted 405 signal (Figure 11, middle). Lastly, filter the dF/F signal using a zero-phase digital filter to remove noise (Figure 11, bottom).
Figure 11. Normalization of fluorescent traces. Example showing the 405 nm control signal (red) fitted to the raw 465 nm signal (blue, top), the normalized dF/F signal (middle), and the filtered dF/F signal (bottom).
Load the EEG data (EEG channel, EMG channel, sampling frequency, and TTL time stamps) into MATLAB.
Load scored onsets and durations into MATLAB and create binary vectors for each state that will be used for plotting and subsequent alignment with fiber photometry data.
Divide wake bouts into micro-arousals < 15 s and awakening excluding micro-arousals (≥ 15 s).
To align with fiber photometry data, cut the EEG data and scoring before the first TTL pulse sent from the fiber photometry setup and plot scoring and traces to check alignment.
Next, you can extract event-related epoch traces if you are interested in certain events such as state transitions.
Acknowledgments
The protocol described in this paper was developed in the making of Kjaerby et al. (2022).
Funding: Lundbeck Foundation, R386–2021–165; Independent Research Council Denmark, 7016–00324A; Augustinus Foundation, 16–3735.; Novo Nordisk Foundation, NNF20OC0066419.
We thank Myles Billard and Palle Koch for technical support.
Competing interests
We declare no competing interests.
Ethical considerations
All experiments were approved by the Danish Animal Experiments Inspectorate and were overseen by the University of Copenhagen Institutional Animal Care and Use Committee, in compliance with the European Communities Council Directive of 22 September 2010 (2010/63/EU) legislation governing the protection of animals used for scientific purposes.
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Neuroscience > Basic technology > Electromyography
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The authors used this protocol in eLIFE Jul 2022
Abstract
T cells are endowed with T-cell antigen receptors (TCR) that give them the capacity to recognize specific antigens and mount antigen-specific adaptive immune responses. Because TCR sequences are distinct in each naïve T cell, they serve as molecular barcodes to track T cells with clonal relatedness and shared antigen specificity through proliferation, differentiation, and migration. Single-cell RNA sequencing provides coupled information of TCR sequence and transcriptional state in individual cells, enabling T-cell clonotype-specific analyses. In this protocol, we outline a computational workflow to perform T-cell states and clonal analysis from scRNA-seq data based on the R packages Seurat, ProjecTILs, and scRepertoire. Given a scRNA-seq T-cell dataset with TCR sequence information, cell states are automatically annotated by reference projection using the ProjecTILs method. TCR information is used to track individual clonotypes, assess their clonal expansion, proliferation rates, bias towards specific differentiation states, and the clonal overlap between T-cell subtypes. We provide fully reproducible R code to conduct these analyses and generate useful visualizations that can be adapted for the needs of the protocol user.
Key features
• Computational analysis of paired scRNA-seq and scTCR-seq data
• Characterizing T-cell functional state by reference-based analysis using ProjecTILs
• Exploring T-cell clonal structure using scRepertoire
• Linking T-cell clonality to transcriptomic state to study relationships between clonal expansion and functional phenotype
Graphical overview
Keywords: Single-cell analysis T-cell receptor TCR Transcriptomics T-cell clone Reference projection scRNA-seq scTCR-seq
Background
T cells are crucial players in the adaptive immune response with the capacity to recognize and eliminate infected and malignant cells. The antigen specificity of T cells is conferred by their T-cell receptors (TCRs). These are heterodimeric proteins in which each of the two protein chains—typically one alpha (α) and one beta (β)—is produced through somatic rearrangement of V, (D), and J gene segments, as well as the addition or deletion of nucleotides between spliced gene segments, to form a unique V(D)J exon. This recombination process is largely random and generates a large repertoire of TCRs, with an estimated diversity in the order of 108–1010 unique TCR chains in a single individual (Qi et al., 2014; Lythe et al., 2016). Such a large repertoire, and in particular the hypervariable complementary-determining region 3 (CDR3) of the TCR, allows recognizing with high specificity a vast array of foreign antigens, while maintaining tolerance to self-antigens. Due to this huge diversity, each naïve αβ T cell has a virtually unique combination of TCR α and β chains. Thus, paired αβ TCR sequences serve as molecular barcodes to track T-cell clones through processes such as proliferation, differentiation, and migration.
The emergence of single-cell technologies has enabled the coupled sequencing of full-length TCRs with transcriptome-wide RNA sequencing in individual cells (Pai and Satpathy, 2021). This is achieved by either computationally reconstructing TCR chain transcript sequences from single-cell RNA sequencing reads (Eltahla et al., 2016; Stubbington et al., 2016; Bolotin et al., 2017) or by specifically amplifying the TCR locus, also known as single-cell TCR-seq (e.g., using Chromium 5′ V(D)J sequencing). Coupled with scTCR-seq, scRNA-seq enables the connection between T-cell states, clonotypes, and potential antigen specificity (Han et al., 2014).
We have recently proposed computational pipelines for the analysis of single-cell T-cell repertoires (Borcherding and Bormann, 2020) and for reference-based analysis of single-cell transcriptomics data (Andreatta et al., 2021a), based on the tools scRepertoire and ProjecTILs, respectively. In this protocol, we describe how to combine these computational tools to analyze paired scRNA-seq and scTCR-seq data to track individual clonotypes projected in a reference map, assess their clonal expansion, proliferation rates, their bias towards specific differentiation states, and the clonal overlap between T-cell states. We will focus the examples on human CD8+ T cells from tumor biopsies, but the protocol is applicable to any single-cell transcriptomics data with TCR sequence information in humans and mice. We invite the reader to follow this protocol while interactively running the associated R Notebook (see Software and datasets section).
Equipment
Personal computer (minimum 16 GB of RAM) or high-performance computing cluster. All software runs on Linux, Windows, or MacOS machines.
Software and datasets
This protocol requires basic R programming skills: installing packages, running an R notebook, and adapting the code to the needs of the user.
All software used for this protocol is free and open source.
R version 4.2 or higher
scRepertoire (version ≥ 1.7) (Borcherding and Bormann, 2020) (https://github.com/ncborcherding/scRepertoire)
ProjecTILs (version ≥ 3.0) (Andreatta et al., 2021a) (https://github.com/carmonalab/ProjecTILs)
Seurat (version ≥ 4.3) (Hao et al., 2021) (https://github.com/satijalab/seurat)
In addition, it is recommended to install R Studio Desktop (https://posit.co/downloads/) to interactively run the R Notebook that reproduces the results of this protocol (https://github.com/carmonalab/Tcell_clonal_analysis).
To download the repository to your machine, run the following from command line:
git clone [email protected]:carmonalab/Tcell_clonal_analysis.git
Then, move to the newly created directory and open the project file (with .Rproj extension). Open the protocol notebook (protocol_CD8TIL_clonalAnalysis.Rmd) in R Studio and execute all commands in order. Note that the R Notebook makes use of the renv package (https://rstudio.github.io/renv/articles/renv.html) for straightforward installation of all required packages with the correct version and to ensure reproducibility of the results shown in this protocol.
The protocol assumes the user has generated a single-cell transcriptomics dataset with TCR sequencing information for the same T cells or a subset thereof. There is no restriction on the sequencing technology used, if it generates i) a count matrix quantifying gene expression in single cells; and ii) TCR sequences, for paired αβ chains or single chains, with barcodes that can be mapped to transcriptomics measurements of the same cells.
Procedure
The protocol details all steps required to go from scRNA-seq and scTCR-seq count matrices to T-cell clonal analysis in the context of a T-cell reference map. Each step includes example code snippets that highlight the R commands that accomplish the step. For the complete list of R commands that reproduce the results of this protocol, refer to the accompanying R Notebook (see Software and datasets section).
Single-cell data pre-processing
scRNA-seq data
Several protocols and technologies are available for transcriptomics quantification using scRNA-seq. Sequencing protocols differ in terms of library preparation, read alignment to a reference genome, and quantification of transcripts, as reviewed in multiple publications (Vieth et al., 2019; Mereu et al., 2020). Sequencing facilities commonly offer read mapping and gene expression quantification to obtain a raw count expression matrix (for instance, using the Cell Ranger pipeline from 10× Genomics). From a raw counts matrix, generate a Seurat (Hao et al., 2021) object to store the counts:
seurat <- CreateSeuratObject(counts = matrix)
Note that Seurat also implements functions to load data from specific technologies, for example the Read10X() function to read count matrices from the popular 10× sequencing platform (see https://satijalab.org/seurat/reference/read10x).
scTCR-seq data
Obtaining single-cell TCR sequences requires specific protocols for amplification and sequencing of the V(D)J locus, or their reconstruction from whole-transcriptome sequencing. For an overview of scTCR-seq sequencing approaches, see the comprehensive review by Pai and Satpathy (2021).
We assume that the user has performed V(D)J sequences assembly and clonotype calling. For 10× Chromium 5′ V(D)J libraries, such annotated V(D)J sequences (“contigs”) are obtained from FASTQ files using the Cell Ranger V(D)J pipeline (https://support.10xgenomics.com/single-cell-vdj/software/pipelines/latest/using/vdj). scRepertoire (Borcherding and Bormann, 2020) implements useful functions to process the V(D)J contigs annotation files generated by Cell Ranger. These files (usually named all_contig_annotations.csv for total, or filtered_contig_annotations.csv for high-confidence filtered contigs) contain detailed information for each V(D)J contig, including its cell barcode, length, V-D-J-C segments, the number of reads and distinct UMIs aligned to the contig, and a clonotype ID to which the contig was assigned.
Load TCR α and β chains from Cell Ranger output files and combine them by cell barcode, using function combineTCR() from scRepertoire:
S1 <- read.csv("Sample1/outs/filtered_contig_annotations.csv")
S2 <- read.csv("Sample2/outs/filtered_contig_annotations.csv")
contig_list <- list(S1, S2)
combined <- combineTCR(contig_list, cells ="T-AB")
For V(D)J contigs generated using different pipelines, please see the loadContigs() function from scRepertoire, which allows data pre-processing for multiple formats including TRUST4, BD Rhapsody, WAT3R, and AIRR.
Note 1: It is often useful for further processing steps to generate keys for unique clonotype–sample combinations. As it may occur by chance that the same clonotype is observed in different individuals, these keys will allow discriminating between T cells with identical TCR but from different samples. For example, generate a clonotype–sample key as a metadata column named “cdr3s_pat”:
combined <- lapply(combined, function(x){x$cdr3s_pat <- paste(x$CTaa, x$sample, sep="_"); x})
Combine scRNA-seq and scTCR-seq data
Append the TCR information into the previously prepared Seurat object that stores the scRNA-seq counts. If the V(D)J data were processed using combineTCR() from scRepertoire, you may apply the combineExpression() function:
seurat <- combineExpression(combined, seurat,
group.by = "sample",
cloneTypes=c(Single=1, Small=5, Medium=10, Large=20, Hyperexpanded=50))
For V(D)J data pre-processed using different pipelines, add the TCR chains as metadata to the Seurat object:
seurat <- AddMetaData(seurat, tcr.chains)
Reference-based analysis
Load reference map
Several reference single-cell maps for reference-based analysis are available from the ProjecTILs repository (https://github.com/carmonalab/ProjecTILs) and from SPICA (Andreatta et al., 2021b) (https://spica.unil.ch). For example, to analyze human CD8+ T cells, download and load the corresponding map (Figure 1):
ref.file <- "CD8T_human_ref_v1.rds"
download.file("https://figshare.com/ndownloader/files/38921366", destfile = ref.file)
ref.cd8 <- load.reference.map("CD8T_human_ref_v1.rds")
DimPlot(ref.cd8, cols = ref.cd8@misc$atlas.palette)
Figure 1. Reference map for human CD8+ T cells
Project data into the reference
To embed query data into the reference space and obtain cell type annotations, apply the ProjecTILs pipeline (Andreatta et al., 2021a). If the query dataset is composed of different samples (e.g., from different patients or time points), we recommend splitting it and projecting each sample separately into the reference. In this way, ProjecTILs will assume that each sample represents a different experimental batch and will calculate and correct batch effects accordingly:
seurat.list <- SplitObject(seurat, split.by = "patient")
seurat.projected <- Run.ProjecTILs(seurat.list, ref.cd8)
In this case, the output is a list of Seurat objects, each corresponding to a query sample projected in the reference map.
Note 2: For this example, because we chose to use a CD8+ T cell reference map, ProjecTILs will automatically pre-filter CD8+ T cells from the input data (i.e., will remove CD4+ T cells and non-T cells). With ProjecTILs, it is also possible to conduct multi-reference map analysis, for instance using both CD8+ T cells and CD4+ T cells reference maps. An example can be found in the following R notebook: https://carmonalab.github.io/ProjecTILs_CaseStudies/Bassez_BC.html.
Compare marker gene expression profiles of query data with the reference map
To verify the correspondence of transcriptional phenotypes between the reference and query dataset, visualize the average expression profile of each cell subtype for a panel of marker genes (Figure 2):
which.patient <- "su009"
plot.states.radar(ref.cd8, seurat.projected[[which.patient]], genes4radar = genes4radar)
Figure 2. Expression profiles for reference and query dataset for a panel of marker genes. Reference is a human CD8+ T-cell reference (see text); the query is a representative individual (“su009”) from a cohort of basal cell carcinoma (BCC) patients (Yost et al., 2019).
Cell subtype composition of query data
Reference projection of the query data allows embedding them into the same space of the reference. Cell types for the query dataset can be predicted by nearest-neighbor majority voting based on the annotated reference cells. Visualize low-dimensional embeddings and subtype composition for individual samples or other subsets of the projected data (Figure 3):
which.patient <- "su009"
a <- plot.projection(ref.cd8, seurat.projected[[which.patient]])
b <- plot.statepred.composition(ref.cd8, query = seurat.projected[[which.patient]])
a | b
Figure 3. Reference embeddings and cell subtype composition for query dataset. Reference is a human CD8+ T-cell reference (see text); query is a representative patient (“su009”) from a basal cell carcinoma (BCC) cohort (Yost et al., 2019).
Exclude small samples
Robust analyses require a minimum number of cells in each sample. After projection and annotation, remove all samples with a small number of cells (e.g., 100 cells):
sizes <- as.vector(lapply(seurat.projected, ncol))
keep <- names(sizes)[sizes > 100]
seurat.projected <- seurat.projected[keep]
For large enough samples, we can compare their composition in terms of cell subtypes (Figure 4):
plots <- lapply(names(seurat.projected), function(x) {
plot.statepred.composition(ref.cd8, query = seurat.projected[[x]],
metric = "Percent") + ggtitle(x)
})
wrap_plots(plots, ncol=4)
Merge list of objects to obtain a single object
For some analyses (including clonal analysis detailed below), it is useful to merge individual objects/samples (projected by patient) into a single object:
merged.projected <- Reduce(merge.Seurat.embeddings, seurat.projected)
Idents(merged.projected) <- "functional.cluster"
Figure 4. CD8+ T-cell subtype composition in individual tumor biopsies with at least 100 CD8+ T cells. Data are from basal cell carcinoma (BCC) tumor biopsies (Yost et al., 2019); plots are sorted by the fraction of CD8.TEX cells.
Clonal analysis
If the TCR information was loaded into the query Seurat object as outlined in section A, it will be available as metadata for the projected object. This allows linking the transcriptomics state to clonal information. A few examples of analyses are detailed below.
Identify the most expanded clones
Calculate the frequency of unique TCR chains per patient (e.g., as stored in “cdr3s_pat” metadata, see Note 1) to identify the most expanded clones per patient:
freqs <- lapply(seurat.projected, function(x) {
table(x$cdr3s_pat) / sum(!is.na(x$cdr3s_pat))
})
freqs <- Reduce(c, freqs)
sorted <- sort(freqs, decreasing = TRUE)
largest.clones <- head(sorted, 6)
Locate expanded clones on the reference low-dimensional space
TCR chains can be used to subset clones of interest (e.g., the largest clones as identified above) and inspect their distribution on the reference UMAP space (Figure 5):
plots <- list()
for (i in 1:length(largest.clones)) {
ctype <- names(largest.clones)[i]
cells <- which(merged.projected[["cdr3s_pat"]]==ctype)
plots[[i]] <- plot.projection(ref.cd8, merged.projected[,cells])
}
wrap_plots(plots, ncol = 3)
Figure 5. Reference UMAP embeddings highlighting with contour lines the location of the six most expanded CD8+ T-cell clones in basal cell carcinoma (BCC) tumor biopsies (Yost et al., 2019)
Clonal expansion by T-cell subtype
scRepertoire implements several useful functions to visualize clonal expansion and clonal diversity. Plot the number of cells in different categories of expansions, from “Single” clones to large clones (here >50 cells), by T-cell subtype (Figure 6):
occupiedscRepertoire(merged.projected, x.axis = "functional.cluster")
Figure 6. Occupied single-cell repertoire by cell subtype. The y-axis indicates the absolute number of cells, and colors identify the type of clone based on the number of cells it contains. Data from Yost et al. (2019).
Clonotype proliferation rate
High proliferation rate of a specific clonotype may indicate that the T cells with shared specificity are actively recognizing antigens in situ. We can measure proliferation at the clonal level by calculating how many cells of a clone are cycling, according to transcriptomics readouts. ProjecTILs automatically calculates cell cycling signature scores using UCell (Andreatta and Carmona, 2021). These signature scores can be used to define cell cycle stage and proliferative status (Figure 7):
merged.projected$is.cycling <- ifelse((merged.projected$cycling.score.G1_S > 0.1 |
merged.projected$cycling.score.G2_M > 0.1),
yes = "Proliferating",
no = "Resting")
#Only consider expanded clones
clonotypes <- table(merged.projected$cdr3s_pat)
expanded <- names(clonotypes)[clonotypes>=2]
frequency.proliferating <- sapply(expanded, function(x) {
sub <- subset(merged.projected[[]], subset=cdr3s_pat == x)
sum(sub$is.cycling == "Proliferating") / ncol(sub) })
Figure 7. Most proliferative clones in pre-treatment biopsies from a basal cell carcinoma (BCC) cohort (Yost et al., 2019). A. Fraction of proliferating cells for the six most proliferative clones. B. Reference-embedding for the same six clones. Proliferating cells are calculated based on signature scoring of the cell cycling signatures defined by Tirosh et al. (2016).
Note 3: The user may want to use different gene signatures than those automatically applied by ProjecTILs, to quantify activity of additional gene programs. We refer to the UCell online documentation for interacting with Seurat objects and for custom gene signature scoring: https://bioconductor.org/packages/release/bioc/vignettes/UCell/inst/doc/UCell_Seurat.html.
Clonal sharing between T-cell subtypes
Metrics of clonal overlap [e.g., Horn-Morisita index (Horn, 1966)] can be used to assess clonal sharing between samples and between T-cell subtypes. Here, we analyze the clonal sharing between subtypes (Figure 8A):
clonalOverlap(combined, cloneCall = "cdr3s_pat", method = "morisita")
Several additional representations of clonal overlap are available in scRepertoire, for example as circos plots (Gu et al., 2014) (Figure 8B):
circles <- getCirclize(merged.projected,
cloneCall = "cdr3s_pat",
group.by = "functional.cluster")
circlize::chordDiagram(circles)
Figure 8. Clonal overlap between CD8+ T-cell subtypes. A. Morisita index for T-cell receptors (TCR) sharing between CD8+ T-cell subtypes. B. Circos plot visualization of clonal sharing between subtypes. Data from Yost et al. (2019).
Note 4: Cell type/state classification algorithms are not perfect, and there is generally some uncertainty in the predicted subtypes, especially among closely related subtypes (e.g., NaiveLike and CM/Central Memory). Moreover, some cells might display intermediate states of differentiation, transitioning from one state into another. These factors might lead to some background noise for TCR sharing/Morisita index between transcriptionally related cell states (e.g., in Figure 8, a Morisita index of 0.031 between NaiveLike and CM is very unlikely to be meaningful). It is strongly advised to analyze multiple independent samples to support hypotheses of TCR sharing between groups.
Clonotype bias towards specific cell states
In certain settings, it may be of interest to identify clones that are significantly composed of T cells of a certain subtype. We have previously devised a metric to measure clonotype bias and applied it to investigate if virus-specific naïve CD4+ T-cell clones were preferentially differentiating into a specific effector state, or whether multiple differentiation fates were equally likely (Andreatta et al., 2022). scRepertoire implements a function to calculate clonotype bias (Figure 9A):
clonotypeBias(merged.projected, cloneCall = "cdr3s_pat", split.by = "patient",
group.by = "functional.cluster", min.expand = 10)
Figure 9. Clonotype bias towards specific cell states. A. Clonotype bias as a function of clonal size, for expanded CD8+ T-cell clones in basal cell carcinoma (BCC) tumors (Yost et al., 2019). The blue line approximates the upper bound of a 95% confidence interval of the expected clonotype bias distribution (i.e., the clonotype bias expected by chance if there was no biological association between clonotype and cell state). B. T-cell clones with most significant clonotype bias, ranked by Z-score.
The same function can be used to return a table, by setting exportTable=TRUE, from which we can extract the most significantly biased clones according to their Z-score (Figure 9B):
biased <- clonotypeBias(merged.projected, cloneCall = "cdr3s_pat", split.by = "patient",
group.by = "functional.cluster", min.expand = 5, exportTable=TRUE)
most.biased <- biased[order(biased$Z.score, decreasing = TRUE),]
plots <- list()
for (i in 1:6) {
ctype <- most.biased[i, "Clone"]
cells <- which(merged.projected [["cdr3s_pat"]]==ctype)
title <- sprintf("Clone %s - size %s - %s", i, size, patient)
plots[[i]] <- plot.projection(ref.cd8, merged.projected [,cells])
}
wrap_plots(plots, ncol = 3)
Data analysis
Fully reproducible R code that generates the results and figures in this protocol, including all pre-processing steps, is available on GitHub: https://github.com/carmonalab/Tcell_clonal_analysis. A comprehensive vignette with more information on scRepertoire and its functions can be found at:
https://ncborcherding.github.io/vignettes/vignette.html. Several case studies of applications of ProjecTILs for reference-based analysis of single-cell data are available at: https://carmonalab.github.io/ProjecTILs_CaseStudies.
General notes and troubleshooting
Commercially available single-cell RNA-sequencing technologies have opened the opportunity to study the association of T-cell states and clonality at large scale. However, scRNA-seq experiments typically produce less than 10,000 high-quality single-cell transcriptomes per sample. Depending on the tissue analyzed, and whether or not T cells have been specifically purified, the number of sequenced T cells obtained, even from inflamed tissues, can be very low. As a result, only a small fraction of the complete TCR repertoire is typically sampled. Under-sampling leads to inaccurate estimations of clonal diversity (e.g., Shannon entropy). For this reason, in this protocol we suggest to exclude from analysis samples with very few cells and we avoided the use of clonal diversity metrics, such as Shannon entropy, Gini-Simpson index, and Gini coefficient, that are particularly sensitive to under-sampling (Chiffelle et al., 2020). Instead, we focused the analysis on the largest clonotypes in each sample. Clonal sharing between samples (e.g., Morisita index) is also affected by the low number of observations. Thus, clonal diversity and clonal sharing metrics should be interpreted with caution, and importantly, confirmed in independent samples.
Troubleshooting
Download of large objects in R (as in the case of single-cell datasets and reference maps) may occasionally fail due to connection timeout. This commonly manifests in errors such as “object X is invalid.” Try increasing download timeout using the following command within the R session:
options(timeout = max(900, getOption("timeout")))
Acknowledgments
This work was supported by the Swiss National Science Foundation (SNF project 180010 to S.J.C.).
Parts of this protocol were previously described for the study of clonal structure and clonotype-fate relationship in virus-specific CD4+ T cells (Andreatta et al., 2022).
Competing interests
N.B. is an advisor for Santa Ana Bio and Omniscope. M.A., P.G. and S.J.C. have no conflicts of interest to declare.
References
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Immunology > Immune cell function > Lymphocyte
Systems Biology > Transcriptomics > RNA-seq
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Catheterization of Pulmonary and Carotid Arteries for Concurrent Measurement of Mean Pulmonary and Systemic Arterial Pressure in Rat Models of Pulmonary Arterial Hypertension
TS Tanoy Sarkar *
AI Ayman Isbatan *
SM Sakib M. Moinuddin
JC Jiwang Chen
FA Fakhrul Ahsan
(*contributed equally to this work)
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4737 Views: 644
Reviewed by: Ling LaiJordi Boix-i-Coll Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Molecular Pharmaceutics Aug 2019
Abstract
Pulmonary hypertension (PH) is a group of pulmonary vascular disorders in which mean pulmonary arterial pressure (mPAP) becomes abnormally high because of various pathological conditions, including remodeling of the pulmonary arteries, lung and heart disorders, or congenital conditions. Various animal models, including mouse and rat models, have been used to recapitulate elevated mPAP observed in PH patients. However, the measurement and recording of mPAP and mean systemic arterial pressure (mSAP) in small animals require microsurgical procedures and a sophisticated data acquisition system. In this paper, we describe the surgical procedures for right heart catheterizations (RHC) to measure mPAP in rats. We also explain the catheterization of the carotid artery for simultaneous measurement of mPAP and mSAP using the PowerLab Data Acquisition system. We enumerate the surgical steps involved in exposing the jugular vein and the carotid artery for catheterizing these two blood vessels. We list the tools used for microsurgery in rats, describe the methods for preparing catheters, and illustrate the process for inserting the catheters in the pulmonary and carotid arteries. Finally, we delineate the steps involved in the calibration and setup of the PowerLab system for recording both mPAP and mSAP. This is the first protocol wherein we meticulously explain the surgical procedures for RHC in rats and the recording of mPAP and mSAP. We believe this protocol will be essential for PH research. Investigators with little training in animal handling can reproduce this microsurgical procedure for RHC in rats and measure mPAP and mSAP in rat models of PH. Further, this protocol is likely to help master RHC in rats that are performed for other conditions, such as heart failure, congenital heart disease, heart valve disorders, and heart transplantation.
Keywords: Pulmonary hypertension Right heart catheterization Rat model Mean pulmonary arterial pressure Mean systemic arterial pressure PowerLab system Pulmonary artery catheterization Pulmonary artery catheter Pulmonary artery wedge pressure Invasive monitoring Hemodynamic monitoring
Background
Pulmonary hypertension (PH) encompasses a group of pulmonary vascular disorders with varying etiologies. Depending on the cause and site of the pathogenesis, PH is classified into five groups and many subgroups (Sahay, 2019; Sarah et al., 2021). Despite the differences in the pathological basis of various groups of PH, the disease shares a common definition: a pathological condition in which the mean pulmonary arterial pressure (mPAP) is greater than 25 mmHg at rest or 30 mmHg during exercise (Galiè et al., 2016; Simonneau et al., 2019). Of the five groups of PH, pulmonary arterial hypertension (PAH) and its various subgroups fall in Group I (Wijeratne et al., 2018). In the case of PAH, pulmonary arteries and arterioles undergo pathological changes and thus become thicker and stiffer, resulting in elevated mPAP (Lai et al., 2014; Lan et al., 2018). In other forms of PH (Groups II–V), lung or heart diseases or other conditions increase mPAP (Galiè et al., 2016; Simonneau and Hoeper, 2019).
Because the chief manifestation of PH is increased mPAP, animal models for PH are developed to recapitulate elevated mPAP (Liu and Yan, 2022; Wu et al., 2022). Of the various animal models for PH, Sugen/hypoxia and monocrotaline-induced rat models of PH are two commonly used models (Stenmark et al., 2009; Vitali et al., 2014; Sztuka and Jasinska-Stroschein, 2017) (Figure 1). To develop hypoxia-based models, animals are first treated with a subcutaneous injection of 20 mg/kg Sugen 5416, a vascular endothelial growth factor receptor antagonist. Then, they are housed in 10% oxygen for 3–4 weeks (Bhat et al., 2017; Honda et al., 2020). A right heart catheterization (RHC) is performed to measure mPAP soon after the removal of the animals from the hypoxic environment or after three weeks of hypoxia followed by one week of normoxia (V. Gupta et al., 2013; Nahar et al., 2014; N. Gupta et al., 2015 and 2017; Nahar et al., 2016; Rashid et al., 2017, 2018a, 2018b and 2019; Keshavarz et al., 2019; Al-Hilal et al., 2021). In the case of monocrotaline-based models, animals receive a single subcutaneous injection of 30–80 mg/kg monocrotaline (Lachant et al., 2018; Jing et al., 2020), an alkaloid that causes PH by injuring pulmonary arterial endothelium (Xiao et al., 2017). The mPAP is measured 3–4 weeks after injection (Gomez-Arroyo et al., 2012; N. Gupta et al., 2017). In both models, mPAP rises to as high as 40–60 mmHg.
Figure 1. Sugen-5416/Hypoxia-induced pulmonary arterial hypertension (PAH) model and monocrotaline (MCT) model. For the development of Sugen/hypoxia-based models, animals are first treated with a subcutaneous injection of 20 mg/kg Sugen 5416 and then are housed in 10% oxygen for 3–4 weeks. A right heart catheterization (RHC) is performed to measure mean pulmonary arterial pressure (mPAP) soon after the removal of the animals from the hypoxic environment or after three weeks of hypoxia followed by one week of normoxia. In case of monocrotaline-based models, animals receive a single subcutaneous injection of 30–80 mg/kg monocrotaline. The mPAP is measured 3–4 weeks after MCT injection.
In fact, over the past several decades, these models have been used for studying PH pathophysiology and investigating the efficacy of drugs in reducing mPAP (V. Gupta et al., 2013; Maarman et al., 2013; Nahar et al., 2014; N. Gupta et al., 2015 and 2017; Nahar et al., 2016; Rashid et al., 2017, 2018a, 2018b and 2019; Keshavarz et al., 2019; Al-Hilal et al., 2021; Dignam et al., 2022). For mechanistic studies or to evaluate drugs’ efficacy in ameliorating pathological alterations, pulmonary arteries/arterioles are collected, and various histopathological studies are performed. Further, PH is treated with vasodilators to reduce elevated mPAP (Rose-Jones and McLaughlin, 2015). Reduced mPAP increases survival by reducing right ventricular hypertrophy, the chief cause of death in PH patients (Ryan et al., 2015; Zhu et al., 2019; Benza et al., 2022). However, vasodilator therapy also reduces blood pressure, resulting in peripheral hypotension–induced side effects (Packer, 1985; Rich, 2009; Humbert et al., 2014; Chida-Nagai et al., 2020). Thus, for studying the efficacy of investigational and commercially available drugs in animal models of PH, mPAP is measured as a functional endpoint for therapeutic outcomes (V. Gupta et al., 2013; Maarman et al., 2013; Nahar et al., 2014; N. Gupta et al., 2015 and 2017; Nahar et al., 2016; Rashid et al., 2017, 2018a, 2018b and 2019; Keshavarz et al., 2019; Suen et al., 2019; Al-Hilal et al., 2021; Beckmann et al., 2022; Dignam et al., 2022). Mean system arterial pressure (mSAP) is also measured to evaluate the influence of the drug on peripheral blood pressure or to assess whether the drug or formulation has pulmonary selectivity, by comparing the extent of reduction in mPAP and mSAP in the same rats. In a series of studies, we have shown that pulmonary selectivity can be evaluated by comparing mPAP and mSAP upon administration of various drug therapies to PH rats (V. Gupta et al., 2013; Maarman et al., 2013; Nahar et al., 2014; N. Gupta et al., 2015 and 2017; Nahar et al., 2016; Rashid et al., 2017, 2018a, 2018b and 2019; Keshavarz et al., 2019; Al-Hilal et al., 2021; Dignam et al., 2022).
To measure mPAP, an RHC is performed to place the catheter in the pulmonary artery. Likewise, for mSAP, a catheter is placed in the carotid artery of the same rats that underwent RHC. In our published studies, we inserted polyethylene (PE) catheters in both pulmonary and carotid arteries (N. Gupta et al., 2014; Nahar et al., 2014; Rashid et al., 2018b). However, insertion of PE catheter in pulmonary arteries is challenging because catheters require a specialized curvature for maneuvering from the jugular vein through the right atrium and right ventricle and finally to the pulmonary artery. Further, exposure to the jugular vein for RHC and carotid artery requires microsurgical procedures and specialized tools. Similarly, recording mPAP and mSAP entails using a specialized instrument. PowerLab, a system connected to bridge amplifiers and equipped with Lab Chart Pro software, is universally used to record mPAP and mSAP (Doggett et al., 2018). However, this system must be calibrated and set up accurately.
While we and others have been performing RHC to measure mPAP and mSAP using PowerLab systems for many years now (Rey et al., 2012; V. Gupta et al., 2013; Nahar et al., 2014; N. Gupta et al., 2015 and 2017; Nahar et al., 2016; Rashid et al., 2017, 2018a, 2018b and 2019; Keshavarz et al., 2019; Al-Hilal et al., 2021), there are no published protocols that list the surgical tools, explain the steps for preparation and insertion of catheters, and describe the setup and calibration of PowerLab lab data acquisition systems. In the absence of such a protocol, a new investigator may take months to establish the methodologies for RHC in rats and recording of mPAP and mSAP using the PowerLab system. Importantly, the unavailability of a thorough protocol is a major reason for significant delays in training new lab personnel in RHC or the transfer of methods from one investigator to another in the same lab. Thus, there is an important need for a detailed protocol for performing RHC in rats and measuring such; in this protocol paper, we put together all steps and processes involved in RHC and measurement of mPAP and mSAP, so that investigators with no training can perform the surgery and record mPAP and mSAP. This protocol can also be deployed in performing RHC in animal models for heart failure, congenital heart disease, heart valve disorders, and heart transplantation.
Materials and reagents
Air-TiteTM All-Plastic Henke-JectTM syringes (Fisher Scientific, catalog number: 14-817-25)
Stainless-steel flat instrument tray (Medicus Health, catalog number: 2841M1)
Mayo scissors, straight (Roboz, catalog number: RS-6872) (Figure 2A)
Surgical ophthalmic Westcott tenotomy scissors (Codman, catalog number: 28 54-6513) (Figure 2B)
Vannas spring scissors (Fine Science Tools, catalog number: 91500-09) (Figure 2C)
Vannas micro dissecting spring scissors (Roboz, catalog number: RS-5640) (Figure 2D)
Angled vessel cannulation forceps (Fine Science Tools, catalog number: 18403-11) (Figure 2E)
Serrated Semken forceps (Fine Science Tools, catalog number: 11008-13) (Figure 2F)
Extra fine Graefe forceps, quantity: 2 (Fine Science Tools, catalog number: 11150-10) (Figure 2G)
Graefe forceps, quantity: 2 (Roboz, catalog number: RS-5138) (Figure 2H)
Delicate suture tying forceps (Fine Science Tools, catalog number: 11063-07) (Figure 2I)
Micro serrefines (Fine Science Tools, catalog number: 18055-01) (Figure 2J)
Vein pick (Braintree Scientific Inc., catalog number: V-PIC) (Figure 2K)
25 G needle (Sigma Aldrich, catalog number: Z192406) (Figure 2L)
16 G needle (Becton Dickinson, catalog number: 305198)
HSW syringe 1 mL capacity, polypropylene (Grainger, catalog number: 45UC66)
Silk sutures, non-absorbable, 5-0 (Braintree Scientific, catalog number: SUT-S-106)
PE-50 catheter: polyethylene .023" × .038" per ft. (Braintree Scientific, catalog number: PE50)
Umbili-CathTM 3.5 French single lumen polyurethane umbilical vessel catheter (UVC) (Utah Medical Products, catalog number: 4183505)
Surgical stainless-steel suture (Ethicon, catalog number: DS24)
Acrylic sheet, 6 in. × 6 in., 0.25 inch thick (McMaster-Carr, catalog number: 8560K358)
Umbili-CathTM 3.5 French dual lumen silicone UVC, marked to 34 cm, 20/23 gauge (Utah Medical Products, catalog number: 4273505)
Safelet IV catheter 20 gauge 1" Luer tapered end Teflon (Henry Schein Medical, catalog number: 1198184)
High temperature cautery kit (Fine Science Tools, catalog number: 18010-00)
3MTM MicroporeTM surgical paper tape (Fisher Scientific, catalog number: 19-027761)
Betadine® microbicide solution (Fisher Scientific, catalog number: 19-027136)
Gauze pad (Fisher Scientific, catalog number: 22-362178)
Sprague Dawley rats (Charles River Laboratory, stock number: 400SASSD)
Deionized water (Barnstead Mega-Pure D2, Thermo Scientific)
Sodium chloride, 0.9% (w/v), isotonic saline, Ricca Chemical (Fisher Scientific, catalog number: 7647-14-5)
99% isopropyl alcohol, IPA (VWR, catalog number: IX0235)
Isoflurane liquid (Pharmacompass, NDC: 66794-017-10)
Ketamine hydrochloride, 100 mg/mL (Covetrus, NDC: 11695-0703-1)
Xylazine 20 mg/mL (Heartland Veterinary Supply and Pharmacy, catalog number: 343720-RX, NDC: 59399-110-20)
The cannulation tools shown in Figure 2 are listed in detail in the Materials and reagents section.
Figure 2. Canulation tools. (A) Mayo scissors; straight, (B) surgical ophthalmic Westcott tenotomy scissors, (C) Vannas spring scissors, (D) Vannas micro dissecting spring scissors, (E) angled vessel cannulation forceps, (F) serrated Semken forceps, (G) extra fine Graefe forceps, (H) Graefe forceps, quantity: 2, (I) delicate suture tying forceps, quantity: 2, (J) micro serrefines, (K) vein pick, (L) 25 G needle.
Equipment
Windows desktop computer
PowerLab ML880 16 Channel (ADInstruments)
Bridge Amplifier ML221 (ADInstruments)
Anesthesia induction chamber (Harvard Apparatus, catalog number:75-2030)
Wahl® Trimmer Combo kit (Kent Scientific, catalog number: CL9990-KIT)
SP844 medical pressure transducer sensor (Memscap)
Blood pressure transducer cable kit MLT1199 (Harvard Apparatus, catalog number: 77-0124)
Disposable clip-on BP domes (AD Instruments, catalog number: MLA844)
Blue 3-way stopcock, 2 female Luer locks, swivel male Luer lock (Qosina, catalog number: 99740)
3.5×–90× trinocular stereo zoom inverted light microscope (Amscope, catalog number: SM-3TZ-54S-5M)
Red 3-way stopcock, 2 female Luer locks, swivel male Luer lock (Qosina, catalog number: 99761)
Delta-Cal pressure transducer (Utah Medical Products Inc., catalog number: 650-950)
M3000 tabletop isoflurane, non-rebreathing anesthesia machine (Supera Anesthesia Innovations)
Heated small animal operating table (Harvard Apparatus, catalog number: 50-1247)
Software
Lab Chart Pro Version 7.3.8 (ADInstruments)
Procedure
Preparation of PE-50 catheter for the carotid artery catheterization
Cut a 12-inch piece of PE-50 catheter with Vannas spring scissors (Figure 3A).
Carefully insert a beveled 1 inch 25 G needle on the blunt end of the PE-50 catheter (Figure 3B).
Note: If the catheter punctures during insertion, redo steps A1 and A2. The outer diameter of the 25 G needle is smaller than the inner diameter of the PE-50 catheter. When appropriately inserted, the needle and catheter can fit snugly without perforating the latter.
Prepare a slanted catheter end by cutting the other end of the catheter at a 45° angle (Figure 3C) with Vannas spring scissors.
Note: The slanted end of the catheter is to be inserted into the right carotid artery.
Figure 3. PE-50 catheter preparation for the carotid artery catheterization. (A) A 12-inch piece of PE-50 catheter with Vannas spring scissors and a 25 G needle, (B) Insert the 25 G needle into any end of the PE-50 catheter, (C) Zoomed-in view of the other end of the PE-50 catheter cut at 45° angle with Vannas spring scissors.
Preparation of umbilical venous catheter (UVC) for RHC
Bend 3.3 cm of 16 G beveled needle to a curve at a 65° angle (Figure 4A).
Note: The main pulmonary artery arises from the right ventricle and curves posteriorly and slightly to the left before branching into the left and right pulmonary arteries. The 65° angle bending of the needle and the catheter mimics the curved pulmonary artery.
Insert 2.8 cm of the 3.5 French single lumen polyurethane UVC into the curved needle (Figure 4B).
Note: Proper orientation of the curved needle during catheterization is of utmost importance, with the number markings on the UVC being required to face upwards during insertion of the needle. Furthermore, during insertion of the catheter into the right jugular vein, the orientation will ensure that the curvature of the catheter is parallel to the number markings, permitting the surgeon to determine the insertion length of the UVC.
Submerge the UVC containing the bent needle in 45 °C deionized water for 2 min.
After 2 min, remove the UVC containing the bent needle from the 45 °C deionized water and place it in room temperature deionized water immediately.
Remove the catheter from the needle; upon taking it out from cold water, the shape of the UVC should conform to the shape of the bent needle (Figure 4C).
Note: To reuse the curved UVC, keep the curved needle inserted inside the curved catheter to maintain the curvature, as shown in Figure 4B. It is critical to ensure the curved catheter retains the 65° to avoid missing the pulmonary artery during insertion, which can lead to reinsertions and the eventual puncturing of the artery.
Figure 4. Preparation of a curved catheter. (A) Bend 3.3 cm of a 16 G needle to 65°. (B) Diagram showing the 65° bend. (C) Insert 2.8 cm of UVC into the needle. (D) The catheter is curved by placing a curved needle with the catheter in 45 °C deionized water for 2 min and then putting immediately into room-temperature deionized water.
Steel suture as guide wire setup for RHC (assemble specifically for step H6a)
Attach a 3-way stopcock to a pressure transducer (Figure 5A).
Attach the 3.5 French dual lumen silicone UVC to the 3-way stopcock (Figure 5B).
Insert the surgical stainless-steel suture via the 3.5 French dual lumen silicone UVC up to the end of the catheter tip and tighten the knob to secure the steel suture (Figure 5C).
Figure 5. Steel suture for guide wire setup. (A) A 3-Way Stopcock attached to a pressure transducer. (B) 3.5 French Dual Lumen Silicone UVC attached to a 3-Way Stopcock. (C) Surgical stainless-steel suture inserted through the 3.5 French Dual Lumen Silicone UVC up to the end of the catheter tip and tightened knob, to firmly secure the steel suture. Insert the surgical steel into UVC through the knob and ensure the steel is fully inserted but not protruding out of the other end. The excess steel wire must hang from the knob.
Catheter sleeve preparation for RHC (assemble specifically for step H6b)
Open the Safelet IV catheter 20 gauge 1" Luer tapered end Teflon from its packaging (Figures 6A and 6B).
Detach and remove the needle from the uncut catheter sleeve (Figure 6C).
Note: The needle may be safely disposed of, as it does not serve any further purpose for catheterization. The needle protector, on the other hand, functions as a sleeve for the UVC
Cut the syringe fitting with scissors from the top and trim the tip of the catheter sleeve (Figure 6D).
Insert the French single lumen polyurethane UVC into the catheter sleeve until the tip of the catheter is aligned with the tip of the catheter sleeve (Figure 6E–6G).
Sections C and D are two different methods to perform the right heart catheterization. Irrespective of the method chosen, the data collected will be similar since all these methods lead the UVC to the pulmonary artery.
Figure 6. Catheter sleeve preparation. (A) Safelet IV Catheter 20 Gauge 1" Luer Tapered End Teflon in packaging. (B) Safelet IV Catheter 20 Gauge 1" Luer Tapered End Teflon open tube. (C) Detach the needle from the protecting sleeve. (D) Cut the syringe fitting from the top and trim the tip of the catheter sleeve. (E) The prepared catheter sleeve and the bent UVC. (F) Insert the UVC into the catheter sleeve until the tip of the catheter is aligned with the tip of the catheter sleeve. (G) Once the catheter sleeve is inserted into the right jugular vein, the catheter sleeve can be retracted leaving the bent part of the catheter inside the vein.
Setup of pressure transducer sensors and calibration with PowerLab Data Acquisition system
Setup of PowerLab system (Video 1)
PowerLab system converts the signals received from the rat arteries into numerical data for analysis. One end of the catheter is inserted into the carotid or the pulmonary artery of the rat, and the other end of the catheter is connected to a dome attached to a pressure transducer sensor. The pressure changes in the carotid or pulmonary artery of the rat cause domes to deform, and the pressure transducer senses the deformation to determine the pressure. The pressure transducer converts the pressure into electrical signals and transmits the data to the bridge amplifiers. Bridge amplifiers connect the pressure transducer to the PowerLab, which delivers all data to the computer equipped with Lab Chart Pro Software that shows the outputs as pressure curves.
Video 1. Setup of pressure transducer sensors and calibration with PowerLab Data Acquisition System
Connect the PowerLab system to a USB port of a computer and then connect the PowerLab system to the power outlet.
Connect two bridge amplifiers to the PowerLab. For each rat, use two bridge amplifiers to measure mPAP and mSAP—one for mPAP and another for mSAP—at the same time. For each bridge amplifier, connect the output port of the bridge amplifier to the input port of the PowerLab and connect the input port of the amplifier to the MLT1199 SP844 Kit (Figure 7A).
Note: The PowerLab has 16 input ports; thus, each PowerLab can support up to 16 bridge amplifiers. Two amplifiers, one for measurement of mSAP and another for mPAP are required for each rat; therefore, a PowerLab can support up to eight rats for simultaneous measurement of mPAP and mSAP.
Attach the bottom of the pressure transducer sensors to the acrylic sheet with a hot glue gun. For each rat, two pressure transducer sensors (one for mPAP and one for mSAP) are attached to the acrylic sheet (Figure 7B).
Label each bridge amplifier that is connected to input in transducer sensor and output connected with PowerLab wires with the PowerLab port number. For example, if the bridge amplifier is connected to PowerLab Port #1, label the bridge amplifiers, and its input and output wires as 1 (Figure 7C).
Figure 7. Setting up PowerLab, Bridge amplifiers, and transducers. (A) Overall setup of the equipment for simultaneous measurement of mean systemic arterial pressure (mSAP) and mean pulmonary arterial pressure (mPAP). (B) Zoomed-in view of two pressure transducers attached to a plexiglass plate and the domes with their respective 3-way stopcocks attached. One catheter is inserted in the carotid artery and the other in the jugular vein from the 3-way stopcocks. The transducers are connected to the bridge amplifier. (C) The first connection is between the PowerLab, and the bridge amplifier goes directly from the Power lab to the first bridge amplifier. Each consecutive bridge amplifier is also interconnected with digital input output cables (provided by AD Instruments with the PowerLab). The PowerLab is also the energy source for the bridge amplifiers; therefore, all the bridge amplifiers are connected to the front-end interface of the PowerLab system.
Setup of manual fluid (saline) calibration (Video 1)
Attach one red 3-way stopcock with its female Luer lock inserted in the male swivel Luer lock of the disposable clip-on dome, and attach one blue 3-way stopcock with its male swivel Luer lock inserted on the female Luer lock side of the dome (Figure 8).
Note: Male Luer swivel lock of the dome connects to the female Luer lock side of the 3-way stopcock.
Figure 8. Components and connections of the disposable clip-on dome and 3-way stopcocks are used to connect catheters to the PowerLab system and catheters
Turn the red stopcock knob such that the knob blocks saline flow through the left outlet (Figure 9A), which forces the saline into the dome, towards the blue stopcock. Turn the blue stopcock knob to allow saline to flow in all directions except upwards (Figure 9A).
Fill a 1 mL HSW syringe with 0.9% (w/v) isotonic saline and place it in the inlet of the red stopcock (Figure 9A). The syringe does not require a Luer-lock connection. Press down so that saline flows into the dome (make sure no air bubbles are inside the transducer) until it comes through the right outlet of the blue stopcock. Then, turn the red stopcock knob to restrict saline flow in all directions (Figure 9B).
Note: If the red stopcock knob is not restricted, removal of the syringe will induce air bubbles inside the transducer.
Remove the syringe from the inlet of the red stopcock, refill it with saline, and place the syringe in the inlet of the blue stopcock (Figure 9C). Turn the blue knob to prevent flow towards the dome (Figure 9C) and press down the piston until saline comes out of the right outlet of the blue stopcock (make sure no air bubbles are inside the blue stopcock). Turn the blue stopcock knob (Figure 9D) to restrict saline flow in all directions and leave the syringe in the inlet of the blue stopcock.
Use a second syringe and fill it up with saline. Turn the red stopcock knob to allow saline to flow in all directions except towards the dome (Figure 9D) and place the syringe in the inlet of the red stopcock. Press the piston down until saline comes out through the left outlet (make sure no air bubbles are inside the red stopcock). Leave the syringe in the inlet of the red stopcock.
Insert the dome on a transducer sensor and turn the blue stopcock knob to allow saline to flow in all directions except upwards (Figure 9E). The pressure inside the dome is at 0 mmHg now, which is atmospheric pressure.
Figure 9. Dome saline calibration process. (A) This red knob orientation restricts the saline from flowing to the left of the red 3-way stopcock. This orientation allows the saline from the syringe to flow towards the blue valve. (B) After pressing the syringe, the saline flows towards the right side (arrow); once the dome and the stopcocks are filled with saline, shown as blue region, turn the red knob. This will block (shown by the cross-sign) all the inlets of the red 3-way stopcock and allow the liquid to stay within the dome without any leaks when the syringe is removed. (C) Place a saline-filled syringe on the blue 3-way stopcock, press down to fill all the regions of the blue stopcock with saline, and leave the syringe attached. (D) Place another saline-filled syringe on the red 3-way stopcock, turn the red knob in the orientation shown to restrict flow into the dome, and fill up the rest of the red 3-way stopcock to be entirely filled with saline. (E) After the previous step, the dome is ready to be placed on the sensor for 2-point calibration. Follow this orientation of the knob to do zero calibration explained in step E3b. (F) Use this orientation of knobs to conduct 2-point calibration and attach the Delta-Cal male swivel Luer lock to the female Luer lock of the blue 3-way stopcock, as explained in step E3c.
Manual fluid (saline) two-point calibration of the PowerLab system using Lab Chart Pro (Video 1)
Turn on the PowerLab and open the Lab Chart Pro software.
Attach the Delta-Cal Pressure transducer male swivel connector to the blue stopcock female port (Figure 9F) with the stopcock orientation shown in Figure 9E. The stopcock orientation in Figure 9E is used for zero-calibration only. Turn on the Delta-Cal and the pressure reading should be 0. In Lab Chart Pro, click the Start button to start the continuous measurement of the sensor. Under the channel name where there is a plain white region, right-click and select Bridge Amp. Now click the Zero button to zero calibrate the sensor.
Notes:
i. After turning on Delta-Cal, the reading may not be 0 if any of the stopcocks block the flow within the dome, which builds pressure in the dome.
ii. Make sure there are no vibrations or movements near the sensor during calibration.
iii. Zero function allows the sensor to identify when the electrical signal is at 0 V and the pressure is also at 0 mmHg.
Now, turn the red knob of the red 3-way stopcock to prevent upwards and leftwards flow and retain the blue knob of the blue 3-way stopcock orientation (Figure 9F). This will allow the liquid to be confined within the dome when pressure is applied. The stopcock orientation in Figure 9F is to be used when calibrating at 50 mmHg and 120 mmHg, explained in later steps.
Note: This setting keeps liquid inside the dome; when pressure is applied, the diaphragm in the middle of the dome will expand upwards but the volume of saline in the dome will remain the same.
Go to Setup and then Channel settings. This window shows all 16 channels that will be collecting data (Figure 10). On the second column of the window, select by clicking the number of active channels. Channel 1 corresponds to the bridge amplifier connected to port 1 in PowerLab, and so on. Input the number of channels on the bottom right side of the window, based on the total number of active channels. Press OK to continue.
Notes:
i. It is important to change the input for the number of channels based on active channels, or else the software will show all the channels.
ii. Do not change any other parameters on the window, such as the sample rate and range, until after calibration.
Figure 10. LabChart Pro window showing all 16 channels of the PowerLab Data Acquisition System
In the Lab Chart Pro, use the scale button on the bottom right of the screen and change it to 100:1 or 200:1.
Note: Scaling allows x-axis values of the graph (time) to be within the window for two-point calibration. Using the computer mouse, select the region that covers both parts of the chart.
On the left unselected region adjacent to the continuous measurement, right-click and select Auto Scale.
Note: Auto Scale all the y-values of the graph (current with Amps unit) to be within the window for two-point calibration.
Now, turn the Delta-Cal knob clockwise to increase the pressure to 50 mmHg. In the comment section of Lab Chart Pro, type 50 and press Add. This will mark the point in the data to indicate when the pressure was 50 mmHg. Allow the measurement of data at 50 mmHg for at least 3 s. Increase the pressure in the Delta-Cal to 120 mmHg by turning the knob clockwise, type 120 in the comment section, and press Add to indicate that data point corresponds to 120 mmHg pressure; allow at least 3 s of continuous data measurement at 120 mmHg.
Click Stop in the bottom right part of the screen to stop data collection. Select the region covering both parts in the graph (Figure 11A).
Under the channel name, you will see a plain white region; right-click and select Unit Conversion. On the left bottom side of the window, click the plus and minus signs until the selected region is in view.
Note: The entire graph should be visible in the -6 mV to 6 mV range.
Click on Units and select mmHg. Now, select the region corresponding to 50 mmHg (Figure 11B) and click the arrow next to Point 1. This will input an average of the selected region values. Then type 50 in the box adjacent to it (Figure 11C).
Now, select the region corresponding to 120 mmHg (Figure 11D) and click the arrow next to Point 2. This will input an average of the selected region values. Then, type 120 in the box adjacent to it (Figure 11E) and press OK. This completes the two-point calibration (Figure 11F).
To verify the calibration, click Start for continuous data collection. The number (mmHg) on the top of the channel name should be the same as the number (mmHg) on Delta-Cal. You may decrease the pressure in Delta-Cal to 80 mmHg and check if it shows 80 mmHg on the computer screen.
Note: An error of ± 1 is acceptable due to machine error.
Repeat steps E3c–E3m for each pressure transducer sensors, since one transducer requires one channel feedback in the Lab Chart Pro.
Figure 11. 2-point calibration using Lab Chart Pro. (A) Selecting the region from the collected continuous measurement for unit conversions using the 2-point calibration method. (B) Select the region that shows continuous measurement at 50 mmHg and click on the arrow next to Point 1 to average the values selected and type that value. (C) Type 50 next to the second arrow for Point 1 to allow the sensor to convert the value in the first box to reflect 50 mmHg. Change the Units box to mmHg. (D) Select the region that shows continuous measurement at 120 mmHg and click on the arrow next to Point 2 to average the values selected and input that value. (E) Type 120 next to the second arrow for Point 2 to allow the sensor to convert the value in the first box to reflect 120 mmHg. (F) Unit conversion window for completed 2-Point calibration.
A PE-50 catheter or UVC heads can now be fit into the red stopcock male port, with the red knob allowing saline flow to the left and right but no flow upwards and the blue knob turned such that there is no flow (Figure 12A). Press the syringe filled with saline to allow the catheters to be pre-filled with saline. To begin measurement of mPAP and mSAP, match the knob orientation shown in Figure 12B, insert the other end of the catheter into the jugular vein or the carotid artery.
Figure 12. Valve orientations to pre-fill saline and begin measurements of mPAP and mSAP. (A) Valve orientation for catheter ready to pre-fill with saline: turn the red knob such that flow from the left syringe to the left outlet of the red stopcock is allowed and turn the blue knob such that flow is restricted in all directions. (B) Valve orientation for catheter ready to begin measurement. Turn the red knob such that the upwards flow is blocked and turn the blue knob such that flow in all directions is blocked. The arrow indicates the female red knob port that attaches to a PE-50 catheter or UVC.
Preoperative procedures
Prepare surgery tray with the tools shown in Figure 2 before surgery. Clean all tools with soap and water and wipe them with 70% isopropyl alcohol.
Place the heating pad on a flat surface under the inverted light microscope and clean the heating pad surface with 70% isopropyl alcohol.
Note: The light microscope helps to better visualize the arteries and catheterization.
Turn on the heating pad and set the temperature to 37 °C.
Note: Maintain the temperature at 37 °C throughout the surgery and experiment.
Place the rats in the inhalation anesthesia chamber with 2.5% isoflurane. Once the rat falls asleep, inject intraperitoneally 300 μL per 250 g rat body weight with a cocktail of 1 mL of 100 mg/mL ketamine and 100 μL of 20 mg/mL xylazine. Confirm anesthesia by the toe-pinch method. More anesthesia can be administered in small increments if sufficient induction is not obtained with the initial dose.
Note: One dose of intraperitoneal anesthesia will keep the rat anesthetized for at least one hour. To ascertain the state of unconsciousness, the toe-pinch method may be utilized at any point of the surgery. Sudden movements in the rat should be avoided. However, any muscle movements or agitation observed in the rat during the surgical procedures are an indication that the rat is beginning to regain consciousness. In such cases, an additional dose of anesthesia should be administered.
Place the rat in the supine position on the heating pad (Figure 13A). Restrain all four legs with micropore tape and the head by placing a string under the upper incisors, as shown in Figure 13A.
(Optional) Using a hair trimmer, remove hair from the ventral neck and the dorsal area between the scapulae.
Scrub the trimmed area with Betadine® and then with 70% isopropyl alcohol for three cycles.
Note: Steps F6 and F7 are optional for experienced researchers but recommended for those who are new to the process.
Figure 13. Restraining the rat and exposing the ventral neck area for catheterization of the left carotid artery. (A) Position the rat with the chest facing upwards and the head towards the surgeon on the heating pad. (B) Using straight Mayo scissors, make an incision of ~3 cm at the middle line of the ventral neck area and use the scissors as pliers to detach the skin from the subcutaneous tissues. (C) Adjacent to the left side of the trachea, go deeper by cutting the connective tissues and muscles with tenotomy scissors for coarse cutting. (D) Separate the muscle layers to cut deeper into the left side of the trachea until the nerve and the carotid artery is exposed. (E) Using curved tip Graefe forceps, gently separate the nerve that is attached to the left carotid artery. (F) The carotid artery thus exposed and free from innervation is used for catheterization.
Catheterization of the left carotid artery (Video 2)
Video 2. Left carotid artery for mean systemic arterial pressure (mSAP). The animal studies were performed according to the guidelines from the University of Illinois at Chicago approved by the Animal Research Ethics Board of University of Chicago under the protocol #: ACC21-180.
Using straight Mayo scissors, make a skin incision of ~3 cm at the middle line of the ventral neck area and use the scissors as pliers to detach the skin from the subcutaneous tissues (Figure 13B). Adjacent to the left side of the trachea, go deeper by cutting the connective tissues and muscles with tenotomy scissors for coarse cutting and spring scissors for finer cutting, until the left carotid artery is visible (Figure 13B–13D).
Notes:
Be careful not to puncture or cut the trachea and the left carotid artery while removing tissues and muscles.
Minor bleeding may appear due to cutting small blood vessels in the area. So, apply a gauze pad to control the bleeding or cauterize vessels using the cautery kit.
Using Graefe forceps, gently separate the nerve around the left carotid artery (Figure 13E).
Notes:
Use the microscope to locate the left carotid artery to avoid severing surrounding blood vessels or nerves.
Make sure that the nerve is separated from the carotid artery (Figure 13F).
Place a Graefe forceps under the left carotid artery (Figure 14A), lift it up, and keep it exposed for surgical manipulation. Leave the Graefe forceps towards the left side of the rat body away from the trachea.
Note: If the forceps is on the top of the trachea, it may choke the rat.
Using a 5-0 suture, tie one end of the carotid artery (towards the head) by making a double knot and then a single knot (Figure 14B) to stop the blood flow to the brain. Place a micro serrefine on the other end of the carotid artery (towards the heart) (Figure 14C).
Notes:
Use the microscope to assess whether the artery is throbbing; if it is, placing a micro serrefine should help reduce artery throbbing.
Usually, lifting the artery with Graefe forceps (in step G2) prevents blood flow and throbbing.
Do not cut the excess suture, because excess suture will be used to anchor the catheter to the carotid artery in step G8.
Using a 5-0 suture, leave an open double knot on the other end of the carotid artery (towards the heart) but before the micro serrefine (Figure 14D).
Note: The open double knot must not be tied before inserting the catheter through the left carotid artery but tied after insertion of the catheter.
Using a micro dissecting spring scissor, make a ~0.5 mm hole in the carotid artery (Figure 14E).
Notes:
~0.5 mm is slightly larger than the diameter of the catheter tip.
Be careful not to sever the carotid artery when making the hole using micro dissecting spring scissor.
Hold the PE-50 catheter (prepared in Section A) with angled vessel cannulation forceps (Figure 14F and Figure 14G), insert 1.0 cm of the catheter into the hole of the carotid artery (Figure 14H), and secure it with a piece of micropore tape (Figure 14I). Further secure the catheter in place by closing the open double knot, followed by tying a single knot on top of the inserted catheter (Figure 14J). Anchor the catheter to the carotid artery with the first suture (as stated in step G4) by making another double knot and then a single knot. Then, remove the micro serrefine (Figure 14K) and confirm the characteristic carotid artery peaks in the computer monitor (Figure 18). Now the mSAP can be measured.
Note: The flushing of a catheter with saline solution is a crucial procedure to be carried out during arterial catheterization. It serves the purpose of clearing any blockages that may occur in the catheter channel due to the coagulation of blood. A pre-filled syringe containing saline solution is used to exert pressure on the catheter and remove any obstructions. The presence of a clogged catheter can lead to erroneous or non-existent feedback reaching the sensors, thereby compromising the accuracy of any measurements obtained from the catheter.
In addition, secure the dangling part of the catheter from the rat’s body to the transducer with a piece of micropore tape to the heating pad. These will further secure the catheter from unwanted movement.
Place gauze pads to cover the exposed left side of the trachea and secure them with micropore tapes (Figure 14L).
Note: Because this protocol entails continuous measurement of mSAP for at least 6 h, the exposed skin should be covered with gauze pads to prevent the area from drying.
Upon completion of all mSAP measurements, the PE-50 catheter should be flushed with a syringe containing 99% isopropyl alcohol. The catheter may be reused if it remains unpunctured and displays transparency after the cleaning procedure.
Figure 14. Catheterization of the left carotid artery. (A) Place one Graefe forceps under the left carotid artery to lift it up and keep it exposed for surgical manipulation. (B) Using a 5-0 suture, tie one end of the carotid artery (towards the head) by making a double knot and then a single knot. (C) Using the microscope, check whether the artery starts throbbing. If the artery is throbbing, place a micro serrefine on the other end of the carotid artery (towards the heart) to stop throbbing. (D) Using a 5-0 suture, leave an open double knot on the other end of the carotid artery (towards the heart) but before the micro serrefine. (E) Using a micro dissecting spring scissor, make a ~0.5 mm hole in the carotid artery. (F) This angled vessel cannulation forceps has teeth to grip the catheter. (G) Grab the prepared PE-50 catheter for the carotid artery with angled vessel cannulation forceps. (H) Insert 1 cm of the catheter via the hole in the carotid artery. (I) Secure the inserted catheter with a piece of micropore tape. (J) Secure the catheter in place by closing the open double knot followed by tying a single knot on top of the inserted catheter. Shorten all the sutures and clean any blood. (K) Remove the micro serrefine to allow the blood to flow and confirm the characteristic carotid artery peaks in the computer monitor. (L) Place gauze pads to cover the exposed left side of the trachea and secure them with pieces of micropore tape.
Catheterization of right jugular vein and pulmonary artery (Video 3)
Since this protocol involves simultaneous measurement of mSAP and mPAP, the right jugular vein of the same rat used to measure mSAP in Section G should be catheterized for measurement of mPAP.
Video 3. Right jugular vein for mean pulmonary arterial pressure (mPAP).The animal studies were performed according to the guidelines from the University of Illinois at Chicago approved by the Animal Research Ethics Board of University of Chicago under the protocol #: ACC21-180.
Using tenotomy scissors for coarse cutting and spring scissors for finer cutting, separate the muscle layers on the far-right side of the trachea near the scapula, to find and expose the bifurcation of the right jugular vein (Figure 15A).
Note: The same skin incision that was performed to expose the left carotid artery is used to locate and expose the right jugular vein.
Place a Graefe forceps under the vein below the bifurcation (Figure 15B) to lift the jugular vein up and leave the forceps to keep the vein exposed for surgical manipulation. As the vein goes towards the heart, the vein diameter gets larger, which is the area of interest.
Note: You can see the bifurcation of the jugular vein at this step. Identification of the bifurcation is important for catheterization of jugular vein (Figure 15C).
Using a 5-0 suture, tie one end of the jugular vein (towards the head but below the bifurcation) by making a double knot and then a single knot that stops blood flow from the head area (Figure 15D).
Using a 5-0 suture, leave an open double knot on the other end of the jugular vein (towards the heart) (Figure 15E).
Grab the jugular vein, where the lumen diameter is larger, with another Graefe forceps and make a small incision of ~0.5 mm on the jugular vein with a micro dissecting spring scissor like the one shown in Figure 15E.
Catheterization of the jugular vein can be performed using three different methods: (a) catheterization using steel guide wire can be used for rats with health issues or young rats and healthy adult rats; (b) catheterization using a catheter sleeve is recommended only for healthy adult rats; (c) catheterization using only the catheter is recommended for skilled surgeons.
Figure 15. Catheterization of the right jugular vein. (A) Using tenotomy scissors for coarse cutting and spring scissors for finer cutting, separate the muscle layers on the far-right side of the trachea (near scapula) to identify and expose the bifurcation of the right jugular vein. (B) Place a Graefe forceps past the bifurcation (Fig. 12B) to lift the jugular vein up and keep it exposed for surgical manipulation. (C) Microscopic view of the bifurcation. (D) Using a 5-0 suture, tie one end of the jugular vein (towards the head but below the bifurcation) by making a double knot and then a single knot that stops blood flow from the brain. (E) Using a 5-0 suture, leave an open double knot on the other end of the jugular vein (towards the heart). (F) Slowly insert the catheter into the jugular vein for approximately 6–8 cm from the point of insertion, depending on the age of the rat.
Catheterization using steel suture as guide wire (corresponding to section C):
i. Adjust the transducer to adapt to the steel suture as guide wire setup as discussed in section C.
ii. Use the UVC with steel suture as guide wire using angled vessel cannulation forceps similar to that shown in Figure 14G.
iii. With another Graefe forceps, hold one side of the hole made in the jugular vein for easy catheterization.
iv. Slowly insert the catheter into the jugular vein for approximately 6–8 cm from the point of insertion, depending on the weight of the rat, in the specific orientation as shown in Figure 15F. If the weight of the rat is less than 250 g, insert the catheter 6 cm; if the rat weighs more than 250 g, insert 8 cm.
v. Pull the steel suture outward to allow the curved catheter to conform to its original curved shape inside the vein.
Catheterization using a catheter sleeve (corresponding to section D) (method used in Video 3):
i. Grab the catheter sleeve (Figure 16A) with the catheter using angled vessel cannulation forceps.
ii. While keeping the jugular vein grabbed, insert 1 cm of the catheter sleeve into the jugular vein in the specific orientation as shown in Figure 16B.
iii. Now insert the catheter in for approximately 6–8 cm from the point of insertion depending on the age of the rat.
iv. While holding the catheter stationary (Figure 16C), pull the catheter sleeve out to the other end of the catheter.
Figure 16. Using catheter sleeves for RHC. (A) Grab the bent UVC through the catheter sleeve. (B) Ensure that the catheter sleeve and the bent UVC are aligned with each other’s edges in this specific orientation with the black marks facing the surgeon. (C) Insert the catheter sleeve into the right jugular vein, and when 1 cm of the catheter is inside the vein, pull the sleeve backward leaving only the UVC inside the vein.
Catheterization using the UVC only:
i. Grab the catheter using hollowed angled vessel cannulation forceps.
ii. With extra fine Graefe forceps, grab one side of the hole made in the jugular vein for easy insertion of the catheter.
iii. Carefully insert the catheter into the jugular vein for approximately 6–8 cm from the hole made, depending on the age of the rat, in the specific orientation as shown in Figure 15F.
Note: When the curved catheter is inserted into the jugular vein, the shape of the catheter conforms to the vein; as it is maneuvered down the vein, the tip of the catheter starts to assume the curved shape, and once it reaches the pulmonary artery it assumes the original curved shape (60°–65° angle).
The pressure readings and characteristic peaks, displayed by the power system, change as the catheter is maneuvered through the right atrium, right ventricle, and then in the pulmonary artery. The pressure in the right atrium, right ventricle, and pulmonary artery should be 2–6 mmHg, 0–25 mmHg, and 10–25 mmHg, respectively, with their characteristic peaks (Figure 17).
Once the characteristic peak of the pulmonary artery is confirmed, secure the catheter with a piece of micropore tape. Further, secure the catheter by closing the open double knot followed by tying a single knot on top of the inserted catheter. Now the mPAP can be measured.
Place gauze pads to cover the exposed right side of the rat and secure them with micropore tapes.
Note: Because this protocol entails the continuous measurement of mPAP for at least 6 h, the exposed skin needs to be covered with gauze pads to prevent dryness or adverse reaction due to long-time exposure to the environment.
After the completion of data collection, the rat should be euthanized using isopropyl alcohol. A syringe should be removed from the domes and filled with isopropyl alcohol. The alcohol-filled syringe should then be injected into the transducer, which will subsequently travel into the rat. A flat line measurement of mPAP or mSAP confirms rat euthanasia.
Upon completion of all mPAP measurements, the UVC should be flushed with a syringe containing 99% isopropyl alcohol. The catheter may be reused as long as it remains unpunctured and displays transparency after the cleaning procedure.
Figure 17. Characteristic peaks during RHC in the right atrium and right ventricle help to identify the location of the UVC as it maneuvered to the pulmonary artery. Used with permission from Dr. Richard Klabunde https://www.cvphysiology.com/Heart Failure/HF008.
Data analysis
Lab Chart Pro collects mPAP and mSAP data in a continuous fashion every second (Figure 18). Data are first transferred to Excel for further processing.
Figure 18. LabChart Pro Software showing the outputs as pressure curves
Identify mPAP and mSAP data at baseline point or at zero time, as shown in Table 1 and Table 2.
Table 1. Mean pulmonary arterial pressure (mPAP) data points of three groups: saline, pulmonary group; plain PGE, IV (120 μg/kg) group; and plain PGE, pulmonary (120 μg/kg), plotted in Figure 19A
Time (min) Saline, pulmonary Plain PGE, IV (120 μg/kg) Plain PGE, pulmonary (120 μg/kg)
mPAP % decrease mPAP % decrease mPAP % decrease
0 52.00 0.00 49.50 0.00 51.50 0.00
5 45.24 13.00 41.58 16.00 44.81 13.00
10 45.50 12.50 33.17 33.00 36.57 29.00
15 45.76 12.00 34.65 30.00 35.79 30.50
20 52.00 0.00 35.64 28.00 36.82 28.50
25 - - 40.84 17.50 36.57 29.00
30 52.00 0.00 42.08 15.00 38.11 26.00
35 - - 37.62 24.00 36.05 30.00
40 - - 45.05 9.00 39.91 22.50
45 - - 48.02 3.00 40.43 21.50
50 - - 49.50 0.00 49.44 4.00
Table 2. Mean systemic arterial pressure (mSAP) data points of three groups: saline, pulmonary group; plain prostaglandin E (PGE), IV (120 μg/kg) group; and plain PGE, pulmonary (120 μg/kg), plotted in Figure 19B
Time (min) Saline, pulmonary Plain PGE, IV (120 μg/kg) Plain PGE, pulmonary (120 μg/kg)
mSAP % decrease mSAP % decrease mSAP % decrease
0 115.00 0.00 119.00 0.00 118.00 0.00
5 112.13 2.50 73.19 38.50 89.09 24.50
10 108.68 5.50 63.07 47.00 75.52 36.00
15 109.25 5.00 64.26 46.00 75.52 36.00
20 109.83 4.50 69.02 42.00 79.06 33.00
25 110.98 3.50 80.92 32.00 91.45 22.50
30 109.25 5.00 80.33 32.50 94.40 20.00
35 112.13 2.50 95.20 20.00 97.35 17.50
40 117.30 -2.00 116.62 2.00 109.74 7.00
45 - - 121.38 -2.00 109.15 7.50
50 - - 121.38 -2.00 115.05 2.50
Spot and collect mPAP and mSAP at various time intervals (0, 15, 30, 60, 90, 120 to 240 min).
Plot mPAP and mSAP value against time.
Data can also be plotted in percentage decrease mPAP and mSAP, as shown in Figure 19A and Figure 19B. To calculate percent decrease, use the formula , based on initial mPAP, and the formula , based on initial mSAP. The mPAP (at a given time point) and mSAP (at a given time point) refer to any mPAP or mSAP values from the table at any time point, to find the percentage decrease at that time with respect to the original mPAP or mSAP, respectively.
Figure 19. Acute pulmonary hemodynamic efficacy of plain prostaglandin E1 (PGE1) administered either intravenously (IV) or intratracheally (pulmonary). (A) mean pulmonary arterial pressure (mPAP) and (B) mean systemic arterial pressure (mSAP) after administration of plain PGE1. Data represent mean ± SD, n = 4–6; *p < 0.05.
Acknowledgments
This study is supported in parts by a fund from NIH 1R01HL144590-01 (to F.A.), NIH R42HL151045 (to F.A.) and Cardiovascular Medical Research and Education Funds (to F.A). D.S. is supported in part by a fund from the DOD (W81XWH-20-1-0702).
Competing interests
The authors declare no competing interests.
Ethics considerations
Animal studies: the animal studies were performed according to the protocol approved by the IACUC of the University of Illinois at Chicago (Protocol #: ACC21-180)
References
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Simonneau, G., Montani, D., Celermajer, D. S., Denton, C. P., Gatzoulis, M. A., Krowka, M., Williams, P. G. and Souza, R. (2019). Haemodynamic definitions and updated clinical classification of pulmonary hypertension. Eur Respir J 53(1): 1801913.
Stenmark, K. R., Meyrick, B., Galie, N., Mooi, W. J. and McMurtry, I. F. (2009). Animal models of pulmonary arterial hypertension: the hope for etiological discovery and pharmacological cure. Am J Physiol Lung Cell Mol Physiol 297(6): L1013-1032.
Suen, C. M., Stewart, D. J., Montroy, J., Welsh, C., Levac, B., Wesch, N., Zhai, A., Fergusson, D., McIntyre, L. and Lalu, M. M. (2019). Regenerative cell therapy for pulmonary arterial hypertension in animal models: a systematic review. Stem Cell Res Ther 10(1): 75.
Sztuka, K. and Jasinska-Stroschein, M. (2017). Animal models of pulmonary arterial hypertension: A systematic review and meta-analysis of data from 6126 animals. Pharmacol Res 125(Pt B): 201-214.
Vitali, S. H., Hansmann, G., Rose, C., Fernandez-Gonzalez, A., Scheid, A., Mitsialis, S. A. and Kourembanas, S. (2014). The Sugen 5416/hypoxia mouse model of pulmonary hypertension revisited: long-term follow-up. Pulm Circ 4(4): 619-629.
Wijeratne, D. T., Lajkosz, K., Brogly, S. B., Lougheed, M. D., Jiang, L., Housin, A., Barber, D., Johnson, A., Doliszny, K. M. and Archer, S. L. (2018). Increasing Incidence and Prevalence of World Health Organization Groups 1 to 4 Pulmonary Hypertension: A Population-Based Cohort Study in Ontario, Canada. Circulation Cardiovascular quality and outcomes 11(2): e003973.
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Detection and Quantification of Calcium Ions in the Endoplasmic Reticulum and Cytoplasm of Cultured Cells Using Fluorescent Reporter Proteins and ImageJ Software
SS Shunsuke Saito
Kazutoshi Mori
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4738 Views: 1339
Reviewed by: Chiara AmbrogioTanxi Cai Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Nov 2022
Abstract
This protocol describes a method for detecting and quantifying calcium ions in the endoplasmic reticulum (ER) and cytoplasm of cultured cells using fluorescent reporter proteins and ImageJ software. Genetically engineered fluorescent reporter proteins, such as G-CEPIA1er and GCaMP6f, localize to intracellular regions of interest (i.e., ER and cytoplasm) and emit green fluorescence upon binding to calcium ions. In this way, the fluorescence brightness of cells transfected with expression vectors for these reporters reflects the calcium ion concentration in each intracellular region. Here, we describe procedures for observing cultured cells expressing these fluorescent reporters under a fluorescence microscope, analyzing the obtained image using the free image analysis software ImageJ (https://imagej.net/ij/index.html), and determining the average fluorescence brightness of multiple cells present in the image. The current method allows us to quickly and easily quantify calcium ions on an image containing multiple cells and to determine whether there are relative differences in intracellular calcium ion concentration among experiments with different conditions.
Key features
•Detection and quantification of calcium ions in the ER and cytoplasm using fluorescent reporter proteins
•Quick and easy verification of measurement results using ImageJ
•Simultaneous comparison between various experimental conditions (drug treatment, mutants, etc.)
Keywords: Calcium imaging G-CEPIA1er GCaMP6f ImageJ Endoplasmic reticulum Cytoplasm Cultured cells Fluorescence microscopy
Background
Calcium ions function as second messengers that mediate various signal transductions in cells (Berridge et al., 2000). Cytosolic calcium ions are actively taken up into the endoplasmic reticulum (ER) by sarcoplasmic calcium ATPase (SERCA), a calcium ion pump present in the ER membrane (Brini and Carafoli, 2009), to ensure that the calcium ion concentration in the ER is maintained at levels more than 5,000-fold higher than those in the cytoplasm (Laude ant Simpson, 2009). Calcium ions stored in the ER are released into the cytoplasm via the calcium ion channels inositol triphosphate receptor and ryanodine receptor as needed, and then rapidly recovered into the ER via SERCA. Such transient increase in cytosolic calcium ion concentration has been shown to be necessary for many biological phenomena, including muscle contraction and neurotransmission (Berridge et al., 2003).
At the same time, calcium ions are extremely important for maintaining protein homeostasis in the ER. The ER is where newly synthesized secretory and transmembrane proteins destined for the secretory pathway form the correct three-dimensional structure (Ellgaard and Helenius, 2003). BiP (immunoglobulin heavy chain–binding protein) is an Hsp70-type chaperone responsible for protein folding in the ER. For BiP to exert its activity normally, it is essential that a high concentration of calcium ions is maintained in the ER; BiP is a low-affinity and high-capacity calcium-binding protein (Preissler et al., 2020).
Based on these facts, disruption of calcium ion homeostasis in the ER not only impairs muscle and nerve function but also causes normal protein folding in the ER to be inhibited. As a result, the ER can enter a state termed ER stress, in which structurally abnormal proteins accumulate in the ER, which may in turn eventually lead to cell death. Over time, it was realized that such disruption of ER calcium ion homeostasis might actually be involved in aging-associated muscle dysfunction (Delrio-Lorenzo et al., 2020) and various neurodegenerative diseases (Schrank et al., 2020; Saito et al., 2022).
Traditional methods of detecting intracellular calcium ions have long used acetoxymethyl ester (AM)-conjugated versions of fluorescence-indicating chemicals such as Fura-2 AM, which mainly localizes in the cytoplasm, or Mag-Fura-2 AM, which localizes in the cytoplasm and ER. Fura-2 and Mag-Fura-2 themselves are composed of a calcium ion–chelating moiety and a fluorescent cluster moiety. The binding of calcium ions alters their conformation, resulting in changes in the efficiency with which they absorb excitation light and emit fluorescence. Importantly, conjugation of AM with these indicators inhibits their binding to calcium ions outside the cell and simultaneously confers membrane permeability on them (Tsien, 1981). Accordingly, these AM-conjugated indicators penetrate the plasma membrane, followed by removal of their AM parts by cellular esterase. This allows them to accumulate inside cells, bind to intracellular calcium ions, and emit fluorescence. As a result, calcium ion concentration in the cytoplasm can be easily measured using cells treated with Fura-2 AM.
In the case of Mag-Fura-2 AM, this is used by first determining the fluorescence intensity in cells treated with Mag-Fura-2 AM, and then by similarly determining intensity in these cells after treatment with digitonin or saponin, which permeabilize the plasma membrane but not the ER membrane. This second value represents calcium ion concentration in the ER; subtracting this from the first value reveals calcium ion concentration in the cytoplasm (Hofer and Machen, 1993).
In the current protocol, recently developed and genetically engineered fluorescent reporter proteins such as G-CEPIA1er (Suzuki et al., 2014) and GCaMP6f (Chen et al., 2013) are expressed in cultured cells separately by transfection. These are fusion proteins consisting of a circularly permutated fluorescent protein, a calmodulin domain, and the M13 fragment from myosin light chain kinase. Binding of calcium ions to the calmodulin domain causes a conformational change that results in the emission of fluorescence. Because of their optimized affinity for calcium ions, large dynamic range, and specific intracellular localization, fluorescence from G-CEPIA1er or GCaMP6f reports calcium ion dynamics in the ER and cytoplasm, respectively, in a single measurement. We explain how to quickly and easily test the results using ImageJ, a free analysis software. By applying this protocol, it is possible to detect and quantify ER and cytoplasmic calcium ions in a variety of plasmid-transfectable cells without complicated sample preparation procedures using different reagents.
Materials and reagents
Homo sapiens neuroblastoma cell line SH-SY5Y (ATCC, catalog number: CRL-2266)
Homo sapiens colon colorectal carcinoma cell line HCT116 (ATCC, catalog number: CCL-247)
24-well plate (for SH-SY5Y) (Corning, Falcon®, catalog number: 353047)
6-well plate (for HCT116) (Corning, Falcon®, catalog number: 353046)
Dulbecco’s modified Eagle’s medium (DMEM) (Nacalai Tesque, catalog number: 08458-45)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, GibcoTM, catalog number: 10270-106)
100 U/mL penicillin and 100 μg/mL streptomycin (Nacalai Tesque, catalog number: 26253-84)
Opti-MEM (Thermo Fisher Scientific, GibcoTM, catalog number: 31985-070)
Polyethylenimine max (Polyscience, catalog number: 24765-100)
Lipofectamine® LTX and PlusTM Transfect (Thermo Fisher Scientific, InvitrogenTM, catalog number: 15338100)
pCMV G-CEPIA1er (Addgene, plasmid, catalog number: 58215)
pGP-CMV-GCaMP6f (Addgene, plasmid, catalog number: 40755)
pCMV-myc-wtSeipin (Satio et al., 2022)
pCMV-myc-ngSeipin (Satio et al., 2022)
1 mM thapsigargin (Calbiochem, catalog number: T9033) dissolved in DMSO (stored at -20 °C)
Equipment
Fluorescence stereomicroscope (Olympus IX-71-22TFL/PH)
Acquisition software (DP Controller 1.2.1.108)
Software
ImageJ (https://imagej.net/ij/index.html)
Excel (Microsoft)
Procedure
Detection and quantification of calcium ions in the ER
Preparation of cultured cells expressing reporter protein
Culture SH-SY5Y cells in DMEM with glucose (4.5 g/L) supplemented with 10% FBS and antibiotics (100 U/mL penicillin and 100 μg/mL streptomycin) in an incubator at 37 °C and a 5% CO2 atmosphere.
Transfect SH-SY5Y cells (3.0 × 105 cells, approximately 80% confluent on a 24-well plate) with 200 ng of pCMV G-CEPIA1er in 63 μL of Opti-MEM, 3.2 μL of Lipofectamine®︎ LTX, and 0.6 μL of PlusTM Transfect.
Detection and acquisition of images of cells expressing reporter protein
Twenty-eight hours after transfection, observe the cells under an inverted fluorescence microscope (e.g., Olympus IX-71-22TFL/PH) with an appropriate exposure time. If phenol red contained in DMEM causes a strong background signal, the volume of DMEM should be accordingly reduced. If a sufficient number of cells is present on the dish (approximately 95% confluent) and transfection efficiency is sufficient (approximately 15%), images should be taken using a 20× objective lens. This will produce images that contain approximately 100 fluorescent cells per image (Figure 1).
Figure 1. Fluorescence images of wild-type (WT) and Seipin-knockout (KO) SH-SY5Y cells expressing G-CEPIA1er. WT or Seipin-KO SH-SY5Y cells seeded on 24-well plates were transfected with 200 ng of pCMV G-CEPIA1er and observed for fluorescence 28 h later. Scale bar, 100 μm.
Quantification of fluorescence by ImageJ
Export the obtained images in tif or another appropriate file format and open them in ImageJ (Figure 2A).
Select Image > Color > Split Channels to split each image into Red, Green, and Blue channels. Of these, use only the Green-channel image for quantification and close the others (Figure 2B).
Select Process > Subtract Background to perform background subtraction, with Rolling Ball Radius = 50.0 pixels (Figure 2C and 2D).
Select Adjust > Threshold to set the threshold such that the quantitative range is limited only to the area where cells are present (Figure 2E and 2F).
Select Analysis > Set Measurement and check the “Mean grey Value” and “Limit to threshold” checkboxes (Figure 2G).
Select Analysis > Measure to obtain the quantification results (Figure 2H).
Statistical analysis in Microsoft Excel (Figure 3)
Paste the results from Figure 2H (A1:B17 and D1:E16).
Calculate the average of the results of the group that should be used as the reference (B21).
Divide each result by the average of the reference group (A25:B41 and D25:E40).
Calculate the average (H26 and I26) and standard deviation (H27 and I27) of the divided result of each group.
Use the F-test to determine if the variances are equal between the divided results of the two groups being compared (L26) and use Student’s t-test to determine if there is a significant difference between the divided results of the two groups (L27).
Figure 4 shows detection and quantification of calcium ions in the ER of wild-type and Seipin-knockout SH-SY5Y cells using this protocol.
Figure 2. Procedures for analyzing fluorescence images using ImageJ. A. The tif file of the fluorescence image from Figure 1A opened in ImageJ. B. By selecting Image > Color > Split Channels and splitting each image into Red, Green, and Blue channels, a Green-channel image was obtained. C. and D. By selecting Process > Subtract Background, background subtraction was performed with Rolling Ball Radius = 50.0 pixels (C). This produced a background-subtracted version of the image (D). E, F. By selecting Adjust > Threshold, the threshold was set such that the quantitative range was limited to areas where cells were present (E). With this step, cell-free areas (blue) were marked and excluded from the analysis area (F). G. After selecting Analysis > Set Measurement, checkboxes for “Mean grey Value” and “Limit to threshold” were checked. H. Selecting Analysis > Measure provided the quantification results (mean = 26.833).
Figure 3. An example of the statistical data analysis. The results from each group (acquired following the procedure described in Figure 2) are pasted in A1:B17 and D1:E16. The average of results of the reference group (A1:B17 in this example) is calculated in B21. The result of dividing each result by the average of the reference group (B21) is shown in A25:B41 and D25:E40. The results of calculating the average and standard deviation for the divided result of each group are shown in H26 and I26, and H27 and I27, respectively. The results of the F-test to determine if the variances were equal between the divided results of the two groups being compared are shown in L26, and the result of Student’s t-test to determine if there was a significant difference between the divided results of the two groups is shown in L27.
Figure 4. Calcium ion concentrations in the endoplasmic reticulum (ER) of wild-type WT and Seipin-knockout (KO) SH-SY5Y cells. A. WT (1) or Seipin-KO (2) SH-SY5Y cells seeded on 24-well plates were transfected with 200 ng of pCMV G-CEPIA1er and observed for fluorescence 28 h later. Scale bar, 100 μm. B. For both cell types, 5–6 images were taken per sample, and the experimental results for three samples (15–16 images in total) were analyzed following the procedures described in Figure 2 and statistically processed following the procedures described in Figure 3. The results are expressed as relative values, with the mean of measurements of WT SH-SY5Y cells set as 1, along with the standard deviation and presence of significant differences (Student’s t-test, ***: p < 0.001). The results show that Seipin-KO SH-SY5Y cells have a significantly lower calcium ion concentration in the ER than WT cells (Saito et al., 2022).
Detection and quantification of calcium ions in the cytoplasm
Preparation of cultured cells expressing reporter protein
Culture HCT116 cells in DMEM (glucose 4.5 g/L) supplemented with 10% FBS and antibiotics (100 U/mL penicillin and 100 μg/mL streptomycin) in an incubator at 37 °C and a 5% CO2 atmosphere.
Co-transfect HCT116 cells (8.0 × 105 cells, approximately 60% confluent on 6-well plates) with 1 μg of pGP-CMV-GCaMP6f and 100 ng of pCMV-myc-wtSeipin or pCMV-myc-ngSeipin in 300 μL of Opti-MEM and 10 μL of polyethylenimine max solution (1 mg/mL in MilliQ water). HCT116 cells proliferate faster than SH-SY5Y cells; accordingly, they should be less confluent during transfection than SH-SY5Y cells.
Detection and acquisition of images of cells expressing reporter protein
Twenty-eight hours after transfection, observe the cells under an inverted fluorescence microscope (e.g., Olympus IX-71-22TFL/PH) with an appropriate exposure time. If phenol red contained in DMEM causes a strong background signal, the volume of DMEM should be accordingly reduced. If a sufficient number of cells is present on the dish (approximately 95%) and transfection efficiency is sufficient (approximately 40%–50%), images should be taken using a 20× objective lens. This will produce images that contain approximately 150–200 fluorescent cells per image (Figure 5A).
Add 1 µM thapsigargin, an inhibitor of SERCA (Lytton et al., 1991), to the cells to evoke the leakage of calcium ions stored in the ER to the cytoplasm.
Quantification of fluorescence by ImageJ (Figure 5B, C)
Quantify the images according to the procedures described in step A3.
Statistical analysis on Microsoft Excel (Figure 5B, C)
Perform statistical analysis according to the procedures described in step A4.
Figure 5. Calcium ion concentrations in the cytoplasm of HCT116 cells expressing wtSeipin or ngSeipin before or after treatment with thapsigargin. A. HCT116 cells seeded on 6-well plates were co-transfected with 1 μg of pGP-CMV-GCaMP6f and 100 ng of pCMV-myc-wtSeipin or pCMV-myc-ngSeipin. Twenty-eight hours later, cells were treated with or without 1 μM thapsigargin (Tg), which inhibits the sarcoplasmic calcium ATPase (SERCA) (Lytton et al., 1991) and thereby evokes calcium ion leakage from the endoplasmic reticulum (ER) to cytoplasm. 0, 60, and 300 s later, cells were observed for fluorescence. Scale bar, 100 μm. B. For both cell types, images were taken every 30 s after treatment with Tg, and results from three experiments were analyzed according to the procedures described in Figure 2 and statistically processed according to the procedures described in Figure 3. The results are expressed as relative values, with the mean of measurements of cells expressing myc-wtSeipin in 0 s set as 1, along with the standard deviation. C. To estimate the total amount of calcium ions released from the ER to cytoplasm by Tg treatment, the area below the line graph until the broken line (fluorescence intensity in 0 s) in B was calculated and expressed as relative values, with the mean of measurements of cells expressing myc-wtSeipin set as 1, along with the standard deviation and presence of significant differences (Student’s t-test, **: p < 0.01). The results show that ngSeipin decreases the calcium ions stored in the ER (Saito et al., 2022).
Notes
Although this experimental method can be applied to various types of cultured cells, the method for introducing plasmids to express fluorescent reporter proteins (e.g., scale of cell culture, reagent types, volume of plasmid to be introduced, length of incubation time after transfection) needs to be optimized for each cell type. If the chemical gene transfer techniques using Polyethylenimine Max or Lipofectamine® LTX are not effective, the use of viral vectors or electroporation should be considered.
Acknowledgments
The experimental method described here was used in Saito et al. (2022), and the data shown in Figures 1, 4, and 5 are reproduced from that paper. This work was financially supported by AMED-CREST, Japan (23gm1410005 to K.M.).
References
Berridge, M. J., Bootman, M. D. and Roderick, H. L. (2003). Calcium signalling: dynamics, homeostasis and remodelling. Nat Rev Mol Cell Biol 4(7): 517-529.
Berridge, M. J., Lipp, P. and Bootman, M. D. (2000). The versatility and universality of calcium signalling. Nat Rev Mol Cell Biol 1(1): 11-21.
Brini, M. and Carafoli, E. (2009). Calcium pumps in health and disease. Physiol Rev 89(4): 1341-1378.
Chen, T. W., Wardill, T. J., Sun, Y., Pulver, S. R., Renninger, S. L., Baohan, A., Schreiter, E. R., Kerr, R. A., Orger, M. B., Jayaraman, V., et al. (2013). Ultrasensitive fluorescent proteins for imaging neuronal activity. Nature 499(7458): 295-300.
Delrio-Lorenzo, A., Rojo-Ruiz, J., Alonso, M. T. and Garcia-Sancho, J. (2020). Sarcoplasmic reticulum Ca2+ decreases with age and correlates with the decline in muscle function in Drosophila. J Cell Sci 133(6): jcs240879.
Ellgaard, L. and Helenius, A. (2003). Quality control in the endoplasmic reticulum. Nat Rev Mol Cell Biol 4(3): 181-191.
Hofer, A. M. and Machen, T. E. (1993). Technique for in situ measurement of calcium in intracellular inositol 1,4,5-trisphosphate-sensitive stores using the fluorescent indicator mag-fura-2. Proc Natl Acad Sci U S A 90(7): 2598-2602.
Laude, A. J. and Simpson, A. W. (2009). Compartmentalized signalling: Ca2+ compartments, microdomains and the many facets of Ca2+ signalling. FEBS J 276(7): 1800-1816.
Lytton, J., Westlin, M. and Hanley, M. R. (1991). Thapsigargin inhibits the sarcoplasmic or endoplasmic reticulum Ca-ATPase family of calcium pumps. J Biol Chem 266(26): 17067-17071.
Preissler, S., Rato, C., Yan, Y., Perera, L. A., Czako, A. and Ron, D. (2020). Calcium depletion challenges endoplasmic reticulum proteostasis by destabilising BiP-substrate complexes. eLife 9: e62601.
Saito, S., Ishikawa, T., Ninagawa, S., Okada, T. and Mori, K. (2022). A motor neuron disease-associated mutation produces non-glycosylated Seipin that induces ER stress and apoptosis by inactivating SERCA2b. eLife 11: e74805.
Schrank, S., Barrington, N. and Stutzmann, G. E. (2020). Calcium-Handling Defects and Neurodegenerative Disease. Cold Spring Harb Perspect Biol 12(7): a035212.
Suzuki, J., Kanemaru, K., Ishii, K., Ohkura, M., Okubo, Y., Iino, M. (2014). Imaging intraorganellar Ca2+ at subcellular resolution using CEPIA. Nat Commun 5: 4153.
Tsien, R. Y. (1981). A non-disruptive technique for loading calcium buffers and indicators into cells. Nature 290(5806):527-528.
Article Information
Copyright
© 2023 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 imaging > Fluorescence
Biochemistry > Other compound > Ion
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Quantification of Chromosomal Aberrations in Mammalian Cells
IP Inés Paniagua
JJ Jacqueline J. L. Jacobs
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4739 Views: 1218
Reviewed by: Khyati Hitesh ShahAmit Kumar Dey Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Sep 2022
Abstract
Maintenance of genome integrity requires efficient and faithful resolution of DNA breaks and DNA replication obstacles. Dysfunctions in any of the processes orchestrating such resolution can lead to chromosomal instability, which appears as numerical and structural chromosome aberrations. Conventional cytogenetics remains as the golden standard method to detect naturally occurring chromosomal aberrations or those resulting from the treatment with genotoxic drugs. However, the success of cytogenetic studies depends on having high-quality chromosome spreads, which has been proven to be particularly challenging. Moreover, a lack of scoring guidelines and standardized methods for treating cells with genotoxic agents contribute to significant variability amongst different studies. Here, we report a simple and effective method for obtaining well-spread chromosomes from mammalian cells for the analysis of chromosomal aberrations. In this method, cells are (1) arrested in metaphase (when chromosome morphology is clearest), (2) swollen in hypotonic solution, (3) fixed before being dropped onto microscope slides, and (4) stained with DNA dyes to visualize the chromosomes. Metaphase chromosomes are then analyzed using high-resolution microscopy. We also provide examples, representative images, and useful guidelines to facilitate the scoring of the different chromosomal aberrations. This method can be used for the diagnosis of genetic diseases, as well as for cancer studies, by identifying chromosomal defects and providing insight into the cellular processes that influence chromosome integrity.
Graphical overview
Keywords: Chromosomal instability Mitosis DNA repair DNA replication Genome integrity Cancer Congenital malignancies
Background
To maintain a stable genome, cells must accurately duplicate their genetic material before each cell division and ensure efficient signaling and repair of DNA damage. Chromosomal instability (CIN) is a form of genomic instability observed in cancer and many congenital abnormalities, typically associated with numerical or structural chromosome changes (Bakhoum and Cantley, 2018). While numerical CIN is characterized by gain and/or loss of whole chromosomes, structural CIN is characterized by gain, loss, and/or rearrangements of parts of chromosomes. Although significant advances in the detection of CIN have been made with the appearance of quantitative high-throughput imaging cytometry and single-cell genomics, classical cytogenetics still remains as an important method to detect chromosomal aberrations within research and clinical settings (Lepage et al., 2019). Cytogenetic approaches generally involve analyzing metaphase chromosomes stained by DNA dyes such as GIEMSA or DAPI; however, inconsistency in the preparation of chromosome spreads is a major problem. Moreover, the scoring of chromosomal aberrations usually relies on specialized experience, since chromosome spreading artifacts can be easily misinterpreted as genomic changes. Thus, efforts should be made to standardize the assays and refine the analysis of chromosomal aberrations for data interpretation. Here, we report a method that enables fast and reliable preparation of metaphase chromosome spreads from mammalian cells for the purpose of scoring chromosomal aberrations. In brief, cells are treated with a metaphase-arresting substance, harvested, and stained with DNA dyes. Metaphase cells are then analyzed microscopically for the presence of chromosomal aberrations. This method can be used to score gross (i.e., observable with standard staining methods) structural aberrations occurring naturally and following exposure to genotoxic chemicals, such as hydroxyurea, aphidicolin, mitomycin C, etc. While numerical aberrations can also be scored, other methods may be more appropriate for that. In addition, we provide representative images of normal and aberrant metaphases as well as scoring guidelines to facilitate accurate identification and evaluation of chromosomal instability, which is of outmost importance within both research and clinical settings.
Materials and reagents
Gloves and lab coat
10 or 15 cm Petri dish (Greiner or Thermo Fisher Scientific, catalog numbers: 664160 or 168381)
15 and 50 mL screw cap tubes (Sarstedt, catalog numbers: 62554502 and 62547254)
Microscope glass slides with a frosted end (Epredia, catalog number: AB00000112EO1MNZ10)
Glass coverslips (24 mm × 50 mm) (VWR, catalog number: 631-1574)
Paper towel
Mammalian cells of interest and appropriate cell culture medium [suggested complete growth medium: Dulbecco’s modified Eagle’s medium (DMEM, Thermo Fisher Scientific, Gibco, catalog number: 41966-029), supplemented with 10% fetal bovine serum (FBS, Capricorn, catalog number: FBS-12A), 100 U/mL penicillin, 100 μg/mL streptomycin (Thermo Fisher Scientific, Gibco, catalog number: 15140-122), and 2 mM L-Glutamine (VWR, catalog number: 392-0441)]
Note: This protocol is for adherent cells.
Phosphate-buffered saline (PBS) (Thermo Fisher Scientific, Gibco, catalog number: 14190144)
Trypsin-EDTA 0.05% (Thermo Fisher Scientific, Gibco, catalog number: 25300-054), store at 4 °C
Fetal bovine serum (FBS) (Capricorn Scientific, suggested catalog number: FBS-12A)
(Optional) Genotoxic chemical e.g., hydroxyurea (Sigma, catalog number: H8627)
Colcemid, KaryoMax colcemid solution, 10 μg/mL (Gibco, catalog number: 15212012), store at 4 °C
Potassium chloride (KCl) (Sigma, catalog number: 1049360500)
Dimethyl sulfoxide (DMSO) (Sigma, catalog number: 34943)
ProLong Gold antifade reagent with DAPI (Thermo Fisher Scientific, catalog number: P36931)
Transparent nail polish (any brand)
Plastic Pasteur pipette (VWR, catalog number: 612-1684P)
Methanol (Honeywell, catalog number: 32213)
Glacial acetic acid (Sigma, catalog number: A6283)
Hydroxyurea (see Recipes)
Hypotonic solution (see Recipes)
Fixative solution (see Recipes)
Equipment
Pipettes (Gilson)
Vortexer
Centrifuge with swing bucket rotor (e.g., Rotina 380, Hettich)
Heating block (Stretching Table OTS 40, Medite, catalog number: 401520)
Water bath (GFL Shaking Water Baths, catalog number: 1083)
Imaging equipment:
Option 1:
Metafer4/MSearch automated metaphase finder system (MetaSystems) equipped with a Zeiss AxioImager Z2 microscope (Carl Zeiss)
Objectives: 10×/0.45 and 63×/1.40 oil Plan-ApoChromat
Filterset: DAPI (Zeiss Filterset 49, ACR)
Camera: CoolCube1
Option 2:
Wide field microscope e.g., Zeiss AxioObserver Z1 microscope
Objective: 63×/1.40 oil Plan-ApoChromat
Filterset: DAPI (Zeiss Filterset 49, ACR)
Camera: ORCA–Flash4.0 V3 Digital CMOS: C13440-20 CU
Software
MetaSystems (version 3.11) software
Fiji (version 2.0) software
GraphPad Prism (version 9) software
Procedure
Preparation of cultures
Grow cells in standard culture conditions (e.g., at 37 °C in a humidified atmosphere with 5% CO2).
Seed 0.5 × 106 cells into a 10 cm dish 48 h before harvesting, to be ~70% confluent at the time of harvest.
Notes:
Adjust the seeding density according to the doubling time of the cell line. You are aiming for your cells to go through approximately 1.5 cell cycles (since many aberrations are lethal or lost during subsequent cell divisions, so they are best observed at the first or second metaphase).
When working with slow dividing cells, it is better to use a 15 cm dish and seed 1 × 106 cells into it. This will result in a larger cell pellet and a higher percentage of mitotic cells at the time of harvest.
Since the harvesting of the cells is a delicate process, never harvest more than 6–8 samples at once. When you have more samples, harvest them in several rounds. One harvesting round can take up to 1.5–2 h.
(Optional) Alternatively, at the time of seeding, treat cells with a genotoxic chemical (e.g., 4 mM hydroxyurea) for 3–6 h, wash two times with PBS, and culture cells in complete growth medium for approximately 1.5 cell cycles.
Note: When using a 1 M stock of hydroxyurea, add 40 μL of hydroxyurea to 10 mL of medium (for 10 cm dishes).
Arresting cells in metaphase
On the day of harvest, check whether your cells have the right confluency.
Two hours before harvest, add colcemid directly to the medium to a final concentration of 0.2 μg/mL. When using a 10 μg/mL stock of colcemid, add 200 μL of colcemid to 10 mL of medium (for 10 cm dishes).
Notes:
Avoid incubation times longer than 2 h, since prolonged exposure to colcemid results in very compacted chromosomes.
Volumes indicated in the protocol are for a 10 cm dish; adjust accordingly for larger dishes. For instance, add 400 μL of colcemid to 20 mL of medium when using 15 cm dishes.
During the colcemid incubation, label 15 mL screw cap tubes for the different conditions, prepare and prewarm the hypotonic solution (see Recipes) at 37 °C, and prepare ice-cold fixative solution (see Recipes) (keep at -20 °C).
After 2 h of colcemid treatment, the effect should be visible: many cells will look rounded, refractile, and appear as if they are about to detach.
Harvesting of cells
Note: Cells in mitosis round up and lose attachment to the plate. To harvest all mitotic cells (including those floating), save the media and/or the PBS wash as well as the media used to neutralize the trypsin, and collect in a single screw cap tube.
Remove the media.
(Optional) Alternatively, save the media in a 15 mL screw cap tube.
Carefully wash the cells with 5 mL of PBS and recapture them by collecting the PBS in a 15 mL screw cap tube.
Trypsinize and collect the cells in medium containing FBS. Add to the 15 mL tube containing the PBS wash.
Note: To trypsinize the cells, add 1 mL of trypsin to the dish and incubate for 3–5 min at 37 °C until all cells are loosely floating (this can be seen under a microscope). To neutralize the trypsin, add 7 mL of medium to the dish. Collect cells by resuspending.
Spin the samples for 5 min at 186× g at room temperature (RT).
Remove the supernatant completely and carefully tap to the side of the tube so your pellet will be loose.
GENTLY resuspend your cells in 10 mL of hypotonic solution (prewarmed to 37 °C). Pipette the solution onto the tube wall.
Note: The addition of KCl swells the cells, so pipette carefully to prevent cell lysis.
Incubate the cells in the hypotonic solution for 7 min at 37 °C. Slowly invert the tube several times during the incubation to prevent clumping.
Note: Incubation times may differ per cell line and thus need to be determined empirically. For commonly used cell lines, such as HeLa, MEFs, and RPE1-hTERT, 5–10 min of treatment time is sufficient.
Spin the cells for 5 min at 186× g at RT.
Fixation of cells
Decant the KCl and tap the tube to resuspend the cells in the small volume of KCl that remains (approximately 0.5 mL).
Drop by drop, add 1 mL of ice-cold fixative solution while the cells are slowly and gently being mixed on a vortex.
Fill to 10 mL with the fixative and store at 4 °C overnight or longer; cells can be kept at this stage for months.
Metaphase chromosome spread preparation
Note: For additional details pertaining to the procedure, please consult Video 1.
Video 1. Metaphase chromosome spread preparation. This video shows the procedure to prepare metaphase chromosome spreads, as described in section E, including the fixation and dropping of the cells onto microscope slides.
Spin the cells for 5 min at 186× g at RT.
Aspirate the fixative until 0.5–1 mL is left. Resuspend by tapping. Put the tubes on ice.
Note: Starting with 0.5 × 106 cells seeded in a 10 cm dish that reach 70% confluency at time of harvest yields a cell pellet that, when resuspended in 0.5–1 mL fixative, results in an adequately high cell density for chromosome spread preparation. However, the volume in which you resuspend the cells may be adjusted to change the concentration of the sample and obtain a lower or higher density of metaphases on the slides in subsequent steps.
Take some (old) pre-cooled fixative solution and put it on ice as well.
Label microscope glass slides for the different conditions (using a pencil since fixative will dissolve the marker). Place the slides in a beaker containing cold water to pre-cool and wet them.
Prepare a humidified 42 °C heating block (place wet paper towels on top of a heating block set to 42 °C).
Place a piece of cloth on the ground to collect splashes. Then, lay down a 50 mL screw cap tube and place a water-wetted slide against it, tilted at a 45° angle.
Using a plastic Pasteur pipette (the tip is wider than a P1000 pipette tip), pipette up and down and take up the cell suspension from step E2. Let the first drops fall into the tube. Then, drop several drops of cell suspension onto different places on a slide whilst standing up. Aim to cover all parts of the slide with each drop but avoid dropping cells on the same spot.
Note: Slide dropping from a distance (approximately 0.5 m) lets the nuclei fall apart on the slide, so the metaphases will spread. However, make sure that the distance is not too large, since the chromosomes will spread too far apart from each other, which will make it difficult to determine if they are from one or more cells.
Immediately after dropping, quickly wash by dropping fixative solution across the slide using a plastic Pasteur pipette and place the slides (cell side up) for 3 min on a humidified 42 °C heating block.
Note: The high temperature and the vapor that is continuously released from the humidified heating block improves the quality of the spreads (Deng et al., 2003).
Check the slides under a regular light microscope for spreading efficiency. You should see many nuclei and some metaphase chromosomes (Figure 1).
Note: If you can find 3–5 metaphases in 1 min, there is probably plenty present on the slide. If there are less, you can drop some extra. If too many, dilute sample with fixative and repeat steps E2–E9.
Air dry the slides in the fume hood for a few hours at RT.
To mount the slides, add a few drops of Prolong Gold antifade with DAPI to the coverslip, pick up the coverslip using the slide, and slowly lay flat. Avoid air bubbles. Gently press the coverslip with a tissue to remove excess mounting medium.
Seal with nail polish.
Slides can be stored up to one week at 4 °C or at -20 °C for longer storage in the dark. The remainder of the sample in the 15 mL tube can be stored in fresh fixative at 4 °C for months.
Figure 1. Representative metaphase spreads in a cancer cell line (HeLa) at 4× and 10× magnification. After dropping cells onto microscope slides, many cytoplasm-free nuclei, and some metaphase spreads (red arrows) should be visible. At low magnification (4×, left panel), metaphase chromosomes should look like small black dots; at higher magnification (10×, right panel), the arms of the chromosomes should be visible.
Data analysis
Image acquisition and processing
Digital images of metaphases are captured using the Metafer4/MSearch automated metaphase finder system equipped with an AxioImager Z2 microscope and DAPI filter. In this case, the user reviews the images taken by the automated microscope and selects those metaphases that appear suitable for analysis. This method greatly reduces slide reading time and is thus recommended. Alternatively, metaphases can be manually selected and imaged using a high-resolution microscope, such as a Zeiss AxioObserver Z1 microscope. For this method, though, it is important to establish a systematic way of scanning slides to locate the metaphases (e.g., scanning from side to side or along the length of the slide), to avoid imaging and scoring the same metaphase twice.
At least 50 readable metaphases should be imaged at high magnification (63×, oil immersion). Readable metaphases (i.e., metaphases that are suitable for analysis) are identified by the following criteria:
• Chromosome number within the diploid chromosome number of the working cell line.
• Well-spread chromosomes, with minimal overlap of chromosomes and chromosomes arms.
• Clear and defined chromosome structure, with intact centromeres.
If the required total of 50 metaphases is not obtained, then additional slides should be prepared from the reserved fixed sample.
Representative images and scoring
Chromosomal aberrations are quantified from at least 50 metaphase spreads per condition per experiment. Analysis of 50 metaphases per condition and replicate is sufficient to obtain a reliable mean. Data are presented as the average percentage of chromosomal aberrations per metaphase, showing the spread among the replicates of the individual experiments, as previously shown (Paniagua et al., 2022, Figure 3E). Statistical analysis is performed with the appropriate test for multiple comparisons using GraphPad Prism 9.
Classification
Chromosomal aberrations are classified into the following categories (Registre, 2016):
• Chromosome-type (both sister chromatids affected)
- Breaks/deletions (resulting from a DNA break that was not repaired)
- Exchanges [resulting from two or more DNA breaks with inappropriate rejoining/repair, within a single chromosome (intrachanges) or between chromosomes (interchanges)]
• Chromatid-type (only one sister chromatid affected)
- Breaks/deletions
- Exchanges
Additionally, other forms of chromosomal aberrations are occasionally observed; these are recorded but not included in the analysis because they do not necessarily involve chromosome breakage.
• Pulverized chromosome/metaphase (total loss of chromosome architectural integrity).
• Gaps (unstained regions on the chromosomes/chromatids, smaller than the width of one chromatid, and with minimal misalignment with the rest of the chromatid/chromosome).
• Polyploidy and/or other forms of aneuploidy.
It is important to be aware that chromatid-type breaks are the most frequently observed aberrations with a sampling time of approximately 1.5 cell cycles from the start of seeding or following drug exposure. After a further cell cycle, some of these chromatid-type aberrations can be converted into chromosome-type aberrations. This is because DNA double-stranded breaks forming and persisting through S-phase will be replicated and become evident as a chromatid-type aberration. If the chromatid-type is not repaired, then it will also be replicated, resulting in the formation of a chromosome-type aberration. For a more comprehensive view of the formation and scoring of chromosomal aberrations, we refer interested readers to an excellent review (Danford, 2012).
Representative examples of normal and aberrant chromosomes are shown in Figure 2. An example of data quantification is also provided in Table S1.
Data misinterpretation
The analysis of chromosomal aberrations has a subjective component; therefore, misinterpretation can occur and result in the (mis-)classification of a normal chromosome as aberrant. Generally, it is better to err on the side of caution and disregard a metaphase altogether if the apparent aberration is unclear. Some common examples for misinterpretations have been highlighted below:
• Crossing-over of sister chromatids can be mistaken for a dicentric chromosome.
• Chromosomes overlapping near centromeres can resemble a chromosome-type exchange (radial).
• Twisted or overlapping chromosomes can be wrongly scored as chromosome rings.
• Secondary constrictions in chromosomes can appear as chromosome gaps.
Figure 2. Representative examples of common chromosomal aberrations. Schematic illustration of the three common categories of chromosomal aberrations. Representative metaphase spreads, with normal and aberrant chromosomes in a cancer cell line (HeLa) at 63× magnification, are shown below for each category. A. Gaps: dislocation of the chromatid or chromosome arm with no misalignment. B. Breaks: dislocation of the chromatid or chromosome arm, with misalignment, and an unstained region that is wider than the chromatid width. The insert in the lower right corner is derived from an independent metaphase and shows a different example of how chromosomal breaks may appear. C. (Inter-)exchanges: exchanges involving two chromosomes. Examples of quadriradial figures are shown for the chromatid-type exchanges. Scale bar, 10 μm.
Notes
Particular attention should be given to standardizing the conditions of swelling, fixation, and slide dropping, so that high quality preparations can be made regularly. Achieving a high mitotic index (total number of metaphases/number of nuclei) is equally important to maximize the sensitivity to genotoxic agents and reduce the slide reading time. To increase the mitotic index at harvest, one can experiment with the cell culture dishes and/or the cell culture density (confluency should always be avoided for cells growing in monolayers).
In cell lines with high levels of chromosomal instability and in some cancer cell lines, a basal level of chromosomal aberrations can be detected.
Certain genotoxic treatments, including hydroxyurea, have the potential to alter the chromosomal copy number of the treated cells. Thus, it is recommended to control for copy number variation (CNV) by counting the total number of chromosomes per metaphase. In case the manipulation results in CNV, the quantification of chromosomal aberrations should then be plotted as chromosomal aberrations per total number of chromosomes (in contrast to the standard plotting of chromosomal aberrations per metaphase).
Recipes
Hydroxyurea
1 M in DMSO
Dissolve 0.38025 g of hydroxyurea in 5 mL of DMSO
Aliquot in small volumes (0.5 mL) and store at -20 °C for up to four months
Hypotonic solution
0.075 M KCl in dH2O
Prewarm the required amount of 0.075 M KCl in a 37 °C water bath before each use
Fixative solution
Three parts methanol
One part glacial acetic acid
Make fresh each time
Use ice-cold
Acknowledgments
We thank Daniëlle Koot for assistance with the preparation of audiovisual material. I.P. was supported by project grant 11905/2018-2 from the Dutch Cancer Society (KWF), awarded to J.J.L.J. Work in the Jacobs lab was supported by an institutional grant of the Dutch Cancer Society and of the Dutch Ministry of Health, Welfare and Sport. This protocol was adapted from procedures published by (Álvarez-Quilón et al., 2014; Xu et al., 2017; Hustedt et al., 2019; Mukherjee et al., 2019). Figures were created with BioRender.com.
Competing interests
The authors declare no competing interests.
References
Álvarez-Quilón, A., Serrano-Benítez, A., Lieberman, J. A., Quintero, C., Sánchez-Gutiérrez, D., Escudero, L. M. and Cortés -Ledesma, F. (2014). ATM specifically mediates repair of double-strand breaks with blocked DNA ends. Nat Commun 5: 3347.
Bakhoum, S. F. and Cantley, L. C. (2018). The Multifaceted Role of Chromosomal Instability in Cancer and Its Microenvironment. Cell 174(6): 1347-1360.
Danford, N. (2012). The interpretation and analysis of cytogenetic data. Methods Mol Biol 817: 93-120.
Deng, W., Tsao, S. W., Lucas, J. N., Leung, C. S. and Cheung, A. L. (2003). A new method for improving metaphase chromosome spreading. Cytometry A 51(1): 46-51.
Hustedt, N., Saito, Y., Zimmermann, M., Álvarez-Quilón, A., Setiaputra, D., Adam, S., McEwan, A., Yuan, J. Y., Olivieri, M., Zhao, Y., et al. (2019). Control of homologous recombination by the HROB-MCM8-MCM9 pathway. Genes Dev 33(19-20): 1397-1415.
Lepage, C. C., Morden, C. R., Palmer, M. C. L., Nachtigal, M. W. and McManus, K. J. (2019). Detecting Chromosome Instability in Cancer: Approaches to Resolve Cell-to-Cell Heterogeneity. Cancers (Basel) 11(2): 226.
Mukherjee, C., Tripathi, V., Manolika, E. M., Heijink, A. M., Ricci, G., Merzouk, S., de Boer, H. R., Demmers, J., van Vugt, M. and Ray Chaudhuri, A. (2019). RIF1 promotes replication fork protection and efficient restart to maintain genome stability. Nat Commun 10(1): 3287.
Paniagua, I., Tayeh, Z., Falcone, M., Hernández Pérez, S., Cerutti, A. and Jacobs, J. J. L. (2022). MAD2L2 promotes replication fork protection and recovery in a shieldin-independent and REV3L-dependent manner. Nat Commun 13(1): 5167.
Registre, M. P., R. (2016). The In Vitro Chromosome Aberration Test. In: Haley, M. (Ed.). Genetic Toxicology Testing. A Laboratory Manual (pp. 207-267). Academic Press.
Xu, S., Wu, X., Wu, L., Castillo, A., Liu, J., Atkinson, E., Paul, A., Su, D., Schlacher, K., Komatsu, Y., et al. (2017). Abro1 maintains genome stability and limits replication stress by protecting replication fork stability. Genes Dev 31(14): 1469-1482.
Supplementary information
The following supporting information can be downloaded at here:
Table S1. Example data quantification
Article Information
Copyright
© 2023 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
Cancer Biology > Genome instability & mutation > Cell biology assays
Molecular Biology > DNA > DNA damage and repair
Molecular Biology > DNA > DNA structure
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Peer-reviewed
Quantification of Botrytis cinerea Growth in Arabidopsis thaliana
PS Patricia Scholz
KC Kent D. Chapman
TI Till Ischebeck
AG Athanas Guzha
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4740 Views: 1063
Reviewed by: Shweta PanchalAntony Chettoor Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant Physiology Jul 2022
Abstract
Yield losses attributed to plant pathogens pose a serious threat to plant productivity and food security. Botrytis cinerea is one of the most devastating plant pathogens, infecting a wide array of plant species; it has also been established as a model organism to study plant–pathogen interactions. In this context, development of different assays to follow the relative success of B. cinerea infections is required. Here, we describe two methods to quantify B. cinerea development in Arabidopsis thaliana genotypes through measurements of lesion development and quantification of fungal genomic DNA in infected tissues. This provides two independent techniques that are useful in assessing the susceptibility or tolerance of different Arabidopsis genotypes to B. cinerea.
Key features
• Protocol for the propagation of the necrotrophic plant pathogen fungus Botrytis cinerea and spore production.
• Two methods of Arabidopsis thaliana infection with the pathogen using droplet and spray inoculation.
• Two readouts, either by measuring lesion size or by the quantification of fungal DNA using quantitative PCR.
• The two methods are applicable across plant species susceptible the B. cinerea.
Graphical overview
A simplified overview of the droplet and spray infection methods used for the determination of B. cinerea growth in different Arabidopsis genotypes
Keywords: Arabidopsis thaliana Botrytis cinerea Fungal pathogen Pathogen assay Quantitative PCR
Background
Agricultural productivity is constantly under threat from plant pathogens including bacteria, fungi, oomycetes, and viruses, contributing to global losses of 10%–28% in crop production to pests (Savary et al., 2019; IPPC Secretariat, 2021). Among the pathogens, Botrytis cinerea is a necrotrophic fungal phytopathogen distributed worldwide that causes great damage to a large diversity of plants, including various crops, leading to its classification among the top 10 worst fungal pathogens (Dean et al., 2012; Caseys et al., 2021). B. cinerea is considered a generalist necrotroph that kills the host tissues and takes up nutrients from the dead tissue. Unlike other plant pathogens, it has a very broad host range, affecting more than 1,000 plant species (Elad et al., 2016; Caseys et al., 2021; Bi et al., 2022).
The strong economic impact with regard to yield loss and the broad range of plant hosts make B. cinerea an interesting model organism for plant pathogen research, especially as it also infects the model plant species Arabidopsis thaliana. This allowed, for example, extensive transcriptomic analysis of the necrotroph infection (Windram et al., 2012) or studies on how the genetic variation of Arabidopsis or B. cinerea influences disease outcomes (Kliebenstein et al., 2005; Rowe and Kliebenstein, 2008; Soltis et al., 2020). Furthermore, B. cinerea infection assays can be used to observe different aspects of the plant immune responses in Arabidopsis (Genenncher et al., 2016; Guzha et al., 2022). Here, we show two straightforward methods that can be used in parallel to assess the disease tolerance of different Arabidopsis genotypes. One method involves drop inoculation of B. cinerea spores and measuring the average lesion development on the infected leaves (Genenncher et al., 2016; Guzha et al., 2022). The other approach involves the quantification of B. cinerea genomic DNA (gDNA) in infected Arabidopsis tissues (Ettenauer et al., 2014; Guzha et al., 2022). These two methods provided are versatile and can be applied to different plant species susceptible to the pathogen with minimal modifications.
Materials and reagents
Biological materials
Arabidopsis seeds
B. cinerea strain B05-10
Reagents
Vogel buffer
Potato dextrose agar (PDA) (Merck, catalog number: 70139)
Potato dextrose broth (PDB) (Merck, catalog number: P6685)
Glycerol (Thermo Scientific, catalog number: 17904)
Plant/Fungi DNA Isolation kit (Norgen Biotek Corporation, catalog number: 26200)
Takyon No Rox SYBR master mix dTTP Blue (Eurogentec UF-NSMT-B0701)
B. cinerea ACTIN forward primer 5′-TGGAGATGAAGCGCAATCCA-3′ (GenewizTM)
B. cinerea ACTIN reverse primer 5′-AAGCGTAAAGGGAGAGGACG-3′ (GenewizTM)
B. cinerea TUBULIN forward primer 5′-CCGTCATGTCCGGTGTTAC-3′ (GenewizTM)
B. cinerea TUBULIN reverse primer 5′-CGACCGTTACGGAAATCGG-3′ (GenewizTM)
Solutions
Potato dextrose agar (PDA) and potato dextrose broth (PDB) (see Recipes)
Sterile Vogel buffer (see Recipes)
70% ethanol (see Recipes)
15% glycerol (see Recipes)
Recipes
Potato dextrose agar (PDA) and potato dextrose broth (PDB)
Potato dextrose agar (PDA)
For 1 L of PDA, weigh 39 g of commercial PDA powder (contains 20 g of dextrose, 15 g of agar, and 4 g of potato extract), and suspend in 1 L of deionised water. Autoclave the solution at 121 °C for 20 min.
Potato dextrose broth (PDB)
For 1 L of PDB, weigh 24 g of commercial PDB powder (contains 20 g of dextrose, 4 g of potato extract) and dissolve in 1 L of deionised water. Autoclave at 121 °C for 20 min.
Sterile Vogel buffer
For 1 L of Vogel buffer, weigh the compounds as shown in Table 1 and fill up to 900 mL with distilled water. Mix until all compounds are dissolved.
Table 1. Preparation of Vogel buffer
Constituent Amount
Sucrose 15 g
Trisodium citrate·2H2O 2.5 g
K2HPO4 5 g
MgSO4·7H2O 0.2 g
CaCl2·2H2O 0.1 g
NH4NO3 2 g
Adjust to pH 6.0 and fill up the volume to 1 L. Sterilise the buffer by autoclaving.
70% ethanol
Reagent Final concentration Amount
Ethanol (absolute) 70% 700 mL
H2O n/a 300 mL
Total n/a 1,000 mL
15% glycerol
Reagent Final concentration Amount
Glycerol 15% 150 mL
H2O n/a 850 mL
Total n/a 1,000 mL
Sterilize by autoclaving.
Laboratory supplies
10 cm plant pots
25 cm × 51 cm × 6 cm propagation trays
25 cm × 51 cm × 18 cm high standard dome
Arabidopsis soil: Fruhstofer Erde
Chemical-resistant gloves
Miracloth (Merck, catalog number: 475855)
Kimwipes® (Kimtech Science, catalog number: 34120)
2 mL microcentrifuge tube (Sarstedt, catalog number: 72.695.500)
1.5 mL microcentrifuge tube (Sarstedt, catalog number: 72.690.001)
Flat cap PCR tubes (Sarstedt 72.985.002 (tubes) and 65.989.002 (lids))
60 mL glass spray bottle (Uline, catalog number: S24562A)
Equipment
Growth chamber with light and temperature controls (Percival Scientific Inc.)
Oven for sterilisation of soil
Microcentrifuge
Micropipettes
Scalpel
Mortar and pestle
MarCal 16ER digital caliper (Mahr, Göttingen, Germany)
Bio-Rad iQ5 Multicolor Real-Time PCR Detection System
NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific)
Fuchs-Rosenthal counting chamber (Paul Marienfeld, Lauda-Königshofen, Germany)
Software and datasets
MarCom Standard Software for MarCal digital caliper
iQ5 Optical System Software (Bio-Rad Laboratories GmbH-Munich, Germany)
Microsoft Excel
Procedure
Preparation of fungal spores
When working with spores of B. cinerea, take care to avoid any infection of Arabidopsis plants. Keep working areas for B. cinerea and Arabidopsis plants distant from each other and avoid working with plants after you prepared B. cinerea spores (see Notes).
Inoculate 5 μL of spore suspension onto the centre of PDA plates at room temperature under sterile conditions. Incubate the closed plate with spores at room temperature. Do not seal plates until mycelium on the agar is covered with grey spores (4–5 days).
Propagate the spores by transferring clean mycelium grown on the PDA plate to new PDA plates. To that end, cut out a small piece of mycelium-covered agar (1 cm × 1 cm) using a sterile scalpel and transfer it upside down on the new plate. Take care to transfer only areas with a clean layer of B. cinerea mycelium that does not contain any contaminations. An example of a clean B. cinerea culture can be found in Vasquez-Montaño et al. (2020). For long-term storage of spores, harvest the spores from B. cinerea growing for 10 days on PDB plates without contaminations. In case of contaminations, cut out a section in the media with clean hyphae and grow on fresh PDA plates until a clean culture is obtained as described in step A2. Scrape spores from plates using a sterile scalpel before resuspending them with 20 mL of 15% sterile glycerol and filter through Miracloth.
Estimate the harvested spore counts using a Fuchs-Rosenthal counting chamber.
Clean the counting chamber and the coverslip with 70% ethanol. Put the glass cover on the counting chamber. Vortex the spore suspensions and pipette at least 10 μL aliquot slowly against the edge of the coverslip so that the counting chamber is filled by capillary force.
The Fuchs-Rosenthal counting chamber has a grid pattern of 16 squares of 1 mm × 1 mm that are subdivided further into 16 squares of 0.25 mm × 0.25 mm (see Figure 1) and a depth of 0.2 mm. Count the spore numbers in the 1 mm × 1 mm square areas. For one aliquot, count 16 square millimetre areas and calculate the average spore number.
Figure 1. Grid of a Fuchs-Rosenthal counting chamber. Sixteen areas of 1 mm are delimited by triple lines and further subdivided into 0.25 mm × 0.25 mm squares by single lines. The counting chambers depth is 0.2 mm, so a 1 mm area corresponds to 0.2 μL or 0.0002 mL.
Count three independent aliquots of 10 μL from the spore suspension in 15% glycerol and calculate the average spore number per square millimetre area. Vortex the spore suspension before each aliquot to obtain a homogeneous suspension. Calculate the spore concentration as follows:
Adjust the concentration to 2 × 106 spores/mL in 15% glycerol. If the spore concentration is too low, centrifuge the spore suspension and remove some of the supernatant liquid. Determine the spore concentration again (as described in step A3) and then adjust to a final concentration of 2 × 106 spores/mL.
Prepare 1 mL aliquots from this spore suspension for storage. While preparing the aliquots, take care to keep the spores homogeneously suspended by regularly vortexing. Flash-freeze the aliquots in liquid nitrogen and store at -80 °C.
Plant growth
To facilitate the accurate comparison of Arabidopsis genotypes, the plants must be grown under identical conditions and preferably in the same trays, which are rotated at least once every week. As an example, the procedure used in Guzha et al. (2022) and Jacobs et al. (2017) that allow comparisons of Arabidopsis genotypes' susceptibility to plant pathogens is outlined below. In addition, check the Notes section for comments on biological replicates.
Prepare semi-sterile soil sufficient for the desired number of pots by heating it in an oven at 80 °C for 8 h.
Transfer enough soil to fill square 10 cm pots and water the pots by placing water in the trays until the soil reaches saturation, before draining excess water. Place 6–10 Arabidopsis seeds in the four corners and the centre of the pots, which will be thinned out later to leave one plant per station (Figure 2A). Cover with plastic dome.
After stratification at 4 °C, transfer to growth cabinet with short day conditions (8 h light and 16 h darkness) at 22 °C, a relative humidity of 65%, and a light intensity of 120–140 μmol m-2·s-1. Remove the plastic dome after germination of the seeds.
Grow plants for 7–10 days; then, thin out excess seedlings to leave one plant per position (Figure 2A).
Grow until desired stage.
Droplet infection and measurements of lesion diameters
Take care not to infect other plants that are not part of the infection experiment. Keep infected plants separate from the normal plant growth chambers and avoid working with other plants after carrying out infection assays.
Prepare spores by gently thawing the frozen spores on ice. Dilute the thawed spore suspension to a final concentration of 5 × 104 spores/mL in sterile Vogel buffer.
Let the spores germinate (40 mL in a Falcon tube) for at least 4 h at 25 °C (shaking is not necessary). B. cinerea spores germinate readily in Vogel buffer. Germination of spores can be checked under a light microscope.
Use 6-week-old plants grown under short day conditions as described above (section B). For each leaf, place a single 6 μL droplet of the germinated spore suspension onto the adaxial side of the leaf, in the centre next to the middle vein (Figure 2B). Inoculate at least 30 fully expanded leaves of each plant genotype (at least six leaves from each plant in a pot, as shown in Figure 2A). Between application of spore suspension droplets, regularly vortex the spore suspensions to keep a homogeneous spore distribution (every 2–3 droplets). Spore suspension droplets will stay on the leaves due to surface tension; however, take care not to shake the pots to ensure that droplets do not fall off the leaf.
Figure 2. Arabidopsis plants for infection with Botrytis cinerea. A. For each genotype, five plants are grown per pot for use in the infection assays. B. At least 30 leaves per pot are inoculated with 6 μL droplets of a B. cinerea spore suspension. Three days post infection, lesions develop at the infected area.
Add 0.5 L of water to the tray containing the plants and cover them with a lid to maintain high humidity conditions. Return plants to short-day growth conditions (distinct from normal growth chambers) until determination of lesion size.
Analyse lesion size 3 days post infection. To that end, use the MarCal 16ER digital calliper to measure lesion diameters.
Before first use: install the MarCom Standard Software according to manufacturer’s instructions on the notebook you will subsequently use for measurements. Connect your digital calliper to the notebook and install the required software drivers. Use a notebook with Windows operating system that has MS Excel installed (see Notes).
For measurements, connect the digital calliper to the notebook with the installed MarCom software, open the software, and verify that the calliper has been recognised. Detach infected leaves from Arabidopsis plants and measure the diameter of the developed lesion with the calliper. Press the measurement button to transfer the data point to an Excel file, which will open after the first measurement.
Measure leaves from one plant genotype at a time and mark different genotypes in the Excel file with the measured data.
Spray infection and isolation of fungal gDNA
Another way to assess susceptibility of Arabidopsis genotypes to B. cinerea infection is by spray-infection and subsequent quantification of fungal gDNA in plant tissues. When using the spray infection protocol, take care not to infect other plants that are not part of the infection experiment through aerosols generated. Keep infected plants separate from the normal plant growth chambers and avoid working with other plants after carrying out infection assays.
Prepare spores by gently thawing the required number of aliquots on ice. Dilute the thawed spore suspension to a final concentration of 2 × 105 spores/mL in sterile Vogel buffer.
Let the spores germinate (50 mL in a Falcon tube) for at least 4 h at 25 °C. B. cinerea spores germinate readily in Vogel buffer.
Use 6-week-old plants grown under short day conditions as described above (section B). For spray infection, transfer the germinated spores to a spray bottle and spray evenly on the plants until liquid runs off the leaves.
Add 0.5 L water to the tray with the plants and cover them with a lid to keep humid conditions. Return plants to short-day growth conditions (distinct from the normal growth chambers) until harvest for gDNA isolation.
After three or four days, flash-freeze the rosettes in liquid nitrogen and store at -80 °C or proceed to isolation of plant and fungal gDNA.
For isolation of gDNA, the commercially available Plant/Fungi DNA Isolation kit was used according to manufacturer’s instructions. The isolation of gDNA included the following steps:
Keep the harvested plant material cooled with liquid nitrogen and grind into a powder using a mortar and pestle. Transfer ≤ 100 mg of the powder to a 1.5 mL microcentrifuge tube, add 500 μL of Lysis buffer and 1 μL of RNase A, and vortex for 20 s.
Note down the exact mass of powdered plant material for sample normalisation.
Lyse for 10 min at 65 °C, add 100 μL of binding buffer I, mix by inverting the tube, and incubate on ice for 5 min.
Transfer the lysate to a filter column, centrifuge at 20,000× g for 2 min, and transfer the supernatant to a clean 1.5 mL microcentrifugation tube. Add an equal volume of 70% ethanol and vortex.
Transfer 650 μL of the sample to a spin column and centrifuge for 1 min at 10,000× g. Discard the flowthrough and repeat if necessary.
Wash once with 500 μL of Solution WN, centrifuge at 20,000× g for 1 min, and discard the flowthrough.
In a similar manner, wash twice with 500 μL of Wash Solution A. After the final washing step, centrifuge the column for 2 min at 20,000× g.
Transfer the spin column to a fresh elution tube, add 100 μL of Elution Buffer B, and incubate for 1 min at room temperature. Centrifuge for 1 min at 10,000× g. If not all buffer has been eluted, centrifuge again for an additional 1 min at 20,000× g.
Store isolated gDNA at -20 °C. The gDNA isolated in this step will be a combination of both plant and fungal gDNA. Quantification of fungal gDNA is made possible by the use of primers specific for fungal gDNA during quantitative PCR (section F).
Fungal gDNA isolation for calibration
To generate a calibration curve for quantification of fungal gDNA, harvest spores from B. cinerea mycelium that has been growing on PDB plates for 10 days and collect them in a sterile Falcon tube. Harvest the spores by scraping them off with a sterile spatula.
Extract the gDNA using the Plant/Fungi DNA Isolation kit as described above.
Measure the concentration of the genomic DNA using the NanoDrop 2000 spectrophotometer.
Perform a serial dilution (1/2 dilution steps) of the gDNA and measure the concentrations on the NanoDrop 2000 spectrophotometer.
Perform qPCR analysis as described below.
Quantitative PCR for fungal gDNA fungal genomic DNA
For quantification of fungal gDNA, primers for B. cinerea ACTIN or TUBULIN that are specific for fungal gDNA are used. Prepare three technical replicates per sample, including the calibration samples described in section E.
Prepare the reagents for the qPCR in the following proportions:
10 μL of 2× Takyon master mix
2.5 μL of DNA
2 μL of 4 μM forward primer
2 μL of 4 μM reverse primer
3.5 μL of H2O
Final volume: 20 μL
Prepare a master mix with 2× Takyon master mix, primers, and H2O in an autoclaved 1.5 mL tube (all reactions should be performed on ice). Perform each reaction in triplicate.
Load 17.5 μL of the master mix to PCR tubes with flat caps. When loading samples, take note of the arrangement of the samples.
Load 2.5 μL of the genomic DNA to the PCR tubes and carefully seal the flat caps.
Briefly vortex the samples before spinning them down. Ensure that there are no bubbles in the samples as they may interfere with signal detection.
Load samples into the Bio-Rad iQ5 qPCR cycler and wipe the tops of the PCR tubes with a Kimwipe to remove any dust or debris that may interfere with signal detection.
Reaction cycle: 95 °C for 1 min 20 s; (95 °C 20 s, 62 °C 20 s, 72 °C 40 s) × 40; 72 °C 4 min.
Melt curve analysis: 95 °C for 1 min; 60 °C for 1 min; increase to 95 °C in 0.5° increments, hold each temperature for 10 s.
After completion of the qPCR run, export the data to an Excel file to compute the quantifications.
Data analysis
Analysis of drop infections
With the digital calliper, data has already been transferred to an Excel spreadsheet. For calculation of mean, standard deviation, and standard error as well as graphical representation, Excel functions can be used. For statistical analysis beyond Student’s t-test, export the data into software for statistical analysis of your choice. Figure 3 shows exemplary results derived from the drop infection method.
Figure 3. Exemplary results of B. cinerea droplet infection assay. At least 30 leaves were measured per genotype. The respective mean lesion diameter and the standard deviation were calculated in Microsoft Excel and are shown in the bottom two lines. (A) For visualisation and statistical analysis, R-Studio was used in this example. (B) Individual data points are shown as black dots in addition to the mean value (red crossbar) and the standard deviation. Different letters indicate significant differences with p < 0.05 as calculated by ANOVA with post-hoc Tukey-test.
Analysis of spray infections
Calibration of fungal gDNA
Calculate the used volumes of fungal gDNA in nanograms from the concentrations of your different samples from the serial dilution and the used volume per qPCR reaction. Determine the natural logarithm of the gDNA volume and plot the log10 values of gDNA volume vs. respective mean Ct-values. Perform a linear regression analysis on the plot. R2 must be as close as possible to 1.
Determine the parameters m and b of the linear correlation between log-values of gDNA volume and mean Ct-values, for example with the “LINEST” function in Excel.
Note down m and b for subsequent calculation of fungal gDNA in infected plant samples.
Calculation of fungal gDNA in infected plant samples
With the parameters m and b determined above, calculate the volume of gDNA in the infected plant samples.
As only a part of the isolated gDNA was used for the qPCR reaction, multiply the determined mass with a correction factor:
correction factor=(elution volume of gDNA isolation)/(gDNA volume used in qPCR reaction)
Normalise the gDNA volume to the mass of the plant material used for gDNA extraction.
Exemplarily, quantification of fungal gDNA from a spray-infected Arabidopsis leaf sample is shown below.
Sample mass: 190 mg ground Arabidopsis leaf material
Threshold cycle (Ct) value as determined by qPCR: 15.993
Based on the calibration curve obtained, m and b values are determined:
m = -2.179, b = 19.621
Calculate the gDNA in each reaction cycle:
Calculate the correction factor:
Calculate total volume of gDNA:
Normalise amount of fungal gDNA to plant material:
Validation of protocol
This protocol or its parts have been used in the following research articles:
Genenncher et al. (2016). Nucleoporin-Regulated MAP Kinase Signaling in Immunity to a Necrotrophic Fungal Pathogen. Plant Physiol (Figure 1).
Guzha et al. (2022). Cell wall-localized BETA-XYLOSIDASE4 contributes to immunity of Arabidopsis against. Botrytis cinerea (Figure 2C, 2D, 2E, 3C, 3D, and 3E).
General notes and troubleshooting
It is important to carry out independent infection experiments on plants grown at different times to account for variances caused by slight variations in infection or growth conditions. We recommend performing at least three independent experiments with at least 30 infected plant leaves per genotype in case of lesion size measurements, or with at least three biological replicates and at least three technical replicates in case of quantification of fungal gDNA.
In the described protocol, B. cinerea does not contain any resistance or selection markers that would allow growth on selective plates. Work under sterile conditions when preparing B. cinerea spores. When transferring mycelium to fresh PDA plates, make sure that you select regions of the previous PDA plate that contain only B. cinerea mycelium and no contaminations.
As B. cinerea infects plants via air-borne spores, it is important to separate the working area for B. cinerea assays from the area where you work with non-infected plants.
Keep infected plants separate from normal plants in a dedicated growth chamber.
Avoid all normal plant areas after you prepared B. cinerea spores or carried out an infection assay.
After infection assays, clean all tools used for infection by incubation with anti-fungal disinfectant and 70% ethanol. Similarly, clean the working area for infection after use.
During the configuration of your digital calliper, edit the parameters of the measuring instrument so that data will be transferred to Excel. Individual measurements will then be transferred into successive rows of an Excel sheet. Adjust the end row number so that all measurements from a single experiment will fit into the same Excel sheet.
Lesion size is usually not perfectly circular (Figure 2B); so, measuring the diameter becomes more difficult. Measure lesion size diameter for all leaves in a similar angle to the middle vein (e.g., always parallel to the middle vein); all leaves should be measured by the same experimenter. For generation of the standard curve for gDNA quantification, it is important to accurately measure the gDNA concentrations on a NanoDrop after the serial dilutions.
Troubleshooting
Poor fungal growth after infection: maintain the plants under high humidity by placing a dome over the plants after infection.
Acknowledgments
This protocol was derived from Guzha et al. (2022). We are grateful to Prof. Dr. Marcel Wiermer and Dr. Denise Hartken for useful discussions on B. cinerea infection and Dr. Sven Haroth for qPCR advice. This work was supported by German Research Foundation (DFG, IS 273/10-1, IRTG 2172 PRoTECT to T.I), the Studienstiftung des Deutschen Volkes (stipend to P.S.) and a grant from the U.S. Department of Energy, Office of Science, BER program DE-SC0020325 to K.D.C.
Competing interests
The authors have no financial or non-financial competing interests.
References
Bi, K., Liang, Y., Mengiste, T. and Sharon, A. (2022). Killing softly: a roadmap of Botrytis cinerea pathogenicity. Trends Plant Sci: 28(2):211-222.
Caseys, C., Shi, G., Soltis, N., Gwinner, R., Corwin, J., Atwell, S. and Kliebenstein, D. J. (2021). Quantitative interactions: the disease outcome of Botrytis cinerea across the plant kingdom. G3 (Bethesda) 11(8): jkab175.
Dean, R., Van Kan, J. A., Pretorius, Z. A., Hammond-Kosack, K. E., Di Pietro, A., Spanu, P. D., Rudd, J. J., Dickman, M., Kahmann, R., Ellis, J., et al. (2012). The Top 10 fungal pathogens in molecular plant pathology. Mol Plant Pathol 13(4): 414-430.
Elad, Y., Pertot, I., Cotes Prado, A. M. and Stewart, A. (2016). Plant Hosts of Botrytis spp. In: Fillinger, S. and Elad, Y. (Eds.). Botrytis – the Fungus, the Pathogen and its Management in Agricultural Systems (pp. 413-486). Springer International Publishing: Cham.
Ettenauer, J., Piñar, G., Tafer, H., and Sterflinger, K. (2014). Quantification of fungal abundance on cultural heritage using real time PCR targeting the β-actin gene.Front Microbiol 5: 262
Genenncher, B., Wirthmueller, L., Roth, C., Klenke, M., Ma, L., Sharon, A. and Wiermer, M. (2016). Nucleoporin-Regulated MAP Kinase Signaling in Immunity to a Necrotrophic Fungal Pathogen.Plant Physiol 172(2): 1293-1305.
Guzha, A. McGee, R., Scholz, P., Hartken, D., Lüdke, D., Bauer, K., Wenig, M., Zienkiewicz, K., Herrfurth, C., Feussner, I. et al.(2022). Cell wall-localized BETA-XYLOSIDASE4 contributes to immunity of Arabidopsis against Botrytis cinerea. Plant Physiol 189(3): 1794-1813.
IPPC Secretariat. (2021). Scientific review of the impact of climate change on plant pests – A global challenge to prevent and mitigate plant pest risks in agriculture, forestry and ecosystems. Rome. FAO on behalf of the IPPC Secretariat.
Jacob, C., Panchal, S. and Melotto, M., (2017). Surface inoculation and quantification of Pseudomonas syringae population in the Arabidopsis leaf apoplast. Bio Protoc 7(5): e2167.
Kliebenstein, D. J., Rowe, H. C. and Denby, K. J. (2005). Secondary metabolites influence Arabidopsis/Botrytis interactions: variation in host production and pathogen sensitivity.Plant J 44(1): 25-36.
Rowe, H. C. and Kliebenstein, D. J. (2008). Complex genetics control natural variation in Arabidopsis thaliana resistance to Botrytis cinerea. Genetics180(4): 2237-2250.
Savary, S., Willocquet, L., Pethybridge, S. J., Esker, P., McRoberts, N. and Nelson, A. (2019). The global burden of pathogens and pests on major food crops. Nat Ecol Evol 3(3): 430-439.
Soltis, N. E., Caseys, C., Zhang, W., Corwin, J. A., Atwell, S. and Kliebenstein, D. J. (2020). Pathogen Genetic Control of Transcriptome Variation in the Arabidopsis thaliana - Botrytis cinerea Pathosystem.Genetics 215(1): 253-266.
Vasquez-Montaño, E., Hoppe, G., Vega, A., Olivares-Yañez, C. and Canessa, P. (2020). Defects in the ferroxidase that participates in the reductive iron assimilation system results in hypervirulence in Botrytis cinerea. mBio 11(4): e01379-20.
Windram, O., Madhou, P., McHattie, S., Hill, C., Hickman, R., Cooke, E., Jenkins, D. J., Penfold, C. A., Baxter, L., Breeze, E. et al. (2012). Arabidopsis Defense against Botrytis cinerea: Chronology and Regulation Deciphered by High-Resolution Temporal Transcriptomic Analysis.Plant Cell 24(9): 3530-3557.
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Plant Science > Plant physiology > Biotic stress
Plant Science > Plant molecular biology > DNA
Biological Sciences > Biological techniques > Microbiology techniques
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Perforated Patch Clamp Recordings in ex vivo Brain Slices from Adult Mice
SH Simon Hess
HW Helmut Wratil
PK Peter Kloppenburg
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4741 Views: 1036
Reviewed by: Miao HeXiaoliang Zhao Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Jul 2022
Abstract
Intracellular signaling pathways directly and indirectly regulate neuronal activity. In cellular electrophysiological measurements with sharp electrodes or whole-cell patch clamp recordings, there is a great risk that these signaling pathways are disturbed, significantly altering the electrophysiological properties of the measured neurons. Perforated-patch clamp recordings circumvent this issue, allowing long-term electrophysiological recordings with minimized impairment of the intracellular milieu. Based on previous studies, we describe a superstition-free protocol that can be used to routinely perform perforated patch clamp recordings for current and voltage measurements.
Keywords: Perforated patch clamp Brain slices Current clamp Voltage clamp Electrophysiology
Background
Neuronal activity is controlled to a large extent by intracellular signaling systems. This study describes how to perform perforated-patch clamp recordings for high-quality single-electrode whole-cell current and voltage clamp recordings with minimized impact on the cytosolic integrity.
Whole-cell patch clamp recordings have largely replaced intracellular recordings with sharp microelectrodes for single-electrode current and voltage clamp experiments. Because of the high seal resistance between the cell membrane and the recording electrode, this patch clamp configuration allows recording with a high signal-to-noise ratio, even from very small neurons. In addition, the whole-cell configuration provides low access resistance, which helps to ensure that the patch electrode solution can exchange freely with the cytoplasm. The whole-cell configuration is therefore also ideally suited to control the composition of the intracellular milieu, such as ion concentration, and to load the recorded neurons with tracers, sensors, and pharmacological agents. However, the free exchange of molecules between the cytoplasm and the patch electrode has a downside, as it impairs neuronal function by interfering with the cytosolic signaling system. Practically, this makes it impossible to use the whole-cell recording configuration for long-term measurements without significantly altering the physiological state of the recorded neurons.
The perforated-patch configuration, introduced initially by Lindau and Fernandez (1986) and Horn and Marty (1988), minimizes or even overcomes this drawback of the whole-cell configuration. Instead of rupturing the membrane under the recording electrode, which mediates the exchange between the cytosol and the electrode solution, pore-forming substances (ionophores) provide electrical access to the cell interior while largely maintaining the integrity of the cytoplasmic components of the neuron.
The original and most used perforating agents have been the antibiotic polyenes nystatin, amphotericin B, and the antibiotic polypeptide gramicidin (Horn and Marty, 1988; Akaike, 1994; Akaike and Harata, 1994; Kyrozis and Reichling, 1995; Klöckener et al., 2011; Könner et al., 2011; Hess et al., 2013). While polyenes and the peptide exhibit differences in their pore-forming mechanisms and ion selectivity (Myers and Haydon, 1972; de Kruijff and Demel, 1974; Russell et al., 1977; Tajima et al., 1996), their pores share key common properties: they are permeable to small molecules with a molecular weight up to ~200 Da, including monovalent ions (Urry, 1971; de Kruijff et al., 1974; Kyrozis and Reichling, 1995). However, they are neither permeable to divalent ions like Ca2+ nor intracellular signaling molecules of larger molecular weight. Building on previous studies, we describe here a protocol that can be used to routinely perform perforated-patch clamp recordings in brain slices of adult mice. The procedure is based on the use of amphotericin B as ionophore, as it has been, in our hands, the most suitable to achieve low access resistance and reproducibility. We describe how this approach can be used for current and voltage measurements and how this can be combined with single-cell labeling. We have applied this method to a variety of neuron types (see Table 1 for examples), but, for consistency, we only show data from substantia nigra pars compacta (SNpc) dopaminergic (DA) neurons here. In the context of DA neuron recording, one reviewer has strongly suggested mentioning the work of Cattaneo et al. (2021) describing cell-attached and whole-cell patch clamp recordings from DA neurons in the SNpc.
Materials and reagents
Animal preparation
Dopaminergic neurons of the substantia nigra pars compacta (SNpc) from brain slices of adult (12–14 weeks old) mice
Isoflurane (AbbVie Deutschland GmbH and Co KG, catalog number: B506); storage: room temperature (RT)
Pattex Ultra Gel Matic (Henkel AG, EAN: 4015000444972)
Feather® double edge blades (Plano, Feather, catalog number: 121-9)
Glycerol-based modified artificial cerebrospinal fluid (GACSF) (Ye et al., 2006) (see Recipes)
Artificial cerebrospinal fluid (ACSF) for current clamp recordings (see Recipes)
Artificial cerebrospinal fluid (ACSF) for Ca2+ current recordings in voltage clamp (see Recipes)
Artificial cerebrospinal fluid and electrode solutions
Calcium chloride 2-hydrate (CaCl2·2H2O), powder for analysis, ACS (AppliChem, catalog number: 131232); storage: RT
Potassium chloride (KCl) (Reag. USP) for analysis, ACS, ISO (AppliChem, catalog number: 131494); storage: RT
Magnesium chloride 6-hydrate (MgCl2·6H2O) (BP, Ph. Eur.) pure, pharma grade (AppliChem, catalog number: 141396); storage: RT
Sodium hydrogen carbonate (NaHCO3) (Reag. USP) for analysis, ACS, ISO (AppliChem, catalog number: 131638); storage: RT
Sodium chloride (NaCl) for analysis, ACS, ISO (AppliChem, catalog number: 131659); storage: RT
Sodium di-hydrogen phosphate 1-hydrate (NaH2PO4·H2O) (Reag. USP, Ph. Eur.) for analysis, ACS (AppliChem, catalog number: 131965); storage: RT
Sodium hypochlorite 14% Cl2 (ClNaO) in aqueous solution (VWR, catalog number: 27900); storage: RT
Sodium hydroxide (NaOH) 1 mol/L (1 N) in aqueous solution (VWR, catalog number: 31627); storage: RT
D(+)-glucose anhydrous BioChemika (AppliChem, catalog number: A1422); storage: RT
HEPES for molecular biology (AppliChem, catalog number: A3724); storage: RT
Glycerol bioreagent, suitable for (insect) cell culture, suitable for electrophoresis, ≥ 99% (GC) (Sigma-Aldrich, catalog number: G2025); storage: RT
C6H11KO7, potassium-D-gluconate (Sigma-Aldrich, catalog number: G4500); storage: RT
CsCl, cesium chloride (Sigma-Aldrich, catalog number: C4036); storage: RT
CsOH, cesium hydroxide 99.9% (Sigma-Aldrich, catalog number: 232041); storage: RT
EGTA, ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid (Sigma-Aldrich, catalog number: 03777-10G)
TEA, Tetraethylammonium chloride (Sigma-Aldrich, catalog number: T2265); storage: RT
Picrotoxin ≥ 97.5% (Roth, catalog number: 7093.1); storage: RT
CNQX, 6-Cyano-7-nitrochinoxalin-2,3-dion ≥ 98% (HPLC), solid (Sigma-Aldrich, catalog number: C127); storage: RT
DAP-5, DL-2-amino-5-phosphopentanoic acid solid (Sigma-Aldrich, catalog number: A5282); storage: RT
TTX, tetrodotoxin citrate (Biotrend, catalog number: ARCD-0640-1); storage: 4 °C
Tetramethylrhodamine-dextran, 3000 MW, anionic, lysine fixable (Thermo Fisher Scientific, Invitrogen, catalog number: D3308); storage: -20 °C
Biocytin (Sigma-Aldrich, catalog number: B4261); storage: -20 °C
Electrode solution (current clamp) (see Recipes)
Electrode solution (voltage clamp: Ca2+ currents) (see Recipes)
Ionophore preparation and patch clamp recordings
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: D8418); storage: RT
Amphotericin B from Streptomyces sp. ~80% (HPLC), powder (Sigma-Aldrich, catalog number: A4888); storage: 4 °C
Microtubes, PP, 1.5 mL, with attached cap, BIO-CERT® PCR QUALITY (Brand, catalog number: 780420)
Silver wire for electrodes (Science Products, catalog number: AG-10W)
Amphotericin B stock solution (see Recipes)
Equipment
Pipette puller (Narishige, model: PC-10)
Electrode borosilicate glass (Science Products, catalog number: GB150-8P)
Micromanipulator (Luigs-Neumann, model: mini 23)
Data acquisition unit [Cambridge Electronic Design Ltd. (CED), model: Micro 1401 MkII]
Patch clamp amplifier (HEKA, model: EPC-10 USB)
Temperature controller (Warner Instrument Corp., model: TC-324B)
Inline heater (Warner Instrument Corp., model: SH-27B)
Fixed stage upright microscope (Olympus, model: BX51WI) equipped with a 20× water-immersion objective (XLUMPLFL, 0.95 numerical aperture, 2 mm working distance, Olympus), a 4× magnification changer (U-TVAC, Olympus), infrared differential interference contrast optics (Dodt and Zieglgänsberger, 1990), and fluorescence optics
Vibratome (Leica, model: VT1200S)
Heating circulator (Julabo, model: ED)
Anesthesia machine (Groppler Medizintechnik, Univet, model: Porta)
Magnetic stirrer (VWR, catalog number: 12365-428)
Vortex mixer (Scientific Industries, Vortex Genie 2, model: SI-0256)
Ultrasonic bath (Bandelin Electronic, model: DT 31 H)
Centrifuge (Eppendorf AG, model: 5415 D)
Peristaltic pump (Ismatec, model: ISM597D) with Tygon tube (1.6 and 2.4 inner diameters, Ismatec, Tygon, catalog number: T3350)
Fine scissors, sharp (FST, catalog number: 14060-09)
Surgical scissors, blunt (FST, catalog number: 14001-14)
Double spatulas, micro (VWR, catalog number: 231-2261)
Slice anchor (Warner Instruments, model: SHD-26H/15)
Software
PatchMaster (HEKA, Ver. 2.32, www.heka.com)
Spike2 (Cambridge Electronic Design Ltd. (CED), www.ced.co.uk)
Procedure
Animals and preparation of mature/adult brain slices
Lightly anesthetize the animal with isoflurane (5%).
Decapitate the mouse with surgical scissors.
Insert the fine sharp scissors into the spinal cord canal and make an incision by cutting along the midline of the skull from caudal to rostral to approximately the frontonasal suture.
To open the skull, grab one side of the skull with the groove of the tissue forceps and rotate the forceps 180 degrees. Repeat the procedure with the other side of the skull.
Remove the brain with a spatula: insert the spatula between the olfactory bulb and the rest of the brain and pull the mouse brain out.
Note: Be as quick as possible (≤ 30 s), especially when dissecting older animals!
Make two trim cuts (caudal, rostral) with a standard razor blade.
Glue the brain onto the vibratome plate using Pattex Ultra Gel Matic.
Cut the brain with the vibratome under 4 °C cold carbogenated (Carbogen, 95% O2, 5% CO2) GACSF (see Recipes) using a Feather® razor blade.
Vibratome settings: speed 0.06 mm/s; amplitude 1.05 mm; thickness 290 μm.
Incubate the brain slices for 25–50 min in a recovery bath (carbogenated ACSF, see Recipes) at 37 °C.
Keep the brain slices in carbogenated ACSF at RT. The slices should be used within 5 h.
Place the brain slices in the recording chamber for the recordings held down with a slice anchor and superfused with carbogenated ACSF at the desired experimental temperature.
Preparation and usage of the ionophore (amphotericin B)
Preparation
Make a stock solution of amphotericin B by dissolving 4 mg of amphotericin B in 100 μL of DMSO.
Sonicate the stock solution until the solution becomes a uniformly turbid yellowish solution.
Vortex the stock solution several times.
Prepare two 1.5 mL microtubes, each filled with 1 mL of electrode solution (for current or voltage clamp). A defined volume of amphotericin B stock solution (see Table 1) is added to 1 mL of electrode solution, while the solution in the other microtube remains free of amphotericin (tip fill). In our experience, a final amphotericin concentration of 160–200 μg/mL (4–5 μL stock solution) ensures excellent perforation and recording conditions for many neuron types. However, depending on cell type and recording mode (current clamp or voltage clamp), optimizing the amphotericin concentration might be useful or even necessary (for examples, see Table 1).
Table 1. Recommended amphotericin B concentrations for different neuron types
Recording mode Neuron type Amount of stock solution (μL) Final amphotericin B concentration (μg/mL)
Current clamp Dopaminergic (DA) neuron (midbrain) (Hess et al., 2013) 4.5 180
Serotonergic neuron (dorsal raphe nucleus) (Xiao et al., 2021) 4 160
POMC neuron (hypothalamus) (Paeger et al., 2017) 4 160
AgRP neuron (hypothalamus) (Steculorum et al., 2016) 4 160
V2a neurons (spinal cord) (Reinoss et al., 2020) 20 800
Voltage clamp Dopaminergic (DA) neuron (midbrain) (Siller et al., 2022) 10 400
Shake (by hand) the amphotericin B–containing electrode solution several times.
Add 1% (0.1 mg/100 μL) tetramethylrhodamine-dextran to the amphotericin B–containing electrode solution. Tetramethylrhodamine-dextran is used to check the integrity of the cell membrane once the ionophore has perforated the membrane (for further explanation, see below.)
Put all three solutions (stock, tip fill, and amphotericin B–containing electrode solution) on ice. The amphotericin B–containing electrode solution can be used for up to three hours. After that, prepare a fresh solution.
Setting up the perforated-patch clamp recording for current clamp
Pull electrodes with resistances between 3 and 5 MΩ.
Fill the electrode tips with plain electrode solution (tip fill without amphotericin B).
For the backfill, use the amphotericin B–containing electrode solution.
Put the electrode in the bath but do not apply positive pressure!
Note: Commonly, in patch clamp recordings from brain slices, positive pressure is applied when the electrode is advanced to the cell to prevent clogging of the electrode tip and to clean the target cell. However, in this case, the amphotericin B–containing solution would be driven into the electrode tip, severely compromising seal formation.
Only when the electrode is positioned directly in front of the cell, apply a slight positive pressure with the mouthpiece and continue to approach the cell. As soon as a dent forms in the cell membrane in front of the electrode, immediately release the positive pressure (Figure 1C).
Apply gentle negative pressure with the mouthpiece and constantly monitor the test pulse (voltage clamp mode, holding potential: 0 mV, test pulse amplitude: 5 mV) and the seal resistance.
Release the pressure after you have reached a seal resistance of > 600 MΩ.
Note: Sometimes, you will not be able to reach a giga seal since the ionophore already starts perforating the cell membrane during the sealing process, which can take up to 20 min.
Switch to current clamp and monitor the perforation process. Ideally, you should see a continuous transition from the on cell to the perforated configuration, as shown in Figure 1A.
Figure 1. Perforation process. Original recordings showing the transition from the on-cell to the perforated-patch recording configuration under current clamp (A) and voltage clamp (B). Bottom: segments of the original traces shown in the top panel in higher time resolution. The numbers indicate the times from which the segments originate. (B). Voltage pulses (5 mV, 5 ms, HP = -60 mV) were applied every 10 s. Bottom left: test pulse before perforation (RS > 100 MΩ), Bottom right: test pulse after perforation has reached a steady state (RS < 20 MΩ). HP, holding potential. (C). Seal formation. Top: slight positive pressure forms a dent in the cell membrane in front of the electrode. Bottom: after releasing the positive pressure, negative pressure supports seal formation. Scale bar: 10 μm.
The perforation process is reflected by an increase in action potential amplitude, indicating a decrease in series resistance. During this process, spontaneous conversions to the whole-cell configuration can occur, which become evident by an abrupt increase in action potential amplitude (Figure 2A). In many cases, this conversion is accompanied by hyperpolarization and the cessation of spiking activity (e.g., due to the opening of KATP channels).If no further changes in the parameters mentioned above occur, the perforation process is complete, and the actual measurements can begin.
Note: To get meaningful measurements of electrophysiological properties of the investigated cell, the series resistance should ideally be well below 60 MΩ.
Figure 2. Tetramethylrhodamine-dextran as a marker for rupture of the membrane. Spontaneous conversion from the perforated patch to the whole-cell configuration. (A). Current clamp recording with corresponding frequency plot showing the spontaneous rupture of the membrane patch during the perforation process. The arrow marks the jump in spike amplitude due to the rupture of the membrane. (B). Tetramethylrhodamine-dextran fluorescence is confined to the tip of the electrode, indicating the integrity of the membrane patch (top). After rupturing the membrane, tetramethylrhodamine-dextran has diffused into the neuron (bottom). Scale bar: 20 μm.
During the recording, check the tetramethylrhodamine-dextran fluorescence regularly to see if the membrane patch in the electrode is still intact (Figure 2B). If not, terminate the recording.
Adding tetramethylrhodamine-dextran helps significantly to monitor the integrity of the perforated patch. This is very useful since spontaneous ruptures of the membrane patch are often not immediately reflected in noticeable changes in the action potential amplitude and the series resistance once the membrane has been perforated. Due to its molecular weight of 3,000 Da, tetramethylrhodamine-dextran can neither permeate through the cell membrane nor the pores formed by the ionophores, making it possible to detect whether the membrane patch has been ruptured.
Note: In our experience, tetramethylrhodamine-dextran does not compromise the quality of the recording. Although other fluorescent molecules may be suitable, checking whether they interfere with the recording is crucial.
Once the perforated configuration is established, long-term recordings can be performed with a significantly minimized rundown compared to the whole-cell configuration. Figures 3–5 show examples of perforated-patch clamp recordings in different experimental settings, illustrating the possibilities of this recording configuration, e.g., in long-lasting current and voltage clamp experiments. A direct comparison between the whole-cell and perforated configurations is shown in Figure 3. In the whole-cell configuration, action potential frequency decreased dramatically within the first 10 min (Figure 3A). In contrast, the frequency remained stable in the perforated-patch clamp recordings (Figure 3B). This configuration is, therefore, particularly suitable for long-term (> 2 h) pharmacological experiments where it is desirable to demonstrate reversibility and reproducibility in the same recording. Such an experiment is shown in Figure 4, where cocaine was bath-applied and washed out twice.
Figure 3. Time course of spontaneous action potential frequency from 30 min measurements in whole-cell (A) and perforated-patch recordings (B). Whole-cell recordings (n = 9); perforated-patch recordings (n = 11). The traces at the bottom correspond to the red symbols in the respective frequency plots.
Figure 4. Long-term perforated patch clamp recording. Example of a neuropharmacological experiment (~2 h) using the perforated patch configuration. Top: action potential frequency (Hz) of a mouse dopaminergic (DA) substantia nigra pars compacta (SNpc) neurons during two exposures to 10 μM cocaine (COC). Blue dots indicate the instantaneous frequency, and the red line indicates the average frequency (bin size: 10 s). Middle: original recording. The numbers correspond to the sections displayed at the bottom. Current pulses were applied at the sections where gaps in the recordings occurred.
Setting up the perforated patch clamp recording for voltage clamp
To perform voltage clamp experiments, follow steps 1–7 of section C and then continue with the following steps.
Note: Higher amphotericin B concentrations might be useful (see Table 1).
Switch the holding potential to -60 mV.
Continue applying 5 mV test pulses in voltage-clamp mode (Figure 1B).
Wait until the perforation has reached a steady state (stable current amplitude).
Check the integrity of the membrane patch occasionally, as already described.
Examples of successful voltage clamp recordings are given in Figure 5.
Figure 5. Perforated patch voltage clamp recordings of voltage-activated Ca2+ currents (ICa). ICa was induced by depolarizing voltage steps to 0 mV (A) or -10 mV (B) from a holding potential of -60 mV every 10 s. A, B. Example of peak ICa in a mouse dopaminergic (DA) substantia nigra pars compacta (SNpc) neuron plotted over time during cadmium (A; 1 mM) or nifedipine (B; 10 μM) bath application. The numbers correspond to calcium current traces shown at the bottom.
Complementary application: single-cell labeling
Use an electrode solution containing a tracer (e.g., biocytin).
Perform the perforated-patch clamp experiment as described.
After completion of the electrophysiological experiment, rupture the membrane patch either by gentle suction or by a large hyperpolarizing voltage pulse (~1–1.5 V) to allow cell labeling by diffusion of the tracer into the cell.
Further processing of the brain slice containing the labeled neuron can be performed by using standard histological and immunohistochemical procedures.
An example of successful labeling of a recorded neuron is given in Figure 6.
Figure 6. Morphological and immunohistochemical characterization. Post-recording immunohistochemical identification of a dopaminergic (DA) neuron filled with biocytin after the electrophysiological experiment (current clamp). Top: single-cell staining of the DA neuron with biocytin-streptavidin. Scale bar: 50 μm. Inset: immunohistochemical co-labeling against tyrosine-hydroxylase (TH), a marker for DA neurons. Scale bar: 20 μm. Bottom: perforated-patch clamp recording (current clamp) of the labeled neuron showing pacemaker activity and a prominent ‘sag’ potential during hyperpolarization, both characteristic features of DA substantia nigra pars compacta (SNpc) neurons.
Data analysis
Data were recorded with the PatchMaster software (HEKA). Data were sampled at 10 kHz and low-pass filtered at 2 kHz with a four-pole Bessel filter. Data were analyzed with Spike2 and custom-made scripts written in Python 3.x.
Recipes
In the following recipes, the measured osmolarity is given. When composing solutions, we recommend that the osmolarity of the electrode solution (intracellular solution) be 10–20 mOsmol·L-1 lower than the extracellular solution.
Animal preparation
Glycerol-based modified artificial cerebrospinal fluid (GACSF)
225 mM glycerol
2.5 mM KCl
2 mM MgCl2·6H2O
2 mM CaCl2·2H2O
1.2 mM NaH2PO4
21 mM NaHCO3
10 mM HEPES
5 mM glucose
Gassed with carbogen and adjusted to pH = 7.2 with NaOH, resulting in an osmolarity of 300–310 mOsmol/L.
Artificial cerebrospinal fluid (ACSF) for current clamp recordings
125 mM NaCl
2.5 mM KCl
2 mM MgCl2·6H2O
2 mM CaCl2·2H2O
1.2 mM NaH2PO4
21 mM NaHCO3
10 mM HEPES
5 mM glucose
Gassed with carbogen and adjusted to pH = 7.2 with NaOH, resulting in an osmolarity of ~310 mOsmol/L.
Note: Block of glutamatergic and GABAergic synaptic input can be achieved by adding 0.1 mM picrotoxin, 0.05 mM DAP-5, and 0.01 mM CNQX to the ACSF.
Artificial cerebrospinal fluid (ACSF) for Ca2+ current recordings in voltage clamp
66.5 mM NaCl
20 mM CsCl
40 mM TEA-Cl
4 mM MgCl2·6H2O
3 mM CaCl2·2H2O
21 mM NaHCO3
10 mM HEPES
5 mM Glucose
0.001 mM TTX
0.1 mM picrotoxin
0.05 mM DAP-5
0.01 mM CNQX
Gassed with carbogen and adjusted to pH = 7.2 with HCl, resulting in an osmolarity of ~305 mOsmol/L.
Patch clamp recordings
Electrode solution (current clamp)
141 mM K-gluconate
10 mM KCl
10 mM HEPES
0.1 mM EGTA
2 mM MgCl2·6H2O
Adjusted to pH = 7.2 with KOH, resulting in an osmolarity of ~290 mOsmol/L.
Electrode solution (voltage clamp: Ca2+ currents)
146 mM CsCl
10 mM HEPES
0.1 mM EGTA
2 mM MgCl2·6H2O
Adjusted to pH=7.2 with CsOH, resulting in an osmolarity of ~290 mOsmol/L.
Ionophore (amphotericin B) solutions
Stock solution
100 μL of DMSO
4 mg of amphotericin B
Acknowledgments
Work in the lab of PK is supported by grants KL 762/7-1 (401832153) and CRC 1451 TP A01 (431549029) and of the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) and the DFG’s Germany Excellence Strategy—EXC 2030-390661388. We thank Debora Fusca for providing Figure 1C.
This protocol was derived from the original work of Siller et al. (2022).
Competing interests
The authors report no declarations of interest.
Ethics
All animal procedures were conducted in compliance with guidelines approved by local government authorities (§4.17.020; §4.22.028; LANUV NRW, Recklinghausen, Germany).
References
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Cattaneo, S., Regoni, M., Sassone, J. and Taverna, S. (2021). Cell-attached and Whole-cell Patch-clamp Recordings of Dopamine Neurons in the Substantia Nigra Pars Compacta of Mouse Brain Slices. Bio Protoc 11(15): e4109.
de Kruijff, B. and Demel, R. A. (1974). Polyene antibiotic-sterol interactions in membranes of Acholeplasma laidlawii cells and lecithin liposomes. III. Molecular structure of the polyene antibiotic-cholesterol complexes. Biochim Biophys Acta 339(1): 57-70.
de Kruijff, B., Gerritsen, W. J., Oerlemans, A., Demel, R. A. and van Deenen, L. L. (1974). Polyene antibiotic-sterol interactions in membranes of Acholeplasma laidlawii cells and lecithin liposomes. I. Specificity of the membrane permeability changes induced by the polyene antibiotics. Biochim Biophys Acta 339(1): 30-43.
Dodt, H. U. and Zieglgänsberger, W. (1990). Visualizing unstained neurons in living brain slices by infrared DIC-videomicroscopy. Brain Res 537(1-2): 333-336.
Hess, M. E., Hess, S., Meyer, K. D., Verhagen, L. A., Koch, L., Bronneke, H. S., Dietrich, M. O., Jordan, S. D., Saletore, Y., Elemento, O., et al. (2013). The fat mass and obesity associated gene (Fto) regulates activity of the dopaminergic midbrain circuitry. Nat Neurosci 16(8): 1042-1048.
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Könner, A. C., Hess, S., Tovar, S., Mesaros, A., Sánchez-Lasheras, C., Evers, N., Verhagen, L. A., Brönneke, H. S., Kleinridders, A., Hampel, B., et al. (2011). Role for insulin signaling in catecholaminergic neurons in control of energy homeostasis. Cell Metab 13(6): 720-728.
Kyrozis, A. and Reichling, D. B. (1995). Perforated-patch recording with gramicidin avoids artifactual changes in intracellular chloride concentration. J Neurosci Methods 57(1): 27-35.
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Paeger, L., Pippow, A., Hess, S., Paehler, M., Klein, A. C., Husch, A., Pouzat, C., Brüning, J. C. and Kloppenburg, P. (2017). Energy imbalance alters Ca2+ handling and excitability of POMC neurons. eLife 6: e25641.
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Mass Spectrometry-based Lipidomics, Lipid Bioenergetics, and Web Tool for Lipid Profiling and Quantification in Human Cells
LC Liang Cui
MY Meisam Yousefi
XY Xin Yap
CK Clara W.T. Koh
KT Kwan Sing Leona Tay
YO Yaw Shin Ooi
KC Kuan Rong Chan
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4742 Views: 1335
Reviewed by: Alka MehraNeelanjan BoseJianhong ChingPrajita Pandey
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Original Research Article:
The authors used this protocol in PLOS Pathogens Aug 2022
Abstract
Lipids can play diverse roles in metabolism, signaling, transport across membranes, regulating body temperature, and inflammation. Some viruses have evolved to exploit lipids in human cells to promote viral entry, fusion, replication, assembly, and energy production through fatty acid beta-oxidation. Hence, studying the virus–lipid interactions provides an opportunity to understand the biological processes involved in the viral life cycle, which can facilitate the development of antivirals. Due to the diversity and complexity of lipids, the assessment of lipid utilization in infected host cells can be challenging. However, the development of mass spectrometry, bioenergetics profiling, and bioinformatics has significantly advanced our knowledge on the study of lipidomics. Herein, we describe the detailed methods for lipid extraction, mass spectrometry, and assessment of fatty acid oxidation on cellular bioenergetics, as well as the bioinformatics approaches for detailed lipid analysis and utilization in host cells. These methods were employed for the investigation of lipid alterations in TMEM41B- and VMP1-deficient cells, where we previously found global dysregulations of the lipidome in these cells. Furthermore, we developed a web app to plot clustermaps or heatmaps for mass spectrometry data that is open source and can be hosted locally or at https://kuanrongchan-lipid-metabolite-analysis-app-k4im47.streamlit.app/. This protocol provides an efficient step-by-step methodology to assess lipid composition and usage in host cells.
Graphical overview
Keywords: Mass spectrometry Lipid profiling Lipidomics Seahorse assay Bioenergetics Virus–lipid interactions Web tool Clustergram
Background
Lipids are critical for metabolism, signaling, transport across membranes, regulating body temperature, and inflammation. The dysregulation of lipid metabolism or lipid transport can thus disrupt the cell homeostasis and promote oxidative stress that leads to excessive inflammation and cell death. Interestingly, some viruses such as the enveloped viruses have evolved to hijack host cell lipids to augment viral fusion, formation of replication complexes, assembly, and egress. In addition, viruses can also modulate cellular lipid metabolic pathways and leverage intracellular lipid stores to promote viral replication. For instance, dengue virus (DENV) infection can upregulate and re-localize fatty acid synthase (FASN) to the replication complex to increase beta-oxidation for energy production (Heaton et al., 2010; Tang et al., 2014). DENV can also target lipid droplet stores to utilize triglycerides for the production of fatty acids, which are subsequently transported to the mitochondria where they undergo beta-oxidation to synthesize ATP required for viral replication (Zhang et al., 2017). However, much work remains to be done to better understand the virus and lipid interactions responsible for productive virus infection.
RNA interference and CRISPR technologies are useful tools to probe into the lipid metabolic processes required for viral infection. For example, we have recently identified that TMEM41B and VMP1 play a central role in lipid mobilization, mitochondrial beta-oxidation, and global metabolic regulations, to facilitate the replication of flaviviruses and coronaviruses (Yousefi et al., 2022). Deficiency of these proteins resulted in impaired beta-oxidation capacity and dysregulation of the cell lipidome that severely compromised viral infection. To gain insights into the lipid metabolism pathways that are differentially modulated before and after virus infection, we can leverage mass spectrometry to characterize the lipidome. The current protocol describes the detailed procedures required for mass spectrometry in target cells. We have used liquid chromatography–mass spectrometry (LC–MS), the preferred technique for use in metabolomics and lipidomics due to its ability to identify metabolites, even at low concentrations. The method also allows for untargeted lipidomics analysis, so users can simultaneously quantify a variety of lipids in host cells. Furthermore, to evaluate the effects of fatty acid oxidation on cellular bioenergetics, a modified Seahorse assay can be used to ascertain the lipid dependencies for energy production. In combination, these assays provide molecular insights into the role of lipid metabolism in cellular energetics and their potential contribution to virus infection outcome. Finally, we developed a web tool (https://kuanrongchan-lipid-metabolite-analysis-app-k4im47.streamlit.app/) that allows users to plot clustergrams and heatmaps based on the intensity measurements from the LC–MS. While we have used these methods to study virus–lipid interactions in human cells, as lipids are involved in multiple cell processes, we believe that these methods can be more broadly applied to study the role of lipids for other diseases, such as diabetes, obesity, atherosclerosis, and cancer.
In summary, we will describe the detailed protocols for:
Sample preparation and cell extraction for LC–MS
LC–MS analysis and interpretation
Bioinformatics analysis and web tool for data analysis
Seahorse bioenergetics analysis for fatty acid oxidation
Materials and reagents
HEK 293FT cells (Invitrogen, catalog number: R70007)
Dulbecco’s modified Eagle medium (DMEM) with L-glutamine and sodium pyruvate (Thermo Fisher Scientific, Gibco, catalog number: 11995)
0.22 μm filter (Merck Millipore, catalog number: SLGP0033RS)
Seahorse XF RPMI medium (without phenol red, bicarbonate, glucose, pyruvate, or glutamine, contains 1 mM HEPES, adjusted to pH 7.4) (Agilent, catalog number: 103576-100)
Seahorse XF calibrant solution [Agilent, 100 mL (catalog number: 03059-000) or 500 mL (catalog number: l100840-000)]
L-glutamine (200 mM) (Thermo Fisher Scientific, Gibco, catalog number: 25030081)
Fetal bovine serum (FBS) (HyClone, catalog number: SH30396.03)
Trypsin-EDTA (0.25%), phenol red (Thermo Fisher Scientific, Gibco, catalog number: 25200072)
10× PBS (Sigma-Aldrich, catalog number: BUF-2040-10X4L)
Additional materials required for mass spectrometry
Cell scraper (Thermo Fisher Scientific, catalog number: 08-100-241)
Tert-butyl methyl ether (MTBE), HPLC Plus (Sigma-Aldrich, catalog number: 650560-1L)
Methanol, hypergrade for LC-MS LiChrosolv® (Merck, Supelco, catalog number: 1.06035.2500)
Acetonitrile (ACN), hypergrade for LC-MS LiChrosolv® (Merck, Supelco, catalog number: 1.00029.2500)
Isopropanol, hypergrade for LC-MS LiChrosolv® (Merck, Supelco, catalog number: 1.02781.2500)
Ammonium formate (Honeywell Research Chemicals, Fluka, catalog number: 55674-50g-F)
Formic acid, LC/MS grade (Fisher Chemical, catalog number: A117-50)
Ultrapure water (H2O) (Sartorius Stedim Biotech, arium pro VF)
HPLC vial (Agilent, catalog number: 5182-0716)
Additional materials required for Seahorse assays
Seahorse XFe24 V7 PS cell culture microplate (Agilent, catalog number: 100777-004)
Linoleic acid (Sigma-Aldrich, catalog number: L9530-5ML)
Oleic acid (Sigma-Aldrich, catalog number: O3008-5ML)
Oligomycin A (Sigma-Aldrich, catalog number: 209-437-3)
Carbonyl cyanide-4-(trifluoromethoxy) phenylhydrazone (FCCP) (Sigma-Aldrich, catalog number: 206-730-8)
Rotenone (Sigma-Aldrich, catalog number: 83-79-4)
Antimycin A from Streptomyces sp. (Sigma-Aldrich, catalog number: A8674)
Substrate-limited media (see Recipes)
Seahorse assay media (see Recipes)
10× drugs dilution (see Recipes)
Equipment
Seahorse XFe24 analyzer (Agilent, catalog number: 102238 or S7801A or S7801B)
Biological safety cabinet (Esco, BSC Class II)
5% CO2, 37 °C incubator
CO2-free 37 °C incubator
Sonicator (Elmasonic S 100 H, model: S100H)
Vorterxer (Thermo Scientific, model: M37610-33)
Centrifuge (Thermo Fisher Scientific, model: PICO 17)
TissueLyzer II (Qiagen)
Centrifuges for speed vacuum concentration (Labogene, model: Scan speed 40)
Cooling Trap for speed vacuum concentration (ScanLaf A/S, model: Coolsafe 110-4)
Mass spectrometry (Agilent Technologies iFunnel QTOF LC-MS, model: G6550B) (Note 2)
HPLC system (Agilent 1290 Infinity II, including High Speed Pump, Multisampler, Multicolumn Thermostat)
HPLC column, particle size of 1.8 μm, 2.1 mm × 100 mm (Agilent rapid resolution HD Zorbax SB-C18 column, catalog number: 858700-902)
Software
Seahorse Wave software (Agilent, https://www.agilent.com/en/product/cell-analysis/real-time-cell-metabolic-analysis/xf-software/seahorse-wave-desktop-software-740897)
Streamlit (Snowflake Inc., https://streamlit.io/)
Python (Python Software Foundation, https://www.python.org/)
MassHunter Qualitative Analysis 10.0 (Agilent Technologies)
MassHunter Profinder 10.0 (Agilent Technologies)
Mass Profiler Professional 15.1 (Agilent Technologies)
Procedure
We describe two protocols: one for the measurement of lipid metabolites in cells and the other to measure lipid utilization for energy production. Most of our experiments have been optimized for HEK 293FT cells, so we recommend users to optimize the cell counts if different cell types are used. Note that users using the mass spectrometry and the Seahorse XF analyzer for the first time should seek training or be accompanied by an experienced user before proceeding.
Part I: Mass spectrometry protocol
The protocol for extracting lipid metabolites is as follows:
Prepare 108 HEK 293FT cells per replicate for lipid extraction. Users can grow these cells in T75 flasks or in Petri dishes, in DMEM growth medium supplemented with 8% FBS (DMEM GM). We recommend optimizing the cell counts if different cell types are used for measurement.
Check the condition of cells before harvesting them for lipid extraction. Ensure that the cells form a confluent monolayer. Floating cells indicate cell death; users should minimize cell death before proceeding with lipid extraction.
Pre-chill extraction solvent (made of two parts of methanol mixed with five parts of Milli-Q water) at 4 °C. The solvent is pre-chilled to stop the enzymatic reactions of the cells, so as to preserve the cell states during the harvesting and extraction process.
Discard the growth media by either decanting or aspirating with a serological pipette.
Wash the cells by adding 5 mL of 1× PBS at room temperature. Discard by decanting or aspirating with a serological pipette.
Repeat the wash two times. After the final wash, discard by decanting or aspirating with a serological pipette.
Add 560 μL of pre-chilled extraction solvent to the cell culture.
Scrape cells with a cell scraper and transfer them into 2 mL Eppendorf tubes.
Add 800 μL of pre-chilled MTBE to the tubes (Note 3).
Shake the tubes using a TissueLyzer at 30 Hz for 5 min at 4 °C. Repeat two times.
Sonicate the tubes in an ultrasonic cleaner pre-filled with ice and water for 15 min. Sonication reduces emulsion formation and assist in the phase separation in liquid–liquid extraction.
Centrifuge the tubes at 845× g (3,000 rpm) for 15 min at 4 °C.
Transfer the upper MTBE layer to a new Eppendorf tube and dry the solvent with a vacuum evaporator at room temperature.
Store dried samples in a -80 °C freezer until use.
The protocol for mass spectrometry analysis is as follows:
Reconstitute dried samples in 100 μL of isopropanol/methanol (1:1) (Note 4).
Vortex rigorously for 30 s and sonicate for 5 min.
Centrifuge at 20,000× g (15,000 rpm) for 10 min at 4 °C.
Transfer the supernatant into LC vial for LC-QTOF MS analysis.
Take 5 μL of each sample for pooling into one vial as the pooled QC sample (Note 5).
Prepare mobile phase A water/acetonitrile (60:40) with 10 mM ammonium formate and 0.1% formic acid.
Prepare mobile phase B isopropanol/acetonitrile (90:10) with 10 mM ammonium formate and 0.1% formic acid (Castro-Perez et al., 2010).
Analyze samples using an Agilent rapid resolution HT Zorbax SB-C18 (2.1 mm 100 mm, 1.8 μm). The autosampler is set at 4 °C and the injection volume is 5 μL. For users who are not familiar with the protocol or instrument, we recommend doing technical replicates to ensure that the readings obtained are consistent and reproducible.
The column temperature is set at 40 °C and the flow rate is set at 0.3 mL/min.
The solvent gradient is as follows: 0–2 min, 40% solvent B; 2–12 min, 40%–100% solvent B; 12–15 min, 100% solvent B; 15–15.5 min, 100%–40% solvent B; 15.5–20 min, 40% solvent B.
Mass spectrometric analysis is performed in both ESI+ and ESI- modes using the Agilent iFunnel QTOF LC–MS (Figure 1, Note 6). Collect mass data between m/z 100 and 1,000 Dalton at a rate of two scans per second. The major parameter settings are as follows: the ion spray voltage is set at 4,000 V and the heated capillary temperature is maintained at 350 °C. The drying gas and nebulizer nitrogen gas flow rates are 12.0 L/min and 50 psi, respectively. Note that these settings may differ depending on the instrument used, so users will need to optimize their settings for different instruments.
Figure 1. Schematic diagram of Agilent QTOF mass spectrometer. Picture taken and modified from https://www.creative-proteomics.com/support/agilent-6540-uhd-quadrupole-time-of-flight-accurate-mass-mass-spectrometer.htm.
Part II: Seahorse XF substrate oxidation stress test protocol
The protocol to evaluate lipid dependencies for energy production is performed over three days. The detailed steps to execute the protocol are as follows:
Day 1
Seeding cells onto Seahorse XFe24 cell culture plate
Dilute HEK 293FT cells to 5 × 105 cells/mL and plate 200 μL of the cell suspension into each well of a SeahorseXFe24 cell culture plate, such that each well has 1 × 105 cells.
Incubate at 37 °C with 5% CO2 overnight.
Preparation of substrate-limited media and assay media
Prepare the substrate-limited media supplemented with 1 mM glutamine, 1% FBS, and either 2.5 mg/mL of BSA-conjugated linoleic acid plus 2.5 mg/mL of BSA-conjugated oleic acid or 5 mg/mL of BSA only.
Prepare the Seahorse assay media supplemented with either 2.5 mg/mL of BSA-conjugated linoleic acid plus 2.5 mg/mL of BSA-conjugated oleic acid or 5 mg/mL of BSA only.
Day 2
Exchanging cell culture growth media with substrate-limited media
Check the cells under a microscope to ensure that there is a confluent (> 90%) monolayer of cells.
Wash the cells in each condition with the respective substrate-limited media (fatty acid–supplemented media or BSA-only media as control) (Figure 2). Remove 150 μL of growth media and add 150 μL of the substrate-limited media. Each condition should be performed in at least three replicates.
Repeat the wash a second time by removing 150 μL of media but, this time, adding 200 μL of substrate-limited media to make a final volume of 250 μL of media in each well. Perform the washes gently to ensure that the cells are not dislodged.
After the cell washes, check the cells under the microscope to ensure that the cell monolayer is still intact across the wells and that no cells have been dislodged.
Leave the plate to incubate at 37 °C with 5% CO2 overnight.
Figure 2. Example of photos showing how BSA or fatty acids can be used for washing and supplemented to the Seahorse XFe24 cell culture microplate to evaluate lipid dependencies for energy production
Hydrating the sensor cartridges
Hydrate the sensor cartridges according to the manufacturer’s instructions one day prior to running the Seahorse assay.
Fill each well of the utility plate with 1 mL of XF calibrant solution.
Lower the sensor cartridge carefully into the utility plate and ensure that the sensors are submerged.
Leave the plate and cartridge to incubate in a non-CO2 37 °C incubator overnight (Figure 3). Ensure that the incubator is humidified to prevent evaporation of the calibrant.
Figure 3. An example of an XF Prep Station that can be used for incubating the plates at 37 °C overnight
Day 3
Dilute the drugs fresh on the day of assay. Refer to Recipes for details.
Wash cells with assay media. Remove 150 μL of media and replace with 150 μL of assay media for each wash. Repeat the wash three times.
After the third wash, remove 150 μL of media and add 275 μL of media to top up the volume in each well to 375 μL.
Leave the cells to equilibrate in the assay media for at least 30 min in a 5% CO2 37 °C incubator, or in a humidified chamber during transport to a Seahorse facility.
Load the drugs into the cartridge (Figure 4) with the following volumes:
Port A: oligomycin (70 μL)
Port B: FCCP (75 μL)
Port C: rotenone + antimycin A (85 μL)
Figure 4. Seahorse XF analyzer. Left: an example of a cartridge for loading the drugs into the Seahorse XF analyzer. By default, each sample is assigned to each quadrant, where Port A can be assigned at the top left, Port B at top right, and Port C at bottom left. Right: the cartridge can then be loaded in a Seahorse XF analyzer.
Turn on the Seahorse XF analyzer (Figure 4). Run the Wave software and select the mitochondria stress test assay template, which will assign ports A, B, and C to be oligomycin, FCCP, and rotenone + antimycin A, respectively. Click Design to configure the sample positions on the plate map. A standard Seahorse protocol is set up as such:
Calibrate.
Equilibrate.
Baseline readings: loop three times of mix (3 min), wait (2 min), and measure (3 min).
Inject port A: loop three times of mix (3 min), wait (2 min), and measure (3 min).
Inject port B: loop three times of mix (3 min), wait (2 min), and measure (3 min).
Inject port C: loop three times of mix (3 min), wait (2 min), and measure (3 min).
End program.
Load the sensor cartridge with calibrant plate onto the machine tray and start the calibration step.
Once the calibration is done, replace the calibrant plate with the cell culture plate and continue the run with the pre-set protocol.
Data analysis
For molecular feature (a mass spectrometry signal that represents a chemical compound) extraction, raw spectrometric data can be analyzed by MassHunter Profinder and Mass Profiler Professional software (Figure 5). The molecular features are characterized by retention time (RT), chromatographic peak intensity, and accurate mass, which can be obtained by using the Molecular Feature Extractor algorithm. A tolerance window of 0.15 min and 2 mDa was used for alignment of RT and m/z values of the features, and only features with an intensity ≥ 20,000 counts and found in at least 80% of data files in least one sample group were kept for further processing. Lipid identities were assigned based on the accurate mass measurement (mass error within ± 5 ppm) and MS/MS fragmentation patterns. Specific standards can be added to confirm the identities of the lipids.
Figure 5. Data processing pipeline of mass spectrometry data in positive mode using Agilent MassHunter Profinder and Mass Profiler Professional software
Metabolites differential analysis
Intensity values of the different metabolites from LC–MS can be visualized on heatmaps to determine their expression values between the different conditions. Users can consider performing a Z-score transformation to normalize the expression values between the different conditions for each metabolite, allowing for direct comparisons to be made across the different metabolites. Alternatively, users can normalize the expression values as log2-fold changes compared to controls. These calculations can be performed using Microsoft Excel or by using established library packages found in Python and R.
To facilitate data visualization, we have also created a web tool where users can directly upload their Excel files with expression values obtained from mass spectrometry analysis to plot clustergrams and heatmaps. The web tool can be accessed at https://kuanrongchan-lipid-metabolite-analysis-app-k4im47.streamlit.app/. To use the web tool, the first column should contain the identifiers (in this case the lipid metabolites) and the subsequent columns can be filled with expression or intensity values (Figure 6). Alternatively, users can use the demo dataset published by Yousefi et al. (2022) at https://github.com/kuanrongchan/lipid_metabolite_analysis/blob/main/Lipids_VMP1_TMEM41B_KO.csv to understand how the data file can be prepared, and to familiarize themselves with the features of the web tool. After uploading the data file, users can then use the sidebar within the web tool to assign the column(s) that contain the control samples (Figure 7). If replicates of the controls are assigned, the mean value will be used for the fold-change (log2-transformed) (log2FC) calculations. This assignment of the control group will allow the web tool to plot the log2-transformed fold-difference values in the treatment conditions as compared to the control conditions. The web tool can also plot relative values based on a filtered list of metabolites, and users can customize the dimensions of the clustergram on the side bar (Figure 2). Finally, after adjusting the settings, users can click on the finished filtering checkbox, and the clustergrams for Z-scores and log2FC values will be rendered (Figure 7). The codes for the web tool are also provided publicly at the GitHub repository at: https://github.com/kuanrongchan/lipid_metabolite_analysis/blob/main/app.py.
Figure 6. Raw data format for heatmap analysis in the web tool. File containing raw intensity values for the respective conditions from the mass spectrometry can be saved in a .xlsx or .csv format, to be subsequently used in the web tool to plot heatmap and clustermaps. Users can download this demo dataset at https://github.com/kuanrongchan/lipid_metabolite_analysis/blob/main/Lipids_VMP1_TMEM41B_KO.csv.
Figure 7. Web tool interface that renders clustergrams based on Z-score or log2-fold change values. Users can customize the settings on the left side bar and click on the Finished filtering checkbox to render the clustergram. Dataset based on Yousefi et al. (2022). Clustergram based on Z-score is shown.
Seahorse XF substrate oxidation stress test analysis
The mitochondria serve as an important organelle for energy production through oxidative phosphorylation. The mitochondrial respiration test measures the amount of oxygen consumed by the cell, and the supplementation of lipids will determine if more energy is produced via fatty acid oxidation. In addition, various inhibitors of the mitochondria respiratory chain are added to evaluate the contribution of the different mitochondrial complexes in cell respiration. The key respiratory parameters that can be measured are: (i) basal respiration, which is the initial oxygen consumption rate (OCR) minus the non-mitochondrial respiration, (ii) ATP-linked OCR, which is determined after the addition of oligomycin that inhibits ATP synthase, (iii) proton leak, which is the difference between ATP-linked OCR and non-mitochondrial respiration, (iv) maximal respiration, which is induced after addition of carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP), a potent mitochondrial oxidative phosphorylation uncoupler, (v) spare respiratory capacity, which is the difference between maximal respiration and basal respiration, and (vi) non-mitochondrial respiration, which is the OCR after addition of rotenone (complex I inhibitor) and antimycin A (complex III inhibitor). The difference in measurements between these parameters in lipid-supplemented media compared to BSA-supplemented media will indicate the contribution of the fatty acids to mitochondrial respiration (Figure 8).
Figure 8. Measurements of the different respiratory parameters in presence of BSA or with supplementation of fatty acids (Linoleic/Oleic acid) in HEK 293FT cells
In our illustrated example, we used HEK 293FT cells and evaluated the differences in fatty acid oxidation in wild-type cells compared to CRISPR-knockout cells (e.g., TMEM41B knock-out) (Figure 9). The supplementation of linoleic and oleic acid to wild-type cells increases ATP production through ATP-linked respiration. Similarly, TMEM41B-deficient cells promoted ATP production, although the maximal respiration and the spare respiratory capacity were not significantly altered with the addition of linoleic and oleic acid (Figure 9b). More details of the mechanisms of TMEM41B and VMP-1 in facilitating dengue virus infection in HEK 293FT cells can be found in the publication by Yousefi et al. (2022).
Figure 9. Measurements of the different respiratory parameters in presence of BSA or with supplementation of fatty acids (linoleic/oleic acid) in HEK-293 wild-type (WT) or TMEM41B knock-out (TMEM41B KO) cells
Notes
We recommend performing each experiment in at least triplicates to demonstrate the reproducibility and consistency of the data. Technical replicates should be considered if users are inexperienced with the protocol or instruments. Note that the experiments have been optimized in HEK 293FT cells, so if different cells are used, users will need to optimize the cell counts. Finally, the Seahorse conditions have been optimized for the 24-well format, so users will have to scale down the cells and reagents if the 96-well format is used.
High resolution LC-MS systems from other vendors (e.g., Thermo Exploris 240 Orbitrap, SCIEX triple TOF5600, Waters Xevo G2 QtoF, Shimazu LCMS-9030 QTOF) can also be used for the procedure.
MTBE is a volatile and colorless liquid, which is sparingly soluble in water. Studies have shown that MTBE can effectively replace chloroform for lipid extraction and deliver similar or better recoveries of all major lipid classes compared with the gold-standard Folch or Bligh and Dyer Recipes in which chloroform is used. The main advantage of MTBE extraction over conventional two-phase chloroform-containing solvent systems comes from the low density of the lipid-containing organic phase that forms the upper layer during phase separation. This simplifies the collection of the lipid phase and minimizes dripping losses. Furthermore, compared with chloroform, MTBE is nontoxic and noncarcinogenic.
Either chloroform/methanol or isopropanol/methanol can be used for the reconstitution of dried lipid extract. Isopropanol was selected because it is not as toxic as chloroform.
Pooled QC sample was injected periodically together with the samples to monitor the stability of the LCMS system. The coefficient of variance (CV) of each feature was calculated using the QC data and only the features with an CV < 20% were kept for the subsequent data analysis.
QTOF MS is a hybrid mass spectrometer that combines the benefits of two types of mass analyzer: quadrupole (good scan speed/sensitivity and robustness) and time-of-flight (high resolution and high mass accuracy). It offers three data acquisition modes: full scan, data-dependent acquisition, and data-independent acquisition, and is able to obtain high resolution precursor or product ion spectra. These features make QTOF MS an important tool for untargeted omics analysis.
Recipes
Substrate-limited media
Prepare two tubes of substrate-limited media by adding the following to two tubes of Seahorse XF RPMI, pH 7.4:
Add glutamine (200 mM) to a final concentration of 1 mM.
Add FBS to a concentration of 1% (v/v).
To one tube, add a 1:1 mixture of BSA-conjugated linoleic acid and BSA-conjugated oleic acid to a final concentration of 2.5 mg/mL for each fatty acid.
To the second tube, add BSA-only to a final concentration of 5 mg/mL to serve as a control. (As there are two fatty acids added into the fatty acid–supplemented media, the BSA concentration in the BSA-only media should be 2× for the BSA concentration to be equivalent.)
Filter both tubes of substrate-limited media through a 0.22 μm filter and store at 4 °C.
Seahorse assay media
Prepare fatty acid–supplemented assay media by adding a 1:1 mixture of BSA-conjugated linoleic acid and BSA-conjugated oleic acid (2.5 mg/mL each) to Seahorse XF RPMI, pH 7.4
Prepare BSA-only assay media by adding 5 mg/mL of BSA to Seahorse XF RPMI, pH 7.4.
Store both sets of media 4 °C.
10× drugs dilution
Dilute drugs stock (10,000×) to 10× on the same day as the Seahorse assay.
Oligomycin (10,000×,10 mM)
Add 30 μL (10,000×) to 2,970 μL of Seahorse XF RPMI assay media.
FCCP (10,000×, 15 mM)
Add 30 μL (10,000×) to 2,970 μL of Seahorse XF RPMI assay media.
Antimycin-A (10,000×, 10 mM) and rotenone (10,000×, 1 mM)
Add 30 μL of rotenone (10,000×) and 30 μL of antimycin A (10,000×) to 2,940 μL of Seahorse XF RPMI assay media.
Acknowledgments
We would like to thank Dr. Yap Lai Lai from the Department of Biochemistry, NUS, for her assistance with the Seahorse experiments. We would also like to acknowledge Summer Zhang for helping with the photography and Eng Eong Ooi lab for providing the technical support. The seahorse protocol is modified from Low et al. (2022). This work is funded by the Individual Research Grant (MOH-000610-00).
Competing interests
The authors describe no competing interests.
Ethics
No human subjects and samples were used in these protocols.
References
Castro-Perez, J. M., Kamphorst, J., DeGroot, J., Lafeber, F., Goshawk, J., Yu, K., Shockcor, J. P., Vreeken, R. J. and Hankemeier, T. (2010). Comprehensive LC-MS E lipidomic analysis using a shotgun approach and its application to biomarker detection and identification in osteoarthritis patients. J Proteome Res 9(5): 2377-2389.
Low, J. Z. H., Yau, C., Zhang, S. L. X., Tan, H. C., Ooi, E. E. and Chan, K. R. (2022). A protocol to assess cellular bioenergetics in flavivirus-infected cells. STAR Protoc 3(2): 101297.
Heaton, N. S., Perera, R., Berger, K. L., Khadka, S., Lacount, D. J., Kuhn, R. J. and Randall, G. (2010). Dengue virus nonstructural protein 3 redistributes fatty acid synthase to sites of viral replication and increases cellular fatty acid synthesis. Proc Natl Acad Sci U S A 107(40): 17345-17350.
Tang, W. C., Lin, R. J., Liao, C. L. and Lin, Y. L. (2014). Rab18 facilitates dengue virus infection by targeting fatty acid synthase to sites of viral replication. J Virol 88(12): 6793-6804.
Yousefi, M., Lee, W. S., Yan, B., Cui, L., Yong, C. L., Yap, X., Tay, K. S. L., Qiao, W., Tan, D., Nurazmi, N. I., et al. (2022). TMEM41B and VMP1 modulate cellular lipid and energy metabolism for facilitating dengue virus infection. PLoS Pathog 18(8): e1010763.
Zhang, J., Lan, Y. and Sanyal, S. (2017). Modulation of Lipid Droplet Metabolism-A Potential Target for Therapeutic Intervention in Flaviviridae Infections. Front Microbiol 8: 2286.
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Fluorescent Biosensor Imaging of Nitrate in Arabidopsis thaliana
YC Yen-Ning Chen
Cheng-Hsun Ho
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4743 Views: 932
Reviewed by: David PaulLip Nam LOH Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Advances Oct 2022
Abstract
Nitrate (NO3–) is an essential element and nutrient for plants and animals. Despite extensive studies on the regulation of nitrate uptake and downstream responses in various cells, our knowledge of the distribution of nitrogen forms in different root cell types and their cellular compartments is still limited. Previous physiological models have relied on in vitro biochemistry and metabolite level analysis, which limits the ability to differentiate between cell types and compartments. Here, to address this, we report a nuclear-localized, genetically encoded fluorescent biosensor, which we named nlsNitraMeter3.0, for the quantitative visualization of nitrate concentration and distribution at the cellular level in Arabidopsis thaliana. This biosensor was specifically designed for nitrate measurements, not nitrite. Through genetic engineering to create and select sensors using yeast, Xenopus oocyte, and Arabidopsis expression systems, we developed a reversible and highly specific nitrate sensor. This method, combined with fluorescence imaging systems such as confocal microscopy, allows for the understanding and monitoring of nitrate transporter activity in plant root cells in a minimally invasive manner. Furthermore, this approach enables the functional analysis of nitrate transporters and the measurement of nitrate distribution in plants, providing a valuable tool for plant biology research. In summary, we provide a protocol for sensor development and a biosensor that can be used to monitor nitrate levels in plants.
Key features
• This protocol builds upon the concept of FRET biosensors for in vivo visualization of spatiotemporal nitrate levels at a cellular resolution.
• Nitrate levels can be quantified utilizing the biosensor in conjunction with either a plate reader or a fluorescence microscope.
Graphical overview
Keywords: Nitrate Genetically encoded biosensor Yeast transformation Arabidopsis thaliana Light-sheet imaging system Visualization
Background
Genetically encoded sensors have been developed over the past decade. The first fluorescence protein–based calcium sensor, Cameleon, was created and utilized to monitor calcium signaling processes in plant stomata (Allen et al., 2001). Since then, other sensors have been developed and applied in both plant and animal biology. One example is the glucose sensors FLIPglu600μΔ13V and FLIPsuc90μ∆1V, which were created by Chen et al. (2010 and 2012, respectively). These optical sensors have proven useful not only for monitoring analyte levels and fluxes but also for identifying elusive sucrose efflux transporters required for phloem loading. Overall, genetically encoded sensors have proven to be highly versatile tools with broad applicability in a range of research settings.
In our study, we developed a Förster resonance energy transfer (FRET)-based nitrate sensor, named nlsNitraMeter3.0, to successfully monitor the steady-state levels, accumulation, and dynamic conditions of nitrate distribution and content in Arabidopsis thaliana through fluorescence confocal microscopy. Using this sensor, we can directly visualize the spatial and temporal distribution of nitrate with high resolution at the cellular level. In addition, the sensor can be used to acquire images through fluorescence microscopy before and after treatment with different media. Through image analysis software (e.g., Fiji), the images can be quantified. Furthermore, the concept and operation of nlsNitraMeter3.0 can be applied to develop new sensors that identify or characterize other molecules in plants. For instance, we previously reported that NPF1.3 plays a role as a nitrate transporter by some functional in vitro analysis (e.g., Xenopus laevis oocytes) (Chen and Ho, 2022).
Here, we provide the principles of engineering, detection methods, and application of nlsNitraMeter3.0. By following the protocol reported herein it will also be possible to develop further sensors (refer to Supplementary information).
Materials and reagents
Biological materials
Yeast strain: protease-deficient yeast strain BJ5465 (MATa, ura3–52, trp1, leu2Δ1, his3Δ200, pep4::HIS3, prb1Δ1.6R, can1, GAL+) (ATCC, catalog number: 208289TM), which was obtained from the Yeast Genetic Stock Center (University of California, Berkeley, CA).
Sensors: NIT domain/NasR
The full-length open reading frame of NasR from Klebsiella oxytoca (Boudes et al., 2012) in the pDONR221 GATEWAY Entry vector was used as a sensory domain to create the nitrate sensors NiMet3.0 and nlsNiMet3.0. Note: Constructs were inserted using an Entry clone by Gateway LR reactions into the yeast expression vectors pDRFlip30.
The NiMet3.0 is fused to the full-length of NasR with pDRFlip30 vector and sandwiched between an N-terminal Aphrodite t9 (AFPt9) variant (Deuschle et al., 2006), with nine amino acids truncated off the C terminus, and a C-terminal monomeric Cerulean (mCer) (Rizzo et al., 2006).
Note: The pDRFlip30 vector was modified from pDRFlip39 vector (Addgene, catalog number: 65517).
Agrobacterium strain: Agrobacterium tumefaciens strain GV3101 was used here to obtain high transformation rates and high levels of expression, typically leading to high copy insertion into the genome (Koncz and Schell, 1986).
Arabidopsis thaliana: wild type Col-0, a nitrate transporter mutant [npf6.3/NTR1;1/chl1-5 (Leran et al., 2014)], and a nitrate reductase mutant [nia1nia2 (Desikan et al., 2002)] were used.
Reagents
Potassium nitrate (KNO3) (Sigma-Aldrich, catalog number: P8394)
Potassium chlorate (KClO3) (STREM, catalog number: 93-1913)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P5405)
Potassium nitrite (KNO2) (ARO, catalog number: 42306)
Potassium sulfate (K2SO4) (SHOWA, catalog number: 1648-4150-000-23)
Potassium sulfite (K2SO3) (ACROS, catalog number: 44021)
Potassium selenite (K2SeO3) (STREM, catalog number: 931971-000000-18)
Potassium molybdenum oxide, anhydrous (K2MoO4) (Alfa, catalog number: 22898-0000000-17)
Ammonium chloride (NH4Cl) (Merck, catalog number: 1.01145-0500)
Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma-Aldrich, catalog number: M9272)
Gly-Gly (Sigma-Aldrich, catalog number: G1002)
YNB, yeast nitrogen base w/o amino acids w/o ammonium sulfate (BD, Difco, catalog number: 233520)
DO supplement-Ura (Takara Bio Company, Clontech, catalog number: 630416)
D-(+)-Glucose monohydrate (Fluka Analytical, catalog number: 49159)
Sucrose (Merck, catalog number: 1.07687.1000)
Agar (BD BactoTM, catalog number: DIF214530)
Agar (PhytoagarTM, catalog number: 40100072)
MES hydrate (Sigma-Aldrich, catalog number: M2933)
MOPS (Sigma-Aldrich, catalog number: M3183)
Sodium hydroxide (NaOH) (Sigma-Aldrich, catalog number: S5881)
1,4-Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: DTT-RO)
Carrier DNA [UltraPureTM salmon sperm DNA solution (Thermo Fisher Scientific, InvitrogenTM, catalog number: 115632-011)]
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: E6758)
Polyethylene glycol 4000 (PEG4000) (Fluka Analytical, catalog number: 81240)
Lithium acetate dihydrate (LiOAc) (Sigma-Aldrich, catalog number: L4158)
Tris hydrochloride ultrapure bioreagent (Tris-Cl) (J.T. Baker, catalog number: 4103-02)
Sodium chloride (NaCl) (Merck, catalog number: 1.06404.1000)
Sodium hydroxide (NaOH) (Merck, catalog number: 1.06498.1000)
Peptone (BD BactoTM, catalog number: 211677)
Yeast extract (BD BactoTM, catalog number: 212750)
MS modified basal salt mixture without nitrogen (MS) (PhytoTech Labs, catalog number: M531)
MilliQ or distilled water
Spectinomycin
Kanamycin
Solutions
40% (w/v) glucose solution (sterile and filtrated)
MOPS buffer
MES buffer
Yeast extract peptone dextrose (YPD) medium (see Recipe 1)
Solid yeast nitrogen base (see Recipe 2)
PLATE mixture (the acronym of PEG, lithium acetate, Tris, and EDTA) (see Recipe 3)
Wash buffer (see Recipe 4)
Resuspension buffer (see Recipe 5)
Substrate addition (see Recipe 6)
Plant growth solid base (see Recipe 7)
Recipes
Yeast extract peptone dextrose (YPD) mediuma, c
Reagent Final concentration Amount
Peptone 2.0% (w/v) 10 g
Yeast extract 1.0% (w/v) 5 g
Agar 2.4% (w/v) 12 g
40% sterile filtrated glucoseb 2% (w/v) 25 mL
H2O n/a n/a
Total n/a 500 mL
Autoclave, 121 °C, 15 psi, 15 min
For liquid medium, when hand-warm, add glucose from 40% sterile filtrated stock to a final concentration of 2% under a sterile hood (e.g., biosafety cabinet).
For solid medium, add 20 g/L agar before autoclaving. Add sterile filtrated glucose from 40% stock to a final concentration of 2% when the medium is hand-warm before pouring plates.
Solid yeast nitrogen base (-ura DropOut medium)a, c
Reagent Final concentration Amount
Yeast nitrogen base w/o amino acids w/o ammonium sulfate 1.7 g/L 1.7 g
DO supplement-Urad 0.77 g/L 0.77 g
40% sterile filtrated glucoseb 2% (w/v) 50 mL
H2O n/a to 1,000 mL
Total n/a 1,000 mL
Autoclave, 121 °C, 15 psi, 15 min
For liquid medium, when hand-warm, add glucose from 40% sterile filtrated stock to a final concentration of 2% under a sterile hood (e.g., biosafety cabinet).
For solid medium, add 20 g/L agar before autoclaving. Add sterile filtrated glucose from 40% stock to a final concentration of 2% when the medium is hand-warm before pouring plates.
Adjust the pH of the -ura DropOut medium to pH 5.8 with NaOH before addition of agar and autoclaving.
PLATE mixture for yeast transformation
Reagent Amount
45% PEG4000 90 mL
1 M LiOAc 10 mL
1M Tris-Cl (pH7.5) 1 mL
0.5M EDTA 0.2 mL
Total 100 mL
Wash buffer
Reagent Final concentration Amount
MES 50 mM 9.76 g
Total n/a 1,000 mL
Adjust the pH of the MES buffer to pH 5.5 with NaOH and autoclave.
Autoclave, 121 °C, 15 psi, 15 min.
Resuspension buffer
Reagent Final concentration Amount
MES buffer (Recipe 4) 50 mM 250 mL
Agar 0.05% 0.125 g
Total n/a 250 mL
Wait until the medium cools to room temperature (RT) to delay sedimentation of the cells during the measurement.
Plant growth solid basea–d
Reagent Final concentration Amount
MS salts without nitrogen 1/2 strength 0.78 g
20% sterile filtrated sucrose 0.5 % (w/v) 25 mL
Total n/a 1,000 mL
Adjust the pH to pH 5.5 with KOH before addition of agar and autoclaving.
Autoclave, 121 °C, 15 psi, 15 min
For liquid medium, when hand-warm, add sucrose from 20% sterile filtrated stock to a final concentration of 0.5% under a sterile hood (e.g., biosafety cabinet).
For solid medium, add 12 g/L PhytoagarTM before autoclaving. Add sterile filtrated glucose from 20% stock to a final concentration of 0.5% when the medium is hand-warm before pouring plates.
Substrate addition
Reagent Final concentration Amount
KNO3 or other substrates (Reagent 1–11) Depending on the experimental design -
MES buffer 50 mM -
Total n/a 50 mL
Depending on the concentration of the nitrate needed, use the nitrate stock solution and dilute it with MES buffer or MOPS buffer.
Laboratory supplies
96-well microplates (flat bottom clear or black) (Greiner Bio-One, catalog numbers: 650101 and 650209)
Note: Black plates have a lower background.
Multichannel (12) pipette (for 100 μL) (e.g., Sartorius, catalog number: 725240)
50 mL sterile plastic tubes (Falcon®)
Petri dishes (diameter: 9 mm; height: 15 mm; sterile) (Alpha Plus, catalog number: BL6905)
Glass beads (3 mm) (BasicLife, catalog number: CEO-1169)
Vacuum-driven filter system (250 mL, upper cup, 0.22 μm PES) (AGC, catalog number: AGC-VC-PES22-250-1CS)
Equipment
Monochromator-based spectrofluorimeter for 96-well plates [Safire or Infinite® M1000 (Tecan Trading)]
Cell density meter (Amersham, model: Ultrospec 10)
Orbital shaker, with temperature and velocity control (Eppendorf, New Brunswick Scientific, model: Innova 44)
Incubator for 28–30 °C incubation of yeast cells (YIHDER, model: LM-570RD)
Centrifuge with swinging rotor for 50 mL tubes (Eppendorf, model: 5810R)
Inverted confocal plus super resolution microscope (Zeiss, LSM 780 + ELYRA): high sensitivity confocal microscope is equipped with GaAsP spectrum detector, and the super resolution microscope is a Structure Illumination Microscopy.
The laboratory-established light-sheet system was made in cooperation with Microlambda Pte Ltd (Singapore)
Growth chamber
Software and datasets
MetaMorph software (Downingtown, PA)
Fiji (http://fiji.sc/)
GraphPad Prism version 9.0.0 for Mac (www.graphpad.com)
Procedure
FRET sensor design
For a detailed account of the generation of the sensor DNA constructs and the sensor mutants, please refer to Chen and Ho (2022). In this section, we just report a few concepts of the sensor design.
DNA constructs
Based on the FRET characteristic, we designed the Gateway expression clones with an insert of the bacterial (K. oxytoca) NasR/NIT domain (Figure 1).
Figure 1. Map of pDRFlip30-NasR plasmids. NasR, an NO3– binding protein, was fused via attB1 and attB2 linkers to a fluorescent protein FRET pair (donor: Aphrodite, and acceptor: Cerulean). The NasR protein (purple) representation is from a published structure of NasR [PDB 4AKK (Boudes et al., 2012)]. The Aphrodite (yellow) representation is from a published structure of Venus [PDB 1MYW (Rekas et al., 2002)] and the Cerulean (blue) representation is from a published structure of Cerulean [PDB 2WSO (Lelimousin et al., 2009)].
Expression of sensors in yeast and fluorescence analysis
Yeast transformation
The protease-deficient yeast strain BJ5465 is transformed with the sensors containing the desired above (e.g., NiMet-NIT, NiMet1.0, NiMet2.0, NiMet3.0, nlsNiMet3.0, or NiMet3.0-NRs) by using the modified lithium acetate method from Gietz et al. (1992). In brief:
Inoculate cultures in YPD medium and grow at 30 °C overnight to absorbance ~0.5 at OD 600 nm.
Spin down (2,000× g) 1 mL of cells in a microfuge tube (15 s) for each transformation.
Decant the supernatant and resuspend the cells in 100 μL of YPD medium by vortexing.
Add 2 μL of 10 mg/mL carrier DNA and vortex.
Add ~1 μg plasmid and vortex.
Add 0.5 mL of PLATE mixture [100 mL stock containing 90 mL of 45% PEG4000, 10 mL of 1 M lithium acetate, 1 mL of 1 M Tris-Cl (pH 7.5), 0.2 mL of 0.5 M EDTA] and vortex.
Add 20 μL 1 M DTT and vortex.
Incubate at 25 for 6–8 h or overnight.
Heat-shock cells for 10 min at 42 °C.
Place a pipette tip directly into the bottom of the tube, withdraw 50–100 μL of cells, and plate cells on solid -ura DropOut medium. Wrap plates with plastic cling wrap to prevent dehydration.
Incubate plates (lid down) at 30 °C for 2–3 days.
Detection of NitraMeter responses in yeast using a fluorimeter
Select transformed yeast on solid YNB supplemented with 2% glucose and -ura DropOut medium.
Pick single colonies by using sterile pipette tips and grow in a 50 mL tube containing 10 mL of -ura DropOut liquid medium. Pick at least three independent colonies.
Note: Use fresh transformation. To avoid mutations in yeast or plasmid, do not keep colonies for more than one week on plates.
Place tubes in a rack and incubate in an incubator for ~15 h under agitation (220 rpm) at 30 °C until the culture reaches absorbance ~0.5 at OD600nm.
Subculture liquid cultures after dilution to OD600nm 0.01 in the same liquid medium and grow at 30 °C until absorbance reaches ~0.3 at OD600nm.
Collect the cells by centrifugation at 3,000× g for 10 min at RT to precipitate the cells.
Discard the supernatant and resuspend the precipitate by vortexing in 10 mL of wash buffer for 15 s at RT.
Centrifuge at 3,000× g for 10 min at RT again.
Wash the precipitate two more times as in Step e–f to remove traces of growth medium.
Resuspend the precipitate to absorbance ~0.5 at OD600nm in resuspension buffer.
Mix cells well and aliquot 100 μL of the culture into wells of a 96-well flat bottom plate.
Measure the fluorescence in a fluorescence plate reader in bottom reading mode using 7.5 nm bandwidth for both excitation and emission. Typically, emission spectra are recorded with the following instrument settings: λem 470–570 nm for donor (mCer), step size 5 nm, gain: 75; and λem 520–570 nm for AFPt9, step size 5 nm, gain: 75. Fluorescence from pDRFlip30 (donor, mCer), pDRFlip39 (donor, t7.ed.eCFPt9), and pDRFlip42-linker (donor, mCer) was measured by excitation at λexc 428 nm; AFPt9 is measured with excitation at λexc 500 nm.
Use a single or multichannel pipette to add 100 μL of the culture to wells (mix by pipetting up and down) and to add analyte solution to the cells. Set up at least three replicates per treatment. Try to add equal volumes of solutions to reduce variability and use well-calibrated pipettes, since the assays are quantitative and sensitive to differences in volumes/concentration of sensor and analyte.
Record the fluorescence immediately (as fast as possible) after addition of substrate or control solution. It takes approximately 10 min to read a full 96-well plate with the parameters mentioned above. For highly accurate analyses, measure only a few wells at a time to reduce differences in analysis time. It is also possible to use instruments with injectors that allow for immediate recording; use rapid switching between wells to record over time.
Note: The sensor exhibits functional activity when employed as a purified recombinant protein.
Expression of NiMet3.0, NiMet3.0-NR-R176A, and nlsNiMet3.0 in Arabidopsis
DNA constructs for expressing sensors in plants
Insert open reading sequence of NasR or NasR-NR-R176A into the multiple cloning site of the p16-Kan vector (Jones et al., 2014): 5′-, a sequence coding for the SV40-derived nuclear localization signal LQPKKKRKVGG (Schuster et al., 2014); a sequence coding for Aphrodite; a Gateway cassette including attR1, Chloramphenicol resistance gene, ccdB terminator gene, and attR2; a sequence coding for mCerulean (mCer); and a sequence coding for the cMyc epitope tag -3′, or pZPFlip UBQ10-KAN vector under the control of the UBQ10 promoter.
Note: The p16 promoter (Schuster et al., 2014) from the AT3G60245 gene encoding a 16S ribosomal subunit was used to drive the nuclear-localized NiMet3.0 fusion biosensor, whereas the CaMV 35S promoter (Battraw and Hall, 1990) was used to drive the NiMet3.0 and NiMet3.0-NR-R176A fusion biosensor in plants.
Recombine in Gateway LR reactions with NasR or NasR-NR-R176A Entry Clones, resulting in NiMet3.0, NiMet3.0-NR-R176A, and nlsNiMet3.0 expression clones.
Generate transgenic plants using the Agrobacterium floral dip method
Introduce sensors into Agrobacterium tumefaciens GV3101.
Grow healthy Arabidopsis plants in 12 h of light, 50% humidity, and at 22 °C until they begin to bolt and produce floral inflorescences (3–4 weeks in a growth chamber).
Remove siliques and mature flower clusters before floral dipping.
Inoculate a single Agrobacterium colony that was transformed with sensors into 5 mL of liquid LB medium containing the appropriate antibiotics [spectinomycin (final concentration 100 μg/mL)] for binary vector selection. Incubate the culture at 28 °C overnight.
The following morning, use this feeder culture to inoculate 200 mL of liquid LB with spectinomycin (final concentration 100 μg/mL) and grow the culture at 28 °C for 16–24 h.
Collect Agrobacterium cells by centrifugation at 3,000× g for 10 min at RT and discard the supernatant. Then, gently resuspend cells in one volume of the freshly made dipping medium.
Dilute Agrobacterium cells to 6 × 109 cells/mL.
Spray the Agrobacterium on the floral part of the Arabidopsis. Then, lay down the dipped plants in a plastic basin and cover them with plastic wrap for 16–24 h to maintain high humidity.
The next day, remove the cover and allow them to grow normally for one month in the greenhouse or the growth chamber; withhold watering when siliques turn brown.
Select transformants on agar plates containing 1/2× MS medium with vitamins (PhytoTech Labs, M519) and kanamycin (30 mg/L).
Imaging the nitrate sensor in Arabidopsis with fluorescence microscopy confocal microscopy
Note: Although a fluorescence confocal microscope is the standard equipment used, light-sheet microscopy is another option. The settings for laser intensity, detector, and objective are similar to those for confocal microscopy. Please refer to the detailed procedure of the light-sheet system in the Supplementary information section.
Germinate and grow transgenic seedlings
Germinate and grow vertically on 1/2× MS modified basal salt mixture without nitrogen, 1% agar, and 0.05% (w/v) sucrose (pH 5.7) plates.
Place 5- or 6-day-old seedlings in the solution containing 1/2× MS medium [1/2× MS and 0.05% sucrose (pH 5.7)] and prepare for imaging on glass slides.
Nitrate treatments on glass slides for confocal microscopy
Place seedlings on glass slides with 50 μL of solution, surround with a rectangle of vacuum grease, and cover with a square coverslip equal in height and half the width of the vacuum grease rectangle.
Exchange the nitrate treatment solution beneath the coverslip by addition to the left and removal from the right side of the coverslip.
Acquire confocal images on a Zeiss 780 laser scanning microscope and use a 20×/0.8 Plan-Apochromat dry objective or 40×/1.2 C-Apochromat water objective. Excite CFP (440 nm) and yellow fluorescent protein (YFP; 514 nm) with lasers. Detect fluorescence emission using a GaAsP photomultiplier tube (PMT) detector, set to detect 463–508 nm for CFP, and a normal PMT detector, set to 520–585 nm for YFP. Set the laser power between 0.5% and 2% with detector gain set to 700–750 to image CFP or YFP.
Acquire images at time points based on the purpose of the research (refer to the note below for details on time point settings). Acquire three-dimensional images, with a z-step size of 1.5 μm and half the diameter of the primary root axis in Arabidopsis.
Notes:
i. For example, if the purpose is to observe the nitrate distribution in the root after different concentrations of medium supplement, it is suitable to set the range and interval of the time points to less than an hour, unless the sample can be kept moist. Additionally, fluorescence blenching should be considered when continuously recording.
ii. For other methods that can be used to obtain continuous images or video, please refer to the Supplementary information.
Data analysis
Fluorescence emission ratio response of purified NiMet3.0 to NO3− in vitro
Subtract background fluorescence of yeast (using cells transformed with vector only) from all fluorescence values (for both spectra as well as single point measurements).
The solution addition might trigger a change in the energy transfer rate between the emission at 530 nm [Dx acceptor emission (DxAm)] and the emission at 488 nm [Dx donor emission (DxDm)] that could act as a FRET ratio change sensor (ΔDxAm/DxDm). Through several optimizations, we obtained a fusion construct that shows a significantly substrate-triggered positive ratio change (ΔDxAm/DxDm) (e.g., NiMet3.0) (Figure 2).
Notes:
NiMet3.0 expressed in yeast responds to nitrate addition by changing the fluorescence intensity of donor and acceptor emission (obtained with excitation at 428 nm). Aphrodite-t9 emission was unaffected and served as a control or reference for normalization (obtained at 500 nm excitation). Nitrate addition (5 mM) induced a decrease in the emission spectrum of the donor, and the emission of the acceptor increased (Figure 2A). Besides, since the Aphrodite-t9 emission is unaffected by nitrate when excited directly, Aphrodite-t9 emission can be used as a control and for normalization by using ratios instead of absolute values to compare between different cultures.
The various nitrate concentrations from micromolar to millimolar were added externally to the primary root to monitor the NitraMeter sensor responses. The data showed that the FRET ratio changed to external nitrate addition was saturated after approximately 0.25–0.5 mM, indicating either that the NitraMeter sensor in root was all occupied by nitrate or the Vmax of NitraMeter sensor was reached after the concentrations of nitrate addition externally.
Figure 2. Fluorescence response of NiMet 3.0- and nlsNiMet 3.0-expressing yeast cells and Arabidopsis root. (A) Fluorescence emission wavelength scan (B) and emission ratio at 530 nm of purified NiMet3.0 protein with and without NO3−. Nitrate concentration is indicated in the figures. Nitrate was able to trigger responses that were significantly different from the control (*, P < 0.0001, t-test). The presented data are mean ± SD of six biological repeats. (C) Three-dimensional images of nlsNiMet3.0 emission ratios of 5-day-old root meristem zone in transgenic Col-0 before a NO3− pulse, after the NO3− pulse, and after removing the NO3−. NO3− (50 μM) was used. (D) Beeswarm and box plot of NO3− concentration-dependent nlsNiMet3.0 emission ratios for nuclei of root tips. ****, P < 0.0001, Student’s t-test. Means ± SD of three biological repeats are presented.
Quantification of the ratio of the fluorescence pixel intensity of the nitrate sensor in vivo
Use Fiji software to process the image and quantify the fluorescence pixel intensity. Calculate the mean gray values of regions of interest (ROIs) within the root meristem region.
Subtract the background from all measured intensities generated by ROIs where there was no plant material.
Measure the mean intensity values in all four channels (Dx/Dm, Dx/Am, Ax/Dm, and Ax/Am), and subtract that intensity from the entire image.
Create ratio images (DxAm/DxDm) by using the Ratio Plus plug-in for ImageJ (P. Magalhães, University of Padua, Italy) (Figure 2C). Select and analyze ROIs with the help of the ROI manager tool.
Import the fluorescence pixel intensity to GraphPad Prism to generate figures (e.g., Figure 2D) and present the data following the same setting rules as described below (see General notes).
General notes and troubleshooting
General notes
Despite the NitraMeter3.0 being able to visualize the nitrate dynamic within the cells in Arabidopsis thaliana, investigation of nutrient acquisition has relied heavily on techniques that integrate uptake over the entire root system. It is worth to note that the responses of NitraMeter to nitrate indicate the net fluxes of nitrate within where the NitraMeter is located in cell. In addition, net fluxes of NO3– into the roots vary both with position along the root axis and with time. These variations may not be consistent in different plants, in which different cells in different roots may not show exact temporal and spatial patterns of nitrate dynamics.
Present the data by using beeswarm and box plots of raw data. In the beeswarm and box plot graphs, the central rectangle spans the first quartile to the third quartile, while the line inside the rectangle shows the median. The whiskers denote 1.5 interquartile ranges from the box, and outlying values are plotted beyond the whiskers. Perform statistical analyses using GraphPad Prism version 9.0.0 for Mac (www.graphpad.com).
Acknowledgments
We acknowledge W. B. Frommer for discussion and suggestions. We thank Addgene for distributing the plasmids donated by W. B. Frommer. We thank the Advanced Optics Microscope Core Facility at Academia Sinica for technical support for fluorescence imaging. The core facility is funded by Academia Sinica Core Facility and Innovative Instrument Project (AS-CFII-108-116). We thank M.-J. Hung for efforts on this project. We thank A. K. Snyder and M. Loney for English editing.
Funding: This work was supported by Academia Sinica, Taiwan; Ministry of Science and Technology, Taiwan, grant 105-2311-B-001- 045 (C.-H.H.); and Ministry of Science and Technology, Taiwan, grant106-2311-B-001-037-MY3 (C.-H.H.). This protocol was derived from the original work of Chen et al. (2022).
Competing interests
The authors declare that they have no competing interests.
References
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Supplementary information
The supporting information can be downloaded here.
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© 2023 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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In Vitro Analysis of Stalled Ribosomes using Puromycin Incorporation
MS MaKenzie R. Scarpitti
MK Michael G. Kearse
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4744 Views: 1316
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Original Research Article:
The authors used this protocol in The Journal of Biological Chemistry Dec 2022
Abstract
Ribosome footprint profiling has demonstrated that ribosomes can be slowed or stalled on select mRNAs, often due to the presence of rare codons, stalling motifs, or via a ribosome-binding protein (e.g., FMRP). Stalled ribosomes can act as physical roadblocks for trailing ribosomes and ultimately can cause ribosome collisions that stimulate no-go mRNA decay. Detecting stalled or slowed ribosomes in cells by ribosome footprint profiling or classic polysome profiling is laborious, technically challenging, and low throughput. Here, we present a protocol to assay for stalled ribosomes on in vitro–transcribed reporter mRNAs using a robust, commercially available mammalian in vitro translation lysate and an optimized low-speed sucrose cushion. In short, we take advantage of the ability of puromycin to incorporate into the nascent polypeptide and cause the ribosome to dissociate from the mRNA during active elongation, as well as the ability to selectively pellet ribosomes through a low-speed sucrose cushion due to their large molecular weight. Stalled ribosomes are not actively elongating and do not incorporate puromycin, allowing the ribosome-bound mRNA to pellet in the low-speed sucrose cushion. RT-qPCR is used to quantify the amount of ribosome-bound reporter mRNA in the pellet. This workflow allows for direct assessment of stalled ribosomes and is fully amendable to insertion of putative stalling motifs in the target mRNA, as well as supplementation with recombinant proteins or small molecule inhibitors that target translation elongation.
Key features
•This protocol is optimized for cap-dependent in vitro translation in the dynamic linear range.
•Details for generating capped reporter mRNA in one day are provided.
•Requires as little as one day to complete if starting with in vitro–transcribed mRNA.
•This protocol requires access to an ultracentrifuge and a real-time PCR system.
Graphical overview
Keywords: Cycloheximide Elongation inhibition Polysomes Protein synthesis Sucrose cushion Ultracentrifugation
Background
Upon initiating at a start codon, ribosomes proceed through elongation with repetitive cycles of decoding and translocation until they terminate at one of three stop codons (UAA, UAG, or UGA) (Dever et al., 2018). Elongating ribosomes may encounter a multitude of challenges, including rare codons, premature polyadenylation, truncated or damaged mRNA, proline-rich stalling motifs, strong or highly ordered mRNA structure, or mRNA- and ribosome-binding proteins (e.g., FMRP) (Kim and Zaher, 2022). These obstacles can stall ribosomes, which act as physical roadblocks for trailing ribosomes, resulting in ribosome collisions that stimulate the no-go mRNA decay pathway. In yeast, collided ribosomes are recognized by the E3 ligase Hel2 (ZNF598 in human and Caenorhabditis elegans) (Monem et al., 2023), which ubiquitinates ribosomal proteins eS10 and uS10 (Juszkiewicz et al., 2018). In yeast, ubiquitinated and collided ribosomes serve as a unique binding site for the endonuclease Cue2 (N4BP2 in humans, NONU1 in Caenorhabditis elegans) (D’Orazio et al., 2019). Subsequently, the new 5′ and 3′ ends are susceptible to degradation by Xrn1 and the exosome, respectively. Ribosome collisions also drive mRNA-specific feedback translation initiation inhibition to further prevent synthesis of deleterious truncated proteins (Juszkiewicz et al., 2020; Sinha et al., 2020). Upon large-scale ribosome collisions, global translation initiation is inhibited by activation of GCN2-mediated eIF2α phosphorylation (Wu et al., 2020).
Ribosome footprint profiling and classic polysome profiling can be used to detect stalled ribosomes; however, these approaches can be technically challenging, laborious, and rather low throughput. Additionally, ribosome profiling is not cost-effective when testing multiple specific mutations within reporter mRNAs or effector proteins. Here, we present a validated protocol that can be used to assess ribosome stalling in vitro that is medium-to-high throughput and can be performed in as little as one day if starting with in vitro–transcribed mRNA.
We take advantage of the selective nature of puromycin, an amino-acyl transfer RNA analog to incorporate into nascent polypeptides of only actively elongating ribosomes (Yarmolinsky and Haba, 1959). Puromycin incorporation results in ribosomes releasing both the nascent protein and the mRNA, resulting in the collapse of polysomes to monosomes (Yarmolinsky and Haba, 1959; Azzam and Algranati, 1973; Stefani et al., 2004; Sivan et al., 2007). Using an optimized low-speed sucrose cushion and subsequent RT-qPCR, we quantify ribosome-bound reporter mRNA to determine efficacy of puromycin to dissociate ribosomes. Ribosomes that are stalled during elongation do not actively incorporate puromycin; thus, these ribosomes are insensitive to puromycin and consistently pellet bound mRNA in the presence of puromycin. This protocol utilizes a commercially available mammalian translation lysate in conditions that allow for translation in the dynamic linear range and showcases cap and scanning dependency (Kearse et al., 2016). A key feature is an independent reporter that is translated and not treated with puromycin, which serves as a normalizing control for both the low-speed sucrose cushion and RT-qPCR. The low-speed sucrose cushion has been optimized so that only ribosome-bound mRNA is pelleted. Untranslated mRNA does not have sufficient molecular weight to pellet in these conditions. We have published this strategy and validated its efficacy using the Fragile X protein FMRP (an mRNA- and ribosome-binding protein that stalls ribosomes) (Darnell et al., 2011), along with FireFly Luciferase mRNA as the normalizing control reporter and nanoLuciferase mRNA as the experimental reporter (Figure 1) (Scarpitti et al., 2022).
Figure 1. Schematic of overall strategy to assay for stalled ribosomes using puromycin incorporation and an optimized low-speed sucrose cushion.CHX = Cycloheximide.
Materials and reagents
Reagents
2-Propanol (isopropanol) (Fisher Chemical, catalog number: A416P-4)
3′-O-Me-m7G(5′)ppp(5′)G RNA Cap Structure Analog (anti-reverse cap analog; ARCA) (New England Biolabs, catalog number: S1411S)
Agarose LE, quick dissolve (Apex BioResearch Products, catalog number: 20-102QD)
Bromophenol blue (Bio-Rad, catalog number: 1610404)
Chloroform, ethanol stabilized (Millipore Sigma, catalog number: 67-66-3)
10× CutSmart buffer (New England Biolabs, catalog number: B7204S)
Cycloheximide (CHX) (Sigma, catalog number: C1988)
Dimethyl sulfoxide (DMSO) (Millipore Sigma, catalog number: MX1458-3)
Dithiothreitol (DTT) (Thermo Scientific, catalog number: 20290)
DNA Clean & Concentrator-25 kit (Zymo Research, catalog number: 11-305C)
DNase I (RNase-free) (New England Biolabs, catalog number: M0303L)
E. coli Poly(A) polymerase (New England Biolabs, catalog number: M0276L)
500 mM EDTA, pH 8.0 ULTROL grade (Millipore, catalog number: 324504)
Ethanol, 200 proof (Decon Laboratories, Inc., catalog number: 64-17-5)
10 mg/mL ethidium bromide (Thermo Scientific, catalog number: 15585011)
Flexi Rabbit Reticulocyte Lysate System (Promega, catalog number: L4540)
37% (w/v) formaldehyde (Fisher Chemical, catalog number: F79-500)
Glycerol, biotechnology grade (Amresco, catalog number: 0854-4L)
Glycogen, molecular biology grade (Thermo Scientific, catalog number: R0561)
Hi-Di Formamide (Thermo Scientific, Applied Biosystems, catalog number: 4401457)
HiScribe T7 High Yield RNA Synthesis kit (New England Biolabs, catalog number: E2040S)
iScript Reverse Transcription Supermix (Bio-Rad, catalog number: 1708841)
iTaq Universal SYBR Green Supermix, 5 mL (Bio-Rad, catalog number: 1725124)
2 M KCl, pH 7.4 (MOLTOX, catalog number: 26-516.5)
1 M MgCl2 (Thermo Scientific, catalog number: AM9530G)
Millennium RNA size marker (Thermo Scientific, catalog number: AM7150)
10× MOPS buffer (KD Medical, catalog number: RGF-6170)
Nuclease-free water (Thermo Scientific, catalog number: AM9937)
Puromycin dihydrochloride (Sigma, catalog number: P8833)
6× purple gel loading dye (New England Biolabs, catalog number: B7024A)
Quick-Load Purple 1 kb Plus DNA ladder (New England Biolabs, catalog number: N0550S)
RNA Clean & Concentrator-25 kit (Zymo Research, catalog number: 11-353B)
RNase inhibitor, murine (New England Biolabs, catalog number: M0314L)
Sucrose, Ultra-pure, RNase- & DNase-free (VWR, catalog number: 57-50-1)
10× TBE buffer (IBI Scientific, catalog number: IB70154)
1 M Tris-HCl, pH 7.4 (Apex BioResearch Products, catalog number: 18-189)
TRIzol reagent (Ambion, catalog number: 15596018)
Xylene cyanol FF (Bio-Rad, catalog number: 1610423)
pcDNA3.1(+)/nLuc-3XFLAG (Addgene, catalog number: 127299)
Dual promoter plasmid pCR II (Thermo Scientific, catalog number: K207040)
pGL4.13 (Promega, catalog number: E6681)
Solutions
100 mg/mL cycloheximide (CHX) (see Recipes)
10 mg/mL CHX (see Recipes)
5 mg/mL CHX (see Recipes)
1 M DTT, Cleland’s Reagent (see Recipes)
70% (v/v) ethanol (see Recipes)
1 mg/mL ethidium bromide (see Recipes)
10 mM GTP (see Recipes)
1× MOPS buffer (see Recipes)
10 mg/mL (~18 mM) puromycin (see Recipes)
0.6 mM puromycin (see Recipes)
2× ribosome dilution buffer (see Recipes)
RNA loading dye (see Recipes)
RNA sample buffer (see Recipes)
60% (w/v) sucrose (see Recipes)
35% (w/v) sucrose, buffered (see Recipes)
1× TBE buffer (see Recipes)
Recipes
100 mg/mL cycloheximide (CHX) (store at -20 °C)
Reagent Final concentration Quantity
CHX 100 mg/mL 1 g
DMSO n/a To 10 mL
Total n/a 10 mL
10 mg/mL CHX (working solution, aliquot into single-use vials and store at -20 °C)
Reagent Final concentration Quantity
100 mg/mL CHX 10 mg/mL 1 mL
Milli-Q water n/a 9 mL
Total n/a 10 mL
5 mg/mL CHX (working solution, aliquot into single-use vials and store at -20 °C)
Reagent Final concentration Quantity
10 mg/mL CHX 5 mg/mL 3 mL
Milli-Q water n/a 3 mL
Total n/a 6 mL
1 M DTT, Cleland’s Reagent (store at -20 °C)
Reagent Final concentration Quantity
DTT 1 M 1.542 g
Milli-Q water n/a To 10 mL
Total n/a 10 mL
70% (v/v) ethanol
Reagent Final concentration Quantity
100% Ethanol 70% (v/v) 35 mL
Milli-Q water n/a 15 mL
Total n/a 50 mL
1 mg/mL ethidium bromide
Reagent Final concentration Quantity
10 mg/mL ethidium bromide 1 mg/mL 100 μL
Milli-Q water n/a 900 μL
Total n/a 1 mL
10 mM GTP (store at -20 °C)
Reagent Final concentration Quantity
100 mM GTP (from HiScribe T7 High Yield RNA Synthesis kit) 10 mM 50 μL
Nuclease-free water n/a 450 μL
Total n/a 500 μL
1× MOPS buffer
Reagent Final concentration Quantity
10× MOPS buffer 1× 60 mL
Milli-Q water n/a 540 mL
Total n/a 600 mL
10 mg/mL (~18 mM) puromycin (store at -20 °C)
Reagent Final concentration Quantity
Puromycin 10 mg/mL 1 g
Milli-Q water n/a To 100 mL
Total n/a 100 mL
0.6 mM puromycin (store at -20 °C)
Reagent Final concentration Quantity
18 mM puromycin 0.6 mM 66.67 μL
Milli-Q water n/a 1.933 mL
Total n/a 2 mL
2× ribosome dilution buffer (store at 4 °C)
Reagent Final concentration Quantity
1 M Tris-HCl, pH 7.4 40 mM 2 mL
2 M KCl 280 mM 7 mL
1 M MgCl2 20 mM 1 mL
100 mg/mL CHX 0.2 mg/mL 100 μL *Add day of use
1 M DTT 2 mM 100 μL *Add day of use
Milli-Q water n/a 39.8 mL
Total n/a 50 mL
RNA loading dye
Reagent Final concentration Quantity
Glycerol 50% (v/v) 5 mL
500 mM EDTA, pH 8.0 100 mM 2 mL
Bromophenol blue 2.5 mg/mL 25 mg
Xylene cyanol FF 2.5 mg/mL 25 mg
Milli-Q water n/a 3 mL
Total n/a 10 mL
RNA sample buffer (make fresh day of use)
Reagent Final concentration Quantity
37% (w/v) formaldehyde 9% (w/v) 35 μL
Hi-Di formamide 69% (v/v) 100 μL
10× MOPS buffer 0.7× 10 μL
Total n/a 145 μL
60% (w/v) sucrose
Reagent Final concentration Quantity
Sucrose 60% (w/v) 300 g
Milli-Q water n/a To 500 mL
Total n/a 500 mL
35% (w/v) sucrose, buffered (store at 4 °C)
Reagent Final concentration Quantity
60% sucrose (w/v) 35% (w/v) 29.167 mL
1 M Tris-HCl, pH 7.4 20 mM 1 mL
2 M KCl 140 mM 3.5 mL
1 M MgCl2 10 mM 0.5 mL
100 mg/mL CHX 0.1 mg/mL 50 μL *Add day of use
1M DTT 1 mM 50 μL *Add day of use
Milli-Q water n/a 15.733 mL
Total n/a 50 mL
1× TBE buffer
Reagent Final concentration Quantity
10× TBE buffer 1× 100 mL
Milli-Q water n/a 900 mL
Total n/a 1 L
Laboratory supplies
0.2 mL, open-top thick wall polycarbonate tube, 7 mm × 20 mm (Beckman Coulter, catalog number: 343775)
1.7 mL microcentrifuge tube, clear (Olympus Plastics, catalog number: 24-282)
500 mL Erlenmeyer flask
8-strip PCR tubes (Olympus Plastics, catalog number: 27-125UA)
Hard-shell PCR plates, 96-well, thin-well (Bio-Rad, catalog number: HSP9601)
Ice
Ice buckets
Light-dry tissue wipes (VWR, catalog number: 82003-820)
Microseal ‘B’ seals (Bio-Rad, catalog number: MSB1001)
Nitrile gloves
P10, P20, P200, and P1000 calibrated pipettes
P10, P20, P200, and P1000 pipette tips (VWR, catalog numbers: 76323-388, 76323-390, and 76323-456)
Parafilm
Plastic wrap
Tube racks
Equipment
-80 °C freezer
-20 °C freezer
4 °C refrigerator
Aspirator
CFX Connect Real-Time System (Bio-Rad, catalog number: 1855201)
Eppendorf centrifuge 5430 (Eppendorf, catalog number: 022620509)
Fume hood
GelDoc Go Gel imaging system with Image Lab Touch software (Bio-Rad, catalog number: 12009077)
Microwave
MilliporeSigma Synergy ultrapure water purification system (Fischer Scientific, catalog number: SYNS0HFUS)
NanoDrop One Microvolume UV-Vis spectrophotometer (Thermo Scientific, catalog number: ND-ONE-W)
OWL EasyCast B1 Mini Gel electrophoresis system (Thermo Scientific, catalog number: B1-BP)
OWL EasyCast B1A Mini Gel electrophoresis system (Thermo Scientific, catalog number: 09-528-110)
Paper towels
Plate centrifuge, PerfectSpin P (VWR Peqlab, catalog number: PEQL91-PSPIN-P)
Pointed tweezers
S100-AT3 fixed angle rotor (Thermo Scientific, catalog number: 45585)
Sorvall Discovery M120 SE micro-ultracentrifuge (Hitachi)
T100 thermal cycler (Bio-Rad, catalog number: 1861096)
Vortex mixer (Benchmark, catalog number: BV1000)
Software and datasets
Bio-Rad CFX Maestro
Microsoft Excel
Prism GraphPad Molarity Calculator (https://www.graphpad.com/quickcalcs/molarityform/)
Oligo Calc: Oligonucleotide Properties Calculator (http://biotools.nubic.northwestern.edu/OligoCalc.html)
Transcription and Translation Tool (https://biomodel.uah.es/en/lab/cybertory/analysis/trans.htm)
Procedure
In vitro transcription of ARCA-capped reporter mRNAs
Linearize normalizing control and experimental reporter plasmids by restriction enzyme digestion. Separate reporters can be linearized in parallel.
Notes:
i. For the normalizing control reporter, we typically use FireFly Luciferase (FFLuc) subcloned from pGL4.13 into pCR II, which is linearized with HindIII. For the experimental reporter, we typically use nanoLuciferase (nLuc) subcloned from pNL1.1 into pcDNA3.1(+), which is linearized with either XbaI or PspOMI. Both plasmids are available upon request from the corresponding author; pcDNA3.1(+)/nLuc-3XFLAG is also available from Addgene.
ii. Choose a restriction endonuclease that cleaves downstream of the reporter coding sequence and either produces blunt ends or 5 overhangs. Restriction endonucleases that produce 3′ overhangs should be avoided as they can elicit aberrant and antisense transcription (Schenborn and Mierendorf, 1985). Choose a restriction enzyme that will produce the desired 3′ untranslated region (UTR). For example, linearizing pcDNA3.1(+)/nLuc via either XbaI or PspOMI produces a short (10 base or 16 base, respectively) 3′ UTR that works very well in our hands. We have not tested if shorter 3′ UTRs affect translation efficiency of reporter mRNAs in this system, nor have we tested if the length of the 3′ UTR affects the sensitivity of this protocol. In general, 3′ UTRs typically contain inhibitory and stimulatory translational control elements; thus, smaller 3′ UTRs will contain fewer unexpected translational control elements.
In a microcentrifuge tube, combine 100 μL of plasmid DNA (300 ng/μL), 20 μL of 10× CutSmart buffer, 20 μL of restriction enzyme (400–800 U total), and 60 μL of Milli-Q water. Gently mix by inversion 20 times and collect contents by a short centrifugation spin (e.g., 2–3 s in a minicentrifuge or microcentrifuge).
Digest the plasmid for 4 h at 37 °C and then hold at 4 °C. Pause point: Store completed digest at -20 °C.
Note: Despite the excess of restriction enzyme in the reaction, we suggest a 4 h incubation for complete linearization due to the relatively large mass of plasmid DNA.
During the restriction digest, cast a 0.8% (w/v) agarose gel in an OWL EasyCast B1A Mini Gel electrophoresis system with a 1.5 mm 10-well comb. Add 100 mL of 1× TBE buffer with 0.8 g of agarose in a 500 mL Erlenmeyer flask and then loosely plug the flask with a folded-up paper towel. Heat in the microwave for 1.5 min or until the agarose is completely dissolved. Allow to cool for 10 min on the benchtop and then add 5 μL of 10 mg/mL ethidium bromide. Gently swirl to mix and let the flask cool on the benchtop until it can be held comfortably for 10 s (or place in a 65 °C water or bead bath for 30 min). Pour ~70 mL into the casting tray and let the gel solidify at room temperature for ~1 h. Remove combs by gently pulling them up vertically. Remove and rotate the casting tray so that the wells are near the cathode. Fill the gel tank and completely cover the agarose gel with 1× TBE buffer (~350 mL total).
Purify linearized plasmids using the DNA Clean and Concentrator-25 kit and the supplied solutions.
Perform all steps at room temperature and centrifugation at 16,000× g for 30 s, unless specified. To the 200 μL digest, add 800 μL of DNA binding buffer and mix by gentle inversion 20 times. Transfer mixture to the Zymo-spin column in a collection tube. Centrifuge and discard flowthrough. Wash the column with 200 μL of DNA wash buffer by centrifugation. Discard flowthrough and repeat wash step. Transfer column into a fresh and pre-labeled 1.7 mL microcentrifuge tube. Add 30 μL of nuclease-free water directly to the column (the white resin at the bottom of the column) and incubate at room temperature for 1 min. Elute by centrifugation (1 min, 16,000× g, room temperature). Store on ice for immediate use or at -20 °C for long-term storage.
Determine DNA concentration and purity by UV spectrophotometry (e.g., NanoDrop).
Note: The Zymo-spin IICR column has a reported capacity of 25 μg of DNA. This protocol calls for 30 μg of DNA, which allows us to max out the column, resulting in most reporter plasmids eluting at ~1,000 ng/μL.
Confirm linear plasmid integrity by 0.8% (w/v) agarose gel electrophoresis. Add 5 μL of Quick-Load Purple 1 kb Plus DNA ladder to the first well. On a piece of parafilm or in a microcentrifuge tube, gently mix 1 μL of purified linear plasmid DNA (250–500 ng/μL), 1 μL of purple gel loading dye (6×), and 4 μL of Milli-Q water, and then load the entire sample into a single well. Run the agarose gel at 120 V (constant) for 30–45 min or until the desired resolution. Image gel via UV transillumination to confirm a single band that is running at the expected molecular weight.
Note: If using two combs per casting tray, use the bottom half of the agarose gel before using the top half, as ethidium bromide in the gel runs toward the cathode. Unused parts of the gel may be stored for up to one week in plastic wrap at 4 °C.
Synthesize in vitro–transcribed reporter mRNA using the HiScribe T7 High Yield RNA Synthesis kit.
To resuspend the ARCA cap analog to 40 mM, first quickly collect the contents (1 μmol) by a short centrifugation and then add 25 μL of nuclease-free water to the pellet. Gently mix by flicking the tube and collect the contents by brief centrifugation. Repeat the gently mixing procedure for a total of five times.
For a typical 10 μL reaction, combine on ice 1 μL of linear reporter plasmid (500 ng/μL), 0.5 μL of RNase inhibitor, 1 μL of 10× T7 reaction buffer, 1 μL of 100 mM ATP, 1 μL of 100 mM UTP, 1 μL of 100 mM CTP, 1 μL of 10 mM GTP, 2 μL of 40 mM ARCA cap analog, 1 μL of T7 RNA polymerase mix, and 0.5 μL of nuclease-free water in a PCR tube or microcentrifuge tube. It is critical to use 1 μL of 10 mM GTP and not 1 μL of 100 mM GTP. The 8:1 ARCA cap analog to GTP ratio ensures 90% co-transcriptional capping efficiency (Krieg and Melton, 1987).
Notes:
i. If transcribing multiple mRNAs at one time, assemble a master mix of all components except linear plasmid templates. Use 9 μL of master mix with 1 μL of linear plasmid template (~500 ng/μL). Reactions can also be scaled up linearly to 20 μL but beware of exceeding the binding capacity of the RNA clean up columns (see below).
ii. Most T7 RNA polymerase–mediated mRNA synthesis kits with a cap analog pre-mixed with the NTPs use a 4:1 ARCA cap to GTP ratio, which only provides ~80% co-transcriptional capping efficiency. These cap analog pre-mixed kits also do not provide the ability to generate A-capped or non-methylated G-capped mRNAs to test cap-dependency.
Perform in vitro transcription for 2 h at 30 °C using a thermal cycler (for PCR tube) or heat block (for microcentrifuge tube).
Note: In our hands, transcription at 30 °C yielded purer mRNA than transcription at 37 °C.
To remove the DNA template, add 1 μL of DNase I (RNase-free) to each 10 μL reaction and gently mix by inversion. Incubate at 37 °C for 15 min.
If polyadenylation of the RNA is desired, combine and gently mix on ice the 11 μL DNase-treated capped and transcribed mRNA reaction with 5 μL of 10× Poly(A) polymerase buffer, 5 μL of 10 mM ATP, 1 μL of E. coli Poly(A) polymerase, and 28 μL of nuclease-free water, followed by incubation at 37 °C for 1 h.
Note: This protocol uses the nuclease-treated Flexi rabbit reticulocyte lysate (RRL) system from Promega. It is not entirely necessary to polyadenylate in vitro–transcribed mRNA for efficient translation using nuclease-treated RRL. Polyadenylation provides a less than two-fold enhancement of reporter mRNA translation in nuclease-treated RRL (Soto Rifo, 2007). If polyadenylation is skipped, add 39 μL of nuclease-free water to bring the sample volume to 50 μL. It is worth noting that the described cDNA synthesis reagents below use both random hexamers and oligo-dT primers. If using different reagents that only contain oligo-dT primers for cDNA synthesis, polyadenylation is required for detection of pelleted, ribosome-bound mRNA.
If using a PCR tube, transfer the entire contents to a microcentrifuge tube.
Purify mRNA using the RNA Clean and Concentrator-25 kit and the supplied solutions.
Perform all steps at room temperature and centrifugation at 16,000× g for 1 min, unless specified. To each 50 μL of mRNA sample, add 100 μL of RNA binding buffer and gently mix by inversion 20 times. Add 150 μL of 100% ethanol to each reaction and gently mix by inversion 20 times. Transfer the entire sample to the Zymo-Spin IICR Column in a collection tube and centrifuge. Discard the flowthrough. Add 400 μL of RNA prep buffer to the column, centrifuge, discard flowthrough, and place column back into the collection tube. Add 700 μL of RNA wash buffer to the column, centrifuge, discard flowthrough, and place column back into the collection tube. Add 400 μL of RNA wash buffer to the column, centrifuge for 4 min, discard flowthrough, and place column into a new RNase-free 1.7 mL microcentrifuge tube. Add 75 μL of nuclease-free water directly to the column matrix and allow to incubate at room temperature for 1 min. Elute by centrifugation. Store mRNA on ice moving forward.
Determine RNA concentration and purity by UV spectrophotometry (e.g., NanoDrop).
Aliquot mRNA in 3 μL volumes in PCR strips and store at -80 °C. Pause point.
Confirm in vitro–transcribed mRNA quality and purity by denaturing agarose gel electrophoresis.
Cast a denaturing 0.8%–1% (w/v) agarose formaldehyde gel in OWL EasyCast B1 Mini Gel electrophoresis system with a 1.5 mm 10-well comb placed in the top comb slot. Add 0.8–1 g of agarose to 80 mL of Milli-Q water in a 500 mL Erlenmeyer flask loosely plugged with a folded-up paper towel. Microwave for 1.5 min or until the agarose is dissolved and gently swirl to mix. Cool on countertop for 2 min.
In a fume hood, add 10 mL of 10× MOPS buffer and 10 mL of 37% formaldehyde to the flask containing dissolved agarose. Gently swirl to mix and allow to cool for 5–10 min.
Pour 100 mL into the casting tray and let the gel solidify at room temperature for ~1 h in the dark by loosely covering with aluminum foil.
Prepare mRNA samples by mixing 500 ng of in vitro–transcribed mRNA, 1 μL of 1 mg/mL ethidium bromide, 1 μL of RNA loading dye (see Recipes), 5 μL of freshly-prepared RNA sample buffer (see Recipes), and nuclease-free water to 15 μL total volume.
Prepare the RNA ladder by mixing 1 μL of Millennium RNA size marker (1 μg/μL), 1 μL of 1 mg/mL ethidium bromide, 1 μL of RNA loading dye (see Recipes), 3 μL of RNA sample buffer (see Recipes), and 4 μL of nuclease-free water.
Heat ladder and samples at 70 °C for 5 min; then, place the ladder and samples on ice for 2 min.
Remove combs by gently pulling up vertically. Remove and rotate the casting tray so that the wells are near the cathode. Fill the gel tank and completely cover the agarose gel with 1× MOPS buffer (600 mL total, see Recipes).
Load ladder and samples on the gel.
Run at 60 V (constant) (or 5–10 V/cm gel width) in the dark (covered with aluminum foil) for 3–4 h (when the dye front reaches 2 cm from the bottom of the gel) or until the desired resolution.
Image gel via UV transillumination to confirm a single band is running at the expected molecular weight (polyadenylating mRNA will add to the expected molecular weight).
Note: Non-polyadenylated in vitro–transcribed mRNA will run as a single, crisp band at the expected molecular weight on a denaturing agarose gel. The same in vitro–transcribed mRNA that is polyadenylated will appear ~100–200 nt heavier as a slightly broader band. An additional control reaction lacking the Poly(A) polymerase can be included to better define the poly(A) tail length. Large smears spanning far beyond the expected molecular weight indicate RNA degradation, rolling circle transcription due to the presence of non-linear template DNA, nondenatured RNA secondary structure, or errors during RNA clean up steps. We have also found that making fresh, day-of-use RNA sample buffer is critical.
In vitro translation
Pre-cool the Sorvall Discovery M120 SE micro-ultracentrifuge and S100-AT3 rotor to 4 °C prior to setting up in vitro translation reactions. The S100-AT3 rotor can hold up to 20 samples.
Note: For each experimental mRNA, at least six samples should be prepared and translated: three replicates that are translated and not puromycin-treated and three replicates that are translated and puromycin-treated (step B3). Each of the six samples is mixed with a translated normalizing control mRNA-containing reaction (step B2), then diluted and layered on top of individual sucrose cushions (step C1).
In vitro translation of normalizing control mRNA:
Dilute normalizing mRNA to 15 fmol/μL in nuclease-free water and keep on ice. To do so, use the plasmid DNA sequence from the first transcribed nucleotide of the T7 RNA polymerase promoter to the restriction endonuclease cut site and determine the corresponding RNA sequence using the online Transcription and Translation Tool (see Software). Then, calculate the molecular weight of the control mRNA reporter with the online Oligo Calc: Oligonucleotide Properties Calculator (be sure to use the ssRNA setting; see Software). Using its molecular weight from above and the online Prism GraphPad Molarity Calculator (see Software), calculate the mass required for 3 nM in 10 μL. This is the mass of mRNA required in the complete 10 μL in vitro translation reaction for 3 nM mRNA (final). Dilute the purified mRNA in nuclease-free water such that 2 μL of RNA contains the mass calculated above, resulting in a final concentration of 15 fmol/μL. Other similar online tools are available and would suffice.
On ice in a PCR tube, set up the following 10 μL translation reaction: mix 2 μL of 15 fmol/μL in vitro–transcribed mRNA, 3 μL of Flexi RRL (nuclease-treated), 0.1 μL of 1 mM amino acid mixture minus leucine, 0.1 μL of amino acid mixture minus methionine, 0.2 μL of 25 mM Mg(OAc), 0.4 μL of 2.5 M KCl, 0.2 μL of RNase inhibitor, and 4 μL of nuclease-free water. If performing multiple reactions, prepare a master mix of all components and split into 10 μL reactions.
Perform in vitro translation for 15 min at 30 °C in a thermal cycler.
Immediately place samples on ice.
To each sample, add 4 μL of 5 mg/mL CHX (see Recipes) (final 1.43 mg/mL). Gently mix by inversion 20 times and collect contents by a short centrifugation spin. Samples should now be 14 μL. Keep samples on ice until step C1.
Note: Cycloheximide is added in step B2 to robustly inhibit elongation and preserve ribosome-bound mRNAs when added to experimental samples that contain puromycin in step B3.
In vitro translation of experimental mRNA:
Dilute experimental mRNA(s) to 15 fmol/μL (as in step B2a) in nuclease-free water and keep on ice.
On ice in a PCR tube, set up the following 10 μL translation reaction: mix 2 μL of 15 fmol/μL in vitro transcribed mRNA, 3 μL of Flexi RRL (nuclease-treated), 0.1 μL of 1 mM amino acid mixture minus leucine, 0.1 μL of amino acid mixture minus methionine, 0.2 μL of 25 mM Mg(OAc), 0.4 μL of 2.5 M KCl, 0.2 μL of RNase inhibitor, and 4 μL of nuclease-free water. If translating multiple experimental reporters or many biological replicates, prepare a master mix with all components except for the mRNA; add 8 μL of the master mix to 2 μL of 15 fmol/μL in vitro–transcribed mRNA.
Perform in vitro translation for 15 min at 30 °C in a thermal cycler.
Immediately place samples on ice.
To each sample, add 2 μL of nuclease-free water or 0.6 mM puromycin (see Recipes) (final 0.1 mM). Gently mix by inversion 20 times and collect contents by a short centrifugation spin.
Place puromycin-containing samples on a thermal cycler and incubate at 30 °C for 30 min. Keep samples without puromycin (i.e., samples with water added) on ice.
Place all samples on ice and immediately add 2 μL of 10 mg/mL CHX (see Recipes) (final 1.43 mg/mL). Gently mix by inversion 20 times and collect contents by a short centrifugation spin. Keep samples on ice.
All translation reactions should now be 14 μL.
Ribosomal pelleting through low-speed sucrose cushion
Prepare samples for low-speed ribosomal pelleting through a sucrose cushion.
Mix a 14 μL normalizing control mRNA-containing reaction tube and a 14 μL experimental mRNA-containing reaction tube for a total sample volume of 28 μL. Keep samples on ice.
Add 28 μL of ice-cold 2× ribosome dilution buffer (that contains freshly added CHX and DTT, see Recipes). Each sample should now be 56 μL.
Gently mix by inversion 20 times and collect contents by a short centrifugation spin. Keep samples on ice.
Prepare sucrose cushions and overlay samples.
Label 7 mm × 20 mm polycarbonate thick-walled tubes. Mark a single spot on the rim of each tube (Figure 2). Place tubes on a homemade ice bucket made from a P1000 tip box that fits the 7 mm × 20 mm tubes (Figure 3).
Figure 2. Example of how to mark the 7 mm × 20 mm tube to predict the side on which the pellet will form
Figure 3. Example of homemade ice bucket using a rechargeable P1000 tip box to hold 7 mm × 20 mm thick-walled tubes
Add 130 μL of ice-cold 35% buffered sucrose (that contains freshly added CHX and DTT, see Recipes) to the bottom of each tube.
Very carefully, overlay all 56 μL of the diluted sample from step C1 on top of the sucrose cushion.
Low-speed centrifugation
Without disturbing the sample–sucrose interface, place the tubes into the pre-chilled S100-AT3 rotor. Be sure to position the marked spot facing outward and toward the back of the rotor. This spot will indicate the side of the tube where the ribosome pellet will be located after centrifugation (Figure 4). Using pointed tweezers for this step is helpful.
Figure 4. Example of how to orient the 7 mm × 20 mm tube in the S100-AT3 rotor
Carefully place the rotor into the pre-cooled Sorvall Discovery M120 SE micro-ultracentrifuge.
Centrifuge samples at 50,000× g for 1 h at 4 °C. Use an acceleration setting of 9 (fastest setting) and a deceleration setting of 5 (middle setting).
Carefully, remove the tubes from the rotor and place them on ice using the homemade ice bucket for the 7 mm × 20 mm tubes. Using pointed tweezers for this step is helpful.
Using a pipettor, remove and discard the supernatant without disturbing the pellet. The glossy clear pellet should be at the bottom edge of the tube below the mark that was facing outward and toward the back of the rotor during centrifugation.
Resuspend the pellet in TRIzol.
i. First, add 200 μL of TRIzol to a new, labeled, nuclease-free microcentrifuge tube.
ii. Add 100 μL of TRIzol to the ribosome pellet in the 7 mm × 20 mm tube and mix 20 times by gently pipetting up and down. The pellet will dissociate from the tube wall when TRIzol is first added and will float in solution until it ultimately dissolves.
iii. Transfer this 100 μL to the microcentrifuge tube (now containing 300 μL).
iv. Add another 200 μL of TRIzol to the 7 mm × 20 mm tube to wash off any remaining ribosomes by gently pipetting up and down 10 times.
v. Transfer the 200 μL sample to the labeled microcentrifuge tube (now containing 500 μL total)
Mix samples end-over-end at room temperature for 15 min at 15 rpm. Collect contents by a short centrifugation spin. Pause point: Store samples at -80 °C.
RNA extractions, cDNA synthesis, and RT-qPCR
RNA extractions
Thaw samples at room temperature if necessary. Add 100 μL of chloroform to each sample.
Mix vigorously by hand for 1 min (do not vortex).
Centrifuge for 15 min at 12,000× g at 4 °C.
Without disturbing or touching the protein interface, carefully remove the top 200 μL clear aqueous layer and transfer it to a new, labeled, nuclease-free microcentrifuge tube.
Add 1.5 μL glycogen (20 mg/mL) to each sample. Upon dispensing, wash the tip in the aqueous phase by gently pipetting up and down three times.
To each sample, add 500 μL of 100% isopropanol. Gently mix by inversion 20 times.
Centrifuge for 15 min at 12,000× g at 4 °C.
Note: When pelleting RNA in microcentrifuge tubes, place the hinge upright facing the outside of the rotor. This will allow you to predict the location of the pellet at the bottom of the tube on the same side of the hinge.
Aspirate off the isopropanol until ~100 μL remains in the tube, leaving the RNA pellet untouched. Remove the final ~100 μL with a P200 pipette. The pellet should be white but very small.
Add 600 μL of ice-cold 70% ethanol. Vortex each sample for 1 s.
Centrifuge for 15 min at 12,000× g at 4 °C.
Aspirate off the ethanol until ~100 μL remains in the tube, leaving the RNA pellet untouched. Remove the final ~100 μL with a P200 pipette and finally a P10 pipette. Aspirate any remaining ethanol off the tube walls. Allow to air dry with the top open for 2–3 min at room temperature on the bench.
Place tubes on ice, add 30 μL of nuclease-free water to the pellet, and let stand on ice for 2 min. Gently resuspend the pellet by pipetting up and down 20 times. Be sure to wash down the side of the tube (same side as the hinge) to ensure complete resuspension of the RNA pellet.
cDNA synthesis
On ice, combine 16 μL of RNA and 4 μL of 5× iScript Reverse Transcription Supermix. Mix by gentle inversion 20 times and collect the contents by a short centrifugation spin. Store the remaining 14 μL of RNA at -80 °C.
Using a thermal cycler, reverse transcribe using:
i. 25 °C for 5 min (priming)
ii. 46 °C for 20 min (reverse transcription)
iii. 95 °C for 1 min (reverse transcriptase inactivation and RNA cleavage)
iv. Hold at 4 °C
To each 20 μL cDNA sample, add 180 μL of nuclease-free water for a 1:10 dilution. Gently mix by inversion 20 times and collect contents by a short centrifugation spin. Pause point: Store samples at -20 °C.
RT-qPCR
Design a 96-well plate layout scheme for all samples and negative controls. Each cDNA sample will be amplified with two primer sets: one targeting the normalizing control mRNA and the other targeting the experimental mRNA. A no-template negative control (where water is added instead of cDNA) should also be included for each primer set. Perform at least technical duplicates for each sample and primer set.
Dilute qPCR primers by mixing 510 μL of nuclease-free water, 45 μL of 10 μM forward primer, and 45 μL of 10 μM reverse primer. See General notes below for the primer sequences we used for FFLuc and nLuc.
Our typical RT-qPCR reaction is 15 μL. Each well will contain 7.5 μL of iTaq Universal SYBR Green Supermix (2×), 1.5 μL of the 1:10 diluted cDNA, and 6 μL of diluted primers. For both the normalizing control and experimental primer sets, create a master mix of iTaq Universal SYBR Green Supermix and diluted primers. Add 13.5 μL of this master mix to the appropriate wells. Then, carefully add 1.5 μL of the appropriate diluted cDNA samples. The no-template negative control contains 13.5 μL of appropriate master mix and 1.5 μL of nuclease-free water.
Seal the plate with a microseal ‘B’ seal. Be sure not to touch the top of the plastic directly, but rather apply pressure to the seal by using a clean tissue wipe (or Kimwipe). Remove and discard the perforated edges.
Centrifuge the plate in a plate centrifuge for ~30 s to pull the samples to the bottom of the wells.
Place the plate into the Bio-Rad CFX Connect Real-Time System.
Run the following RT-qPCR program:
i. One cycle of 95 °C for 3 min.
ii. Forty cycles of 95 °C for 10 s, 60 °C for 30 s, followed by a Plate Read.
iii. Melt curve from 65 °C to 95 °C with an increment of 0.5 °C for 5 s and a Plate Read.
Note: If using new primer sets, peel back the plastic film after a run and take out 10 μL to confirm the expected size amplicon on a 2% (w/v) agarose gel.
Data analysis
For each experimental mRNA, at least six samples should be prepared and translated: three replicates that are translated and not puromycin-treated and three replicates that are translated and puromycin-treated (step B3). Each of the six samples is mixed with a translated normalizing control mRNA reaction (step B2), then diluted and layered on top of individual sucrose cushions (step C1). During RT-qPCR, each cDNA should be assayed in at least technical duplicates. Primer sets for both the normalizing control mRNA and experimental mRNA should be used in separate wells on the same plate.
Using Bio-Rad CFX Maestro, select the Gene Expression window when the RT-qPCR run is complete. In the Experimental Settings tab, select the normalizing control mRNA as the Reference. This will normalize the signal of the experimental mRNA (i.e., nLuc) to the signal of the normalizing control mRNA (i.e., FFLuc) to account for any error during RNA extraction and/or cDNA synthesis. Once the gene expression has been calculated by the CFX Maestro software, export the data and perform the remaining analysis in Excel. For each experimental reporter, group the without puromycin-treatment replicates and set to 100%. Then, determine the relative signal for each replicate with puromycin-treatment. Statistical comparisons can be made using an unpaired t-test with Welch’s correction. For nLuc as the experimental mRNA reporter, we typically observe a relative ~60% reduction in signal with puromycin treatment (Scarpitti et al., 2022).
Two additional controls should be incorporated in step B3:
1) A no-template negative control, where water is added instead of either reporter mRNA, is critical to ensure specificity of RT-qPCR primers and determine background levels of detection.
2) To confirm that the low-speed sucrose cushions are working as expected to selectively pellet ribosome-bound mRNA, set up translation reactions with the experimental mRNA without puromycin, incubate for 15 min on ice instead of 30 °C (step B3c), and proceed as directed above. The ice-incubated sample should have very few, if any, ribosomes loaded on the experimental mRNA and should be minorly detected by RT-qPCR when compared to the 30 °C–incubated sample.
Validation of protocol
This protocol was validated in Scarpitti et al. (2022) Journal of Biological Chemistry, DOI: 10.1016/j.jbc.2022.102660. See Figure 6E and 6F.
General notes and troubleshooting
These in vitro translation reaction conditions have been optimized to be in the dynamic linear range for time and mRNA input for a range of reporter mRNAs (Kearse et al., 2016). These same conditions are also sufficient for A-capped Internal Ribosome Entry Site (IRES)-mediated reporter mRNAs. The final concentration of reagents in the in vitro translation reactions are 3 nM mRNA, 30% (v/v) RRL, 1 μM amino acid mixture minus leucine, 1 μM of amino acid mixture minus methionine, 0.5 mM Mg(OAc), 100 mM KCl, and 0.8 U/μL RNase inhibitor. Adjust the amount of KCl added to translation reactions if also including recombinant protein or protein synthesis inhibitors that are stored in KCl-containing buffers. The final KCl concentration should be kept constant at 100 mM. Altering the KCl concentrations will alter cap dependency. We have found that any amount of NaCl or LiCl is very inhibitory to in vitro translation using RRL.
We empirically optimized the 0.1 mM puromycin (final) for use with the in vitro translation reaction conditions described in this protocol. Increasing puromycin beyond 0.1 mM did not further reduce the amount of mRNA co-pelleted in a translation-dependent manner. We also optimized the 50,000× g sucrose cushion in the Sorvall Discovery M120 SE micro-ultracentrifuge and S100-AT3 rotor. Increased centrifugal force resulted in loss of specificity for pelleting ribosome-bound mRNAs in a translation-dependent manner. Both variables may have to be re-optimized if using different translation extracts, if making substantial alterations to the translation reaction conditions, and/or if using a different rotor.
If testing ribosome stalling by an mRNA-binding protein (Scarpitti et al., 2022), we recommend an additional control where the messenger ribonucleoprotein (mRNP) is formed but not translated (i.e., step B3c is skipped). Some mRNPs may be heavy enough to pellet alone without any bound ribosomes during the low-speed centrifugation. This control confirms that the mRNA detected in the pellet is translation- and ribosome-dependent. We have shown that FMRP forms an mRNP with nLuc mRNA and stalls ribosomes and that FMRP•nLuc mRNP alone does not pellet in the low-speed sucrose cushion unless translated. In the case of FMRP, a pre-incubation step with FMRP and mRNA was included prior to addition to the in vitro translation reaction (Scarpitti et al., 2022). However, this pre-incubation step may be unnecessary in some cases and should be optimized for each RNA-binding protein of interest.
Lastly, while we have always used FFLuc as the normalizing control mRNA and nLuc as the experimental mRNA, we believe other reporters (e.g., GFP, mCherry, Renilla Luciferase) would function just as well for either the normalizing control or experimental mRNA. We have previously cloned FFLuc from pGL4.13 into the dual promoter plasmid pCR II. The use of pCR II is not required, but pCR II/FFLuc under control of the T7 RNA polymerase promoter is available upon request. pcDNA3.1(+)/nLuc-3XFLAG, which contains a T7 RNA polymerase promoter upstream of nLuc, is available upon request and from Addgene.
See below for reporter coding sequences and qPCR primer sequences.
FireFly Luciferase (FFLuc; from pGL4.13)
ATGGAAGATGCCAAAAACATTAAGAAGGGCCCAGCGCCATTCTACCCACTCGAAGACGGGACCGCCGGCGAGCAGCTGCACAAAGCCATGAAGCGCTACGCCCTGGTGCCCGGCACCATCGCCTTTACCGACGCACATATCGAGGTGGACATTACCTACGCCGAGTACTTCGAGATGAGCGTTCGGCTGGCAGAAGCTATGAAGCGCTATGGGCTGAATACAAACCATCGGATCGTGGTGTGCAGCGAGAATAGCTTGCAGTTCTTCATGCCCGTGTTGGGTGCCCTGTTCATCGGTGTGGCTGTGGCCCCAGCTAACGACATCTACAACGAGCGCGAGCTGCTGAACAGCATGGGCATCAGCCAGCCCACCGTCGTATTCGTGAGCAAGAAAGGGCTGCAAAAGATCCTCAACGTGCAAAAGAAGCTACCGATCATACAAAAGATCATCATCATGGATAGCAAGACCGACTACCAGGGCTTCCAAAGCATGTACACCTTCGTGACTTCCCATTTGCCACCCGGCTTCAACGAGTACGACTTCGTGCCCGAGAGCTTCGACCGGGACAAAACCATCGCCCTGATCATGAACAGTAGTGGCAGTACCGGATTGCCCAAGGGCGTAGCCCTACCGCACCGCACCGCTTGTGTCCGATTCAGTCATGCCCGCGACCCCATCTTCGGCAACCAGATCATCCCCGACACCGCTATCCTCAGCGTGGTGCCATTTCACCACGGCTTCGGCATGTTCACCACGCTGGGCTACTTGATCTGCGGCTTTCGGGTCGTGCTCATGTACCGCTTCGAGGAGGAGCTATTCTTGCGCAGCTTGCAAGACTATAAGATTCAATCTGCCCTGCTGGTGCCCACACTATTTAGCTTCTTCGCTAAGAGCACTCTCATCGACAAGTACGACCTAAGCAACTTGCACGAGATCGCCAGCGGCGGGGCGCCGCTCAGCAAGGAGGTAGGTGAGGCCGTGGCCAAACGCTTCCACCTACCAGGCATCCGCCAGGGCTACGGCCTGACAGAAACAACCAGCGCCATTCTGATCACCCCCGAAGGGGACGACAAGCCTGGCGCAGTAGGCAAGGTGGTGCCCTTCTTCGAGGCTAAGGTGGTGGACTTGGACACCGGTAAGACACTGGGTGTGAACCAGCGCGGCGAGCTGTGCGTCCGTGGCCCCATGATCATGAGCGGCTACGTTAACAACCCCGAGGCTACAAACGCTCTCATCGACAAGGACGGCTGGCTGCACAGCGGCGACATCGCCTACTGGGACGAGGACGAGCACTTCTTCATCGTGGACCGGCTGAAGAGCCTGATCAAATACAAGGGCTACCAGGTAGCCCCAGCCGAACTGGAGAGCATCCTGCTGCAACACCCCAACATCTTCGACGCCGGGGTCGCCGGCCTGCCCGACGACGATGCCGGCGAGCTGCCCGCCGCAGTCGTCGTGCTGGAACACGGTAAAACCATGACCGAGAAGGAGATCGTGGACTATGTGGCCAGCCAGGTTACAACCGCCAAGAAGCTGCGCGGTGGTGTTGTGTTCGTGGACGAGGTGCCTAAAGGACTGACCGGCAAGTTGGACGCCCGCAAGATCCGCGAGATTCTCATTAAGGCCAAGAAGGGCGGCAAGATCGCCGTGTAA
nanoLuciferase (nLuc; from pNL1.1)
ATGGTCTTCACACTCGAAGATTTCGTTGGGGACTGGCGACAGACAGCCGGCTACAACCTGGACCAAGTCCTTGAACAGGGAGGTGTGTCCAGTTTGTTTCAGAATCTCGGGGTGTCCGTAACTCCGATCCAAAGGATTGTCCTGAGCGGTGAAAATGGGCTGAAGATCGACATCCATGTCATCATCCCGTATGAAGGTCTGAGCGGCGACCAAATGGGCCAGATCGAAAAAATTTTTAAGGTGGTGTACCCTGTGGATGATCATCACTTTAAGGTGATCCTGCACTATGGCACACTGGTAATCGACGGGGTTACGCCGAACATGATCGACTATTTCGGACGGCCGTATGAAGGCATCGCCGTGTTCGACGGCAAAAAGATCACTGTAACAGGGACCCTGTGGAACGGCAACAAAATTATCGACGAGCGCCTGATCAACCCCGACGGCTCCCTGCTGTTCCGAGTAACCATCAACGGAGTGACCGGCTGGCGGCTGTGCGAACGCATTCTGGCGTAA
RT-qPCR primers
F_pGL4.13 RT-qPCR: GCAGTACCGGATTGCCCAAG
R_pGL4.13 RT-qPCR: GTCGGGGATGATCTGGTTGC
F_nLuc (pNL1.1) RT-qPCR: CAGCGGGCTACAACCTGGAC
R_nLuc (pNL1.1) RT-qPCR: AGCCCATTTTCACCGCTCAG
Acknowledgments
M.R.S. was supported by The Ohio State University Fellowship and The Ohio State University Center for RNA Biology Graduate Fellowship. This work was supported by NIH grants R00GM126064 and R35GM146924 to M.G.K. This protocol was originally described and validated in Scarpitti et al. (2022). DOI: 10.1016/j.jbc.2022.102660.
References
Azzam, M. E., and Algranati, I. D. (1973). Mechanism of puromycin action: fate of ribosomes after release of nascent protein chains from polysomes. Proc Natl Acad Sci U S A 70(12 Pt 1-2): 3866-3869.
Darnell, J. C., Van Driesche, S. J., Zhang, C., Hung, K. Y., Mele, A., Fraser, C. E., Stone, E. F., Chen, C., Fak, J. J., Chi, S. W., et al. (2011). FMRP stalls ribosomal translocation on mRNAs linked to synaptic function and autism. Cell 146(2): 247-261.
Dever, T. E., Dinman, J. D. and Green, R. (2018). Translation Elongation and Recoding in Eukaryotes. Cold Spring Harb Perspect Biol 10(8).
D’Orazio, K. N., Wu, C. C., Sinha, N., Loll-Krippleber, R., Brown, G. W., and Green, R. (2019). The endonuclease Cue2 cleaves mRNAs at stalled ribosomes during No Go Decay. eLife 8: e49117.
Juszkiewicz, S., Chandrasekaran, V., Lin, Z., Kraatz, S., Ramakrishnan, V., and Hegde, R. S. (2018). ZNF598 Is a Quality Control Sensor of Collided Ribosomes. Mol Cell 72(3): 469-481.e7.
Juszkiewicz, S., Slodkowicz, G., Lin, Z., Freire-Pritchett, P., Peak-Chew, S. Y., and Hegde, R. S. (2020). Ribosome collisions trigger cis-acting feedback inhibition of translation initiation. eLife 9: e60038.
Kearse, M. G., Green, K. M., Krans, A., Rodriguez, C. M., Linsalata, A. E., Goldstrohm, A. C., and Todd, P. K. (2016). CGG Repeat-Associated Non-AUG Translation Utilizes a Cap-Dependent Scanning Mechanism of Initiation to Produce Toxic Proteins. Mol Cell 62(2): 314-322.
Kim, K. Q., and Zaher, H. S. (2022). Canary in a coal mine: collided ribosomes as sensors of cellular conditions. Trends Biochem Sci 47(1): 82-97.
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Monem, P. C., Vidyasagar, N., Piatt, A. L., Sehgal, E., and Arribere, J. A. (2023). Ubiquitination of stalled ribosomes enables mRNA decay via HBS-1 and NONU-1 in vivo. PLoS Genet 19(1): e1010577.
Scarpitti, M. R., Warrick, J. E., Yoder, E. L., and Kearse, M. G. (2022). A noncanonical RNA-binding domain of the fragile X protein, FMRP, elicits translational repression independent of mRNA G-quadruplexes. J Biol Chem 298(12): 102660.
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Sinha, N. K., Ordureau, A., Best, K., Saba, J. A., Zinshteyn, B., Sundaramoorthy, E., Fulzele, A., Garshott, D. M., Denk, T., Thoms, M., et al. (2020). EDF1 coordinates cellular responses to ribosome collisions. eLife 9: e58828.
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Multi-color Flow Cytometry Protocol to Characterize Myeloid Cells in Mouse Retina Research
WX Wei Xiao *
RS Rami A. Shahror *
CM Carol A. Morris
RC Ruth B. Caldwell
AF Abdelrahman Y. Fouda
(*contributed equally to this work)
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4745 Views: 1185
Reviewed by: Vivien J. Coulson-ThomasMunenori Ishibashi Anonymous reviewer(s)
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Abstract
Myeloid cells, specifically microglia and macrophages, are activated in retinal diseases and can improve or worsen retinopathy outcomes based on their inflammatory phenotype. However, assessing the myeloid cell response after retinal injury in mice remains challenging due to the small tissue size and the challenges of distinguishing microglia from infiltrating macrophages. In this protocol paper, we describe a flow cytometry–based protocol to assess retinal microglia/macrophage and their inflammatory phenotype after injury. The protocol is amenable to the incorporation of other markers of interest to other researchers.
Key features
• This protocol describes a flow cytometry–based method to analyze the myeloid cell response in retinopathy mouse models.
• The protocol can distinguish between microglia- and monocyte-derived macrophages.
• It can be modified to incorporate markers of interest.
We show representative results from three different retinopathy models, namely ischemia-reperfusion injury, endotoxin-induced uveitis, and oxygen-induced retinopathy.
Keywords: Retinopathy Mouse retina Flow cytometry Myeloid cells Microglia Macrophages
Background
Retinal injury is associated with activation of microglia and infiltration of bone marrow–derived macrophages and other leukocytes. This immune cell accumulation/activation is known to play a vital role in retinal injury progression and repair. While fluorescent immunolabeling coupled with confocal microscopy imaging of retina flatmounts or sections can be used to assess the retinal immune response, it is often limited by the number of antibodies that can be multiplexed, and quantification tends to be tedious and time-consuming due to the need for staining multiple sections. Furthermore, most myeloid cell markers used in imaging studies such as Iba1 and CD68 are expressed by both microglia and macrophages; also, the markers that are thought to be microglia-specific, such as P2RY12 and TMEM119, can be downregulated in activated microglia, thus complicating interpretation of the results (van Wageningen et al., 2019; Honarpisheh et al., 2020). Alternatively, flow cytometry offers a robust quantitative method to analyze immune cell response in retinal injury models. Using flow cytometry, microglia can be distinguished from myeloid leukocytes based on the relative expression of the CD45 marker coupled with the marker CD11b, where microglia are CD11b+/CD45low while myeloid leukocytes are CD11b+/CD45hi. Furthermore, flow cytometers allow for multicolor staining and hence identification of various markers of interest using the same sample. However, conducting flow cytometry on retina tissue is challenging due to the small tissue size and low cell yield. In this methods paper, we describe a step-by-step protocol to obtain high yield of viable cells and analyze different immune cell populations after retina injury. A panel of antibodies is included to identify the immune cell subsets and myeloid cell inflammatory phenotype using proinflammatory (M1-like) marker, CD11c, and the anti-inflammatory (M2-like) marker, CD206. We present representative flow cytometry results from three different retina injury models—retinal ischemia-reperfusion (IR) injury, endotoxin-induced uveitis (EIU), and oxygen-induced retinopathy (OIR)—that model ischemic retinopathy, uveitis, and retinopathy of prematurity, respectively. The IR and EIU are induced in adult mice and represent two different injury modalities, with the latter leading to a stronger immune response. The OIR model assesses the murine pup’s retina response to hypoxia, which involves vascular regression followed by neovascularization.
Materials and reagents
Ketamine 100 mg/mL (Hikma, NDC 0143-9509-01); note that ketamine is a controlled substance and needs a special Drug Enforcement Administration license to be obtained
Xylazine 100 mg/mL (Covertus, catalog number: 1XYL006)
Sodium chloride for injection vial USP (0.9% saline, 50 mL) (Hospira, NDC 0409-4888-06)
Sodium chloride for injection USP IV bag (0.9% saline, 250 mL) (Baxter, NDC 0338-0049-02)
Bovine serum albumin (BSA) (GeminiBio, catalog number: 700-100P)
DNase I (Sigma-Aldrich, catalog number: NC1539905)
Ethylenediaminetetraacetic acid (EDTA), 0.5 M (Thermo Fisher, catalog number: AM9260G)
Dulbecco’s modified Eagle medium (DMEM), high glucose, GlutaMAXTM supplement, pyruvate (Thermo Fisher, catalog number: 10569010)
Fetal bovine serum (FBS) (Thermo Fisher, catalog number: 10438026)
Phosphate buffered saline (PBS, 10×) (Thermo Fisher, catalog number: 70-011-044)
Hank’s balanced salt solution (HBSS) with calcium and magnesium (Thermo Fisher, catalog number: 14025076)
HEPES buffer solution (Millipore Sigma, catalog number 83264)
LiberaseTM (Sigma, catalog number: NC1179175)
Zombie VioletTM Fixable Viability kit (BioLegend, catalog number: 423113)
Jackson Immuno Research Labs normal rat serum (Fisher, catalog number: NC9834724)
RBC lysis buffer for mouse (Thermo Fisher, catalog number: J62150.AK)
FisherbrandTM round-bottom polystyrene test tubes without cap, FACS tubes (Thermo Fisher, catalog number: FB149563A)
Fluorescent conjugated primary antibodies and unconjugated blocking antibodies (CD16/32) (Table 1)
Table 1. Antibody panel used in this protocol
Antibody Conc. Fluorophore Clone Host Company Catalog # Dilution
CD11b 0.2 mg/mL PerCP M1/70 Rat BioLegend 101230 1:100
CD11c 0.2 mg/mL Brilliant Violet 605TM N418 Armenian Hamster BioLegend 117334 1:100
CD206 0.5 mg/mL FITC C068C2 Rat BioLegend 141704 1:100
CD45 0.5 mg/mL Alexa Fluor® 700 30-F11 Rat BioLegend 103127 1:100
F4/80 0.5 mg/mL PE C1:A3-1 Rat Cedarlane CL8940PE 1:100
Gr-1 (Ly-6G/
Ly-6C)
0.2 mg/mL APC RB6-8C5 Rat BD Bioscience 561083 1:100
CD16/CD32 0.5 mg/mL Used as (FC block) 2.4G2 Rat BD Bioscience 553142 1:100
Solutions
Anesthesia cocktail (see Recipes)
Digestion buffer (see Recipes)
Flow cytometry staining buffer (FACS/EDTA buffer) (see Recipes)
Recipes
Anesthesia cocktail
Mix 2 mL of ketamine (100 mg/mL), 0.2 mL of xylazine (100 mg/mL), and 1.8 mL of 0.9% saline to achieve a final solution of 50 mg ketamine and 5 mg xylazine per 1 mL.
Digestion buffer
Supplement HBSS with 5% FBS prepared from frozen stock (triple filtered by the manufacturer through a 0.1 μm filter) and 10 mM HEPES, 0.5 mg/mL of liberase, and 0.1 mg/mL of DNase I. Mix gently for 5 min at 4 °C and keep protected from light until needed. Make sure that the FBS used in the digestion buffer is filtered. If not, filter the entire digestion buffer through a 0.22 μm filter to avoid any cellular aggregation that may occur due to potential debris in the unfiltered FBS.
Flow cytometry staining buffer (FACS/EDTA buffer)
Prepare by adding 1 mL of 0.5 M EDTA to 50 mL of 5% BSA in 1× PBS.
Equipment
BD LSRFortessa flow cytometer; analyzes up to 16 colors (BD Biosciences)
Digital Peri-StarTM Pro peristaltic perfusion pump (World Precision Instruments)
Eppendorf benchtop centrifuges (for 2, 15, and 50 mL tubes at room temperature and 4 °C)
Lab ArmorTM 37 °C bead bath (Fisher)
Invitrogen Countess III cell counter and counting chamber slides (Thermo Fisher)
Bench-top vortex (Fisher)
Dissection tools (World Precision Instruments, catalog number: MOUSEKIT)
General lab supplies: tubes, pipettes, tips, etc.
Insulin syringes (U-30, short needle, 30 gauge) for ketamine/xylazine administration (MHC medical)
Surgical blades, No. 10 (Medline, catalog number: MDS15010)
Weighing balance (Jscale, catalog number: CJ-4000)
30 G needle (TURMO, catalog number: NN3025R)
25 G needle (Air-Titn, catalog number: 830003786)
Hemostat (World Precision Instruments, catalog number: 15920)
Styrofoam or wood surgical station
Software
FlowJo (software package for analyzing flow cytometry data), https://www.flowjo.com/
Procedure
Ethics Statement: all procedures were performed in accordance with the Association for Research in Vision and Ophthalmology (ARVO) and University of Arkansas for Medical Sciences (UAMS) Institutional Animal Care and Use Committee guidelines. A protocol flowchart is provided in Figure 1.
Figure 1. Protocol flowchart
Euthanasia and retina harvesting
Anesthesia: determine the mouse’s body weight to the nearest gram and inject intraperitoneally 40–50 μL of the anesthesia cocktail for a 25 g mouse, for a dose of 80–100 mg/kg ketamine and 5–10 mg/kg xylazine. Wait a few minutes until the mouse is completely anesthetized as measured by loss of response to toe pinch.
Transcardial perfusion (Figure 2):
Mount the mouse in supine position on Styrofoam or wood surgical station using tape or pins on its four paws.
Expose the xiphoid by making a skin incision on the chest; then, make lateral incisions beneath the ribcage using tissue scissors. Carefully cut through both sides of the rib cage up to the collarbone using tissue scissors.
Pin the sternum up over the head of the mouse with a hemostat. Clear the pericardial sac and any other tissues/organs covering the heart using dissecting forceps to provide a clear view of the heart and vessels.
Make a small incision in the right atrium using iris scissors to create a perfusion outlet. Perform transcardial perfusion using a 25 G needle inserted into the left ventricle with 1× PBS at a constant speed of ~1.5 mL/min using a perfusion pump until the fluid exiting the right atrium is entirely clear. The rate of 1.5 mL/min depends on the pump speed and tube size. It can be adjusted by timing the drip into a 2 mL tube before the experiment and adjusting the pump speed accordingly. It is important that the heart remains beating during the perfusion process to achieve complete perfusion. Inject each mouse with the correct dose of anesthesia cocktail right before perfusion, as giving a high dose or waiting a long time after the cocktail injection may cause the heart to stop and perfusion to fail.
The perfusion step is essential to clear retina vessels from any residual blood, to limit analysis to retinal resident and infiltrating immune cells and exclude cells in the blood circulation.
Figure 2. Transcardial perfusion. 1. Mouse mounted in supine position with arrows indicating the incision line to open the chest cavity. 2. The heart is exposed by making lateral incisions on both sides of the ribcage using tissue scissors. 3. Heart is exposed after clearing the pericardium with the arrow pointing to the apex. It is important that the heart remains beating during the perfusion process to achieve complete perfusion. 4. Insert a 25 G needle connected to the perfusion pump tubing into the left ventricle through the apex of the heart. 5. Make a small incision in the right atrium using iris scissors to create a perfusion outlet. 6. Turn on the perfusion pump to perform transcardial perfusion with 1× PBS at a constant speed of ~1.5 mL/min, until the fluid exiting the right atrium is entirely clear and the liver (denoted by asterisk) becomes pale.
Retina tissue collection (Figure 3):
Put the mouse in prone position.
Place Dumont #7 curved forceps around the rear part of the eyeball close to the optic nerve and apply gentle pressure to protrude the eye.
Make a wide cut in the cornea along the equator using a sharp blade while holding the eyeball with forceps from beneath.
Gently remove the lens and vitreous body with the tip of the blade; then, extract the retina from the eyecup by pulling the forceps upwards.
Note that the retina is an off-white translucent tissue. Make sure that the retina is detached from the retinal pigment epithelium by removing any black tissue that comes out with the isolated retina.
Place the retina from both eyes in 2–3 mL of cold HBSS in a 10 mm Petri dish.
Figure 3. Retina tissue collection. 1. Put the mouse in prone position and hold the eyeball from beneath to cause the eye to protrude. 2. Make a wide cut in the cornea along the equator using a sharp blade while holding the eyeball with the forceps from beneath. 3. Grip the dissected eyeball with forceps to remove the lens and vitreous denoted by arrow. 4. Squeeze the remaining eye cup to extract the retina from the eyeball. 5. Move the retina (off-white translucent tissue) to 2–3 mL of cold HBSS. Asterisk denotes the discarded lens with the vitreous and iris. 6. Remove any remaining retinal pigment epithelium (black tissue, denoted by asterisk). 7. Retinas from both eyes in HBSS are ready for further processing.
Retina digestion and single-cell suspension (Figure 4)
Figure 4. Representative images of retina digestion protocol. 1. Diced retinas using a scalpel. 2. Retinas resuspend in digestion buffer. 3. Digested retinas after incubation for 30 min at 37 °C. 4. Digested retinas after adding 10 mL of DMEM + 10% FBS. 5. Straining the suspension through a mesh. 6. Using the rubber end of a syringe plunger to gently press the samples through the mesh. 7. Cell pellet after centrifugation. 8. Cell suspension in a 5 mL FACS tube. 9. Cell pellet after spinning down. 10. Remaining pellet in 100 μL of PBS after aspiration. 11. Resuspended cells ready for staining.
Use a scalpel to dice the retinas into pieces < 1 mm in HBSS; then, transfer to a 15 mL conical tube and centrifuge at 400× g for 5 min at room temperature.
Resuspend the pelleted tissue in a total of 500 μL of digestion buffer.
Note: You can add an additional step to eliminate any red blood cells (RBC) in the samples by incubating the digested tissue in 1 mL of RBC lysis buffer for 5 min at room temperature.
Place the tubes on a rack and place the rack into a 37 °C water (or dry beads) bath for 30 min. Vortex gently every 10 min.
Add 10 mL of DMEM containing 10% FBS to the reaction to inhibit the enzymatic digestion and increase the cell yield.
Gently strain each sample three times through a 40 μm cell strainer and wash the strainer with 1 mL of PBS after each strain. Use the rubber end of a syringe plunger to gently press the samples through the mesh.
Spin down at 400× g for 5 min at 4 °C.
Resuspend the pelleted tissues in 3 mL of PBS and count the cells. One can start by pooling two to four whole retinas per sample and reduce the number in subsequent experiments based on the cell yield.
Cells can be counted on an automated cell counter or manually using a hemocytometer under a microscope.
Note: Cell counts are approximately 1.5 × 106 cells per three retinas, which is sufficient for each staining panel/sample. Consider pooling more retinas for fluorescence minus one (FMO) and control samples.
Divide the cell samples into the 5 mL FACS test tubes with proper cell number and groups. At least 1.5 million cells per tube are required for further processing. The tube groups should include the following controls along with sample groups: A) negative/unstained cells; B) single stain controls for each color; and C) FMOs for each color. For our antibody panel, we used the following groups: unstained cells, cells + PerCP only, cells + APC only, cells + PE only, cells + FITC only, cells + AF700 only, cells + BV605 only, cells + viability dye only, FITC FMO- cells with all stains except FITC, PE FMO- cells with all stains except PE, APC FMO- cells with all stains except APC, PerCP FMO- cells with all stains except PerCP, AF700 FMO- cells with all stains except AF700, BV605 FMO- cells with all stains except BV605, and viability dye FMO- cells with all stains except viability dye.
Note: Controls are needed to set instrument voltages and compensate the samples, while the FMOs are used to set the positive and negative gates for each color (fluorophore) in the analysis. Controls can be done the first time the samples are run and omitted for subsequent experiments if the same conditions and protocols for tissue lysis and staining are followed.
Spin down at 400× g for 5 min at 4 °C.
Carefully aspirate the supernatant in each tube without disturbing the pellet until approximately 100 μL are remaining. Next, add another 100 μL of PBS and resuspend the pellet immediately to avoid clumping.
Cells are then ready to be stained for flow cytometry. Cells should be protected from light throughout staining and storage.
Staining for flow cytometry
Stain for viability (live/dead stain) by diluting the fixable Zombie Violet UVTM dye at 1:100–1:1,000 in PBS and resuspend 1.0 × 106 cells in diluted 100 μL of Zombie UVTM solution. Incubate the samples for 30 min on ice in the dark. To minimize background staining of live cells, titrate the volume of dye and/or number of cells per 100 μL for optimal performance.
Note: Titrate the dye volume that will be used in preliminary studies to minimize background staining of live cells.
Wash by adding 2–3 mL of FACS/EDTA buffer and spin down at 400× g for 5 min at 4 °C. Gently aspirate supernatant without disrupting the cell pellet.
Block with 1 μg/mL of Fc block anti-mouse CD16/32 and 20% normal rat serum at a volume of 50 mL of PBS/sample for 10 min at room temperature or 20 min at 4 °C.
Without washing, add the appropriate volume of antibodies for surface staining to evaluate the different cell populations. For this protocol, we used the antibody panel listed in Table 1.
Protect samples from light and incubate them as per the antibody manufacturer’s instructions, which is typically 20 min at 4 °C.
Add 1 mL of FACS/EDTA buffer and spin down at 400× g for 5 min at 4 °C.
Gently aspirate the supernatant.
Resuspend the cell pellet in 300 μL of FACS/EDTA buffer and cover the samples with foil for same-day run on the flow cytometer.
Alternatively, samples can be fixed by resuspending in 0.4% PFA and kept overnight at 4 °C and protected from light for next-day run on the flow cytometer. However, we recommend running the sample immediately after staining to avoid any loss of yield or signal.
Data analysis
Data analysis, gating strategy, and identification of immune cells subsets
Selecting cells and excluding debris
Use Forward versus Side Scatter (FSC vs. SSC) density plot (Figure 5A) to identify the cell population and exclude debris. FSC indicates cell size, while SSC indicates cell complexity or granularity. Cell debris is excluded by creating a gate that excludes the bottom left corner (debris).
Figure 5. Gating strategy for retinal immune cells. Gating strategy is shown in various panels, with panels E and F showing the cell population of interest as a percentage of viable cells in panel D, while panels G–J show the cell populations as a percentage of the gated population in the preceding panel.
Selecting singlets and excluding doublets
Use Side Scatter Height versus Side Scatter Area (SSC-H vs. SSC-A) density plot to exclude doublets. A Forward Scatter Height versus Forward Scatter Area (FSC-H vs. FSC-A) plot can also be used (Figure 5B and 5C).
Selecting viable cells
This is achieved by using a viability dye vs. FSC-H density plot and excluding cells stained with the dye (Figure 5D).
Note: Viability dyes are impermeable to live cells and therefore stain only dead cells.
Identifying populations with specific markers
The general leukocyte marker CD45 is used to identify immune cell populations (Figure 5E). Gating on CD45 and CD11b (myeloid cell marker) is used to distinguish microglia (CD11b+ CD45low), myeloid leukocytes (CD11b+ CD45hi), and lymphocytes (CD11bneg CD45hi), as shown in Figure 5F. T-helper cells can be identified as CD4+ (Figure 5G). Microglia (CD11b+ CD45low) and macrophages (CD45hi CD11b+ F4/80+) can be further characterized as M1-like CD11c+ CD206- and M2-like CD11c- CD206+ cells (Figures 5H, 5I). Monocytes and granulocytes were characterized as CD11b+ Gr-1int. and CD11b+ Gr-1hi, respectively (Figure 5J).
Note: Another approach is to distinguish classical pro-inflammatory monocytes (CD11b+ CD45hi Ly6Chi Ly6Gneg) from non-classical anti-inflammatory monocytes or monocyte-derived macrophages (CD11b+ CD45hi Ly6Cneg Ly6Gneg) using antibodies specific for Ly6C and Ly6G, as described in Abcouwer et al. (2021).
Validation of protocol
Representative results from different injury models are presented in Figure 6.
Figure 6. Representative results from three different retinal injury models. Panels from control (A), ischemia-reperfusion (IR) injury (B), oxygen-induced retinopathy (OIR) (C), and endotoxin-induced uveitis (EIU) (D) show density plots of CD11b vs. CD45 to distinguish microglia from myeloid cells and lymphocytes (i), CD45hi leukocytes (ii), and F4/80+ CD11b+ macrophages as a percentage of CD45hi cells (iii). A strong leukocyte and macrophage infiltration was observed in both the IR and EIU models, with a stronger response in the EIU model. On the other hand, the OIR model was associated with more microglial (CD11b+ CD45low) proliferation.
Injury models
Retinal IR injury model was achieved under anesthesia by raising the eye intraocular pressure (IOP) to 110 mm Hg for 60 min unilaterally. The anterior chamber of one eye is cannulated with a 30 G needle attached to a line infusing sterile saline. The IOP is raised to 110 mm Hg to achieve ischemia by elevating the saline reservoir. After 60 min, the needle is withdrawn to allow reperfusion and initiate IR injury.
EIU was induced by intraperitoneal injection of lipopolysaccharide from Salmonella typhimurium (LPS, 4 mg/kg in PBS, Sigma-Aldrich).
OIR model was induced by subjecting 1-week-old mice litters along with their nursing dams to hyperoxia of 75% oxygen in sealed chamber, starting at postnatal day 7 (P7) for five days until P12, and then followed by normoxia or room air. Here in this protocol, we subjected the pups to two days of normoxia (P12–P14).
We used 8–10-week-old male C57BL6J mice for the control, IR-injury, and EIU data, and P14 pups for the OIR model. The IR model was induced as described in Fouda et al. (2018), the EIU was induced as described in Zhang et al. (2009), and OIR was induced as described in Fouda et al. (2022).
Acknowledgments
This work was supported by the following grants from the National Institute of Health (NIH): R01-EY11766 to RBC, and R00 EY029373-03 to AYF. Our protocol is modified/adapted from previously published work by Abcouwer et al. (2021) and O’Koren et al. (2016).
Competing interests
The authors declare no competing interests.
Ethical considerations
All studies were approved by the Augusta University and University of Arkansas for Medical Sciences IACUC committee.
References
Abcouwer, S. F., Shanmugam, S., Muthusamy, A., Lin, C. M., Kong, D., Hager, H., Liu, X. and Antonetti, D. A. (2021). Inflammatory resolution and vascular barrier restoration after retinal ischemia reperfusion injury.J Neuroinflammation 18(1): 186.
Fouda, A. Y., Xu, Z., Suwanpradid, J., Rojas, M., Shosha, E., Lemtalsi, T., Patel, C., Xing, J., Zaidi, S. A., Zhi, W., et al. (2022). Targeting proliferative retinopathy: Arginase 1 limits vitreoretinal neovascularization and promotes angiogenic repair. Cell Death Dis 13(8): 745.
Fouda, A. Y., Xu, Z., Shosha, E., Lemtalsi, T., Chen, J., Toque, H. A., Tritz, R., Cui, X., Stansfield, B. K., Huo, Y., et al. (2018). Arginase 1 promotes retinal neurovascular protection from ischemia through suppression of macrophage inflammatory responses. Cell Death Dis 9(10): 1001.
Honarpisheh, P., Lee, J., Banerjee, A., Blasco-Conesa, M. P., Honarpisheh, P., d’Aigle, J., Mamun, A. A., Ritzel, R. M., et al. (2020). Potential caveats of putative microglia-specific markers for assessment of age-related cerebrovascular neuroinflammation. J Neuroinflammation 17(1): 366.
O'Koren EG, Mathew R, Saban DR. Fate mapping reveals that microglia and recruited monocyte-derived macrophages are definitively distinguishable by phenotype in the retina. Sci Rep . 2016;6:20636.
Van Wageningen, T. A., Vlaar, E., Kooij, G., Jongenelen, C. A. M., Geurts, J. J. G. and van Dam, A. M. (2019). Regulation of microglial TMEM119 and P2RY12 immunoreactivity in multiple sclerosis white and grey matter lesions is dependent on their inflammatory environment.Acta Neuropathol Commun 7(1): 206.
Zhang, W., Baban, B., Rojas, M., Tofigh, S., Virmani, S. K., Patel, C., Behzadian, M. A., Romero, M. J., Caldwell, R. W. and Caldwell, R. B. (2009). Arginase activity mediates retinal inflammation in endotoxin-induced uveitis.Am J Pathol 175(2): 891-902.
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Immunology > Immune cell staining > Flow cytometry
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Peer-reviewed
Analysis of Pectin-derived Monosaccharides from Arabidopsis Using GC–MS
PS Patricia Scholz
KC Kent D. Chapman
TI Till Ischebeck
AG Athanas Guzha
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4746 Views: 510
Reviewed by: Samik BhattacharyaCătălin VoiniciucKumiko Okazaki
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Original Research Article:
The authors used this protocol in Plant Physiology Jul 2022
Abstract
Pectin is a complex polysaccharide present in the plant cell wall, whose composition is constantly remodelled to adapt to environmental or developmental changes. Mutants with altered pectin composition have been reported to exhibit altered stress or pathogen resistance. Understanding the link between mutant phenotypes and their pectin composition requires robust analytical methods to detect changes in the relative monosaccharide composition. Here, we describe a quick and efficient gas chromatography–mass spectrometry (GC–MS)-based method that allows the differential analysis of pectin monosaccharide composition in plants under different conditions or between mutant plants and their respective wild types. Pectin is extracted from seed mucilage or from the alcohol-insoluble residue prepared from leaves or other organs and is subsequently hydrolysed with trifluoracetic acid. The resulting acidic and neutral monosaccharides are then derivatised and measured simultaneously by GC–MS.
Key features
• Comparative analysis of monosaccharide content in Arabidopsis-derived pectin between different genotypes or different treatments.
• Procedures for two sources of pectin are shown: seed coat mucilage and alcohol-insoluble residue.
• Allows quick analyses of neutral and acidic monosaccharides simultaneously.
Graphical overview
Keywords: Cell wall Pectin Gas chromatography–mass spectrometry Monosaccharide Arabidopsis
Background
Primary cell walls consist of up to 90% polysaccharides including cellulose, hemicelluloses, and pectin (Pettolino et al., 2012; Höfte and Voxeur, 2017). Of those, pectin is characterised by high amounts of D-galacturonic acid, which forms the backbone in the pectic polysaccharides homogalacturonan and rhamnogalacturonan II. In a third pectic polysaccharide, rhamnogalacturonan I, the backbone consists of disaccharide units of D-galacturonic acid linked to L-rhamnose (Atmodjo et al., 2013). Further mono- or oligosaccharide side chains are linked to the backbone, resulting in a chemically complex structure (Harholt et al., 2010; Atmodjo et al., 2013).
Pectin is the most abundant component of primary cell walls of many plant species and performs diverse functional roles, including the response to infection by various pathogens (Shin et al., 2021). In this context, it was reported that an Arabidopsis double mutant line, disrupted in the two galacturonic acid–producing enzymes GLUCURONATE 4-EPIMERASE 1 and 6, contained less homogalacturonan and concomitantly showed higher susceptibility towards infection with the pathogens Pseudomonas syringae pv maculicola ES4326 and Botrytis cinerea isolates (Bethke et al., 2016). Also, modifications of the pectin backbone have been connected to plant–pathogen interactions. In Arabidopsis, POWDERY MILDEW RESISTANT5 (PMR5) is responsible for the acetylation of galacturonic acid, and the pmr5 mutant has been shown to possess enhanced resistance to powdery mildew (Vogel et al., 2004). Furthermore, the bxl4 Arabidopsis mutant that lost enzyme activity of BETA-XYLOSIDOSE4 (BXL4) exhibited increased susceptibility to the fungal pathogen B. cinerea. BXL4 has been shown to possess activities on cell wall–derived arabinans and xyloses (Guzha et al., 2022).
Given the reported effects of changes in pectin abundance and composition, fast and straightforward methods to compare the relative composition of pectin in plants under different conditions or of different genotypes are essential. Different methods for compositional analysis have been described so far, including colorimetric assays, gas chromatography–mass spectrometry (GC–MS), and high-pressure liquid chromatography-based approaches (Pettolino et al., 2012; Biswal et al., 2017; Bethke and Glazebrook, 2019; Dean et al., 2019). Among those, GC–MS-based methods require analytes volatile enough for GC separation. Two main methods to obtain volatile analytes are described in the literature: reduction and acetylation to alditol acetates or acidic methanolysis of the sample followed by trimethylsilylation. The latter procedure offers the advantage of analysing neutral and acidic monosaccharides simultaneously; however, methanolysis is a time-consuming process with incubation times of 18 h (Biswal et al., 2017). We therefore intended to combine shorter sample processing times with the simultaneous study of neutral and acidic monosaccharides.
We describe a GC–MS-based analysis of cell wall pectin extracted from cell wall samples as alcohol-insoluble residue (AIR) or seed coat mucilage by water extraction. Subsequently, extracted pectin is hydrolysed with trifluoracetic acid (TFA) that can be evaporated to leave the monosaccharide monomers. These are then derivatised with O-methylhydroxylamine hydrochloride (MOX) and N-methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA) to yield volatile analytes to be analysed by GC–MS.
Materials and reagents
Reagents
Ultrapure water (prepared, for example, with a water purification system)
Liquid nitrogen
Nitrogen (N2) gas
Acetone (Carl Roth, catalog number: 9372.5)
Chloroform (Carl Roth, catalog number: 7331.1)
Undenatured ethanol
Methanol (Fisher Scientific, catalog number: 10124490)
Pyridine anhydrous (Sigma-Aldrich, catalog number: 270970)
Trifluoroacetic acid (TFA) (Sigma-Aldrich, catalog number: 302031)
O-Methylhydroxylamine hydrochloride (MOX) (Sigma-Aldrich, catalog number: 226904)
N-Methyl-N-trimethylsilyltrifluoroacetamide (MSTFA) (Sigma-Aldrich, catalog number: 69479)
allo-Inositol (Sigma-Aldrich, catalog number: 468088)
L-Arabinose (Sigma-Aldrich, catalog number: A91906)
L-Fucose (Sigma-Aldrich, catalog number: F2252)
D-Galactose (Carl Roth, catalog number: 4987.2)
D-Galacturonic acid monohydrate (Sigma-Aldrich, catalog number: 48280)
D-Glucose (Carl Roth, catalog number: 6780.2)
D-Glucuronic acid sodium salt monohydrate (Sigma-Aldrich, catalog number: G8645)
D-Mannose (Sigma-Aldrich, catalog number: M8574)
L-Rhamnose monohydrate (Carl Roth, catalog number: 4655.2)
D-Xylose (Sigma-Aldrich, catalog number: X3877)
Solutions
70% ethanol (see Recipes)
2 M trifluoroacetic acid (TFA) (see Recipes)
allo-Inositol (see Recipes)
Monosaccharide standards (see Recipes)
O-Methylhydroxylamine hydrochloride (MOX) derivatisation solution (see Recipes)
Recipes
70% ethanol
Reagent Final concentration Quantity
Ethanol (absolute) 70% 70 mL
H2O n/a 30 mL
Total n/a 100 mL
2 M trifluoracetic acid (TFA)
Reagent Final concentration Quantity
TFA (13 M) 2 M 1.54 mL
H2O n/a 8.46 mL
Total n/a 10 mL
Add the acid slowly to ultrapure water. 10 mL of 2 M TFA solution is enough for the hydrolysis of 30 samples. Caution: TFA is corrosive and attacks skin, eyes, and mucous membranes. Wear chemical-resistant gloves and safety goggles and work with TFA only under a fume hood.
allo-Inositol
Reagent Final concentration Quantity
allo-Inositol 50 μg/mL 5 mg
H2O n/a 100 mL
Total n/a 100 mL
Monosaccharide standards
Reagent Final concentration Quantity
L-Arabinose 250 mM 37.53 mg in 1 mL of water
D-Xylose 250 mM 37.53 mg in 1 mL of water
L-Fucose 250 mM 41.04 mg in 1 mL of water
L-Rhamnose 250 mM 45.54 mg in 1 mL of water
D-Galactose 250 mM 45.04 mg in 1 mL of water
D-Glucose 250 mM 45.04 mg in 1 mL of water
D-Mannose 250 mM 45.04 mg in 1 mL of water
D-Galacturonic acid 250 mM 53.04 mg in 1 mL of water
D-Glucuronic acid 250 mM 58.54 mg in 1 mL of water
Note down the actual mass that you weighed in and adjust the volume using the following formula:
O-Methylhydroxylamine hydrochloride (MOX) derivatisation solution (see General note 3)
Reagent Final concentration Quantity
MOX 30 mg/mL 15 mg
Pyridine anhydrous n/a 0.5 mL
Total n/a 0.5 mL
Caution: Pyridine is harmful if swallowed, in contact with skin, or if inhaled. Wear chemical-resistant gloves and work with pyridine and MOX solutions only under a fume hood. Use solvent-resistant tips for pipetting.
Laboratory supplies
10 cm plant pots
Soil: Einheitserde SPECIAL Vermehrung, Patzer Erden, Sinntal-Altengronau, Germany; medium clay content, contains peat, perlite, 1% nutrient salts, trace elements and iron, pH 5.8
Chemical-resistant gloves
Safety goggles
Solvent-resistant pipette tips [Biozym, catalog numbers: 693010 (10 μL), 692069 (200 μL), 692078 (1,000 μL)]
Vials, inserts, and lids for GC–MS [Macherey-Nagel, catalog numbers: 702282 (vials), 702716 (inserts), 702287.1 (lids)]
Glass beads, 5 mm diameter (Carl Roth, catalog number: HH56.1)
2 mL microcentrifuge tube (Sarstedt, catalog number: 72.695.500)
1.5 mL microcentrifuge tube (Sarstedt, catalog number: 72.690.001)
DURAN® culture tubes with screw cap (VWR, catalog number: 391-4022)
Equipment
Growth chamber with light and temperature controls
Oven for sterilization of soil at 80 °C
Fume hood
Gas chromatography–mass spectrometry (GC–MS) system:
Agilent Technologies 7890B GC-System coupled to 5977B MSD quadrupole
PAL3 Auto sampler system with Robotic Tool Change (RTC)
HP-5ms Ultra Inert column (Agilent, catalog number: 19091S-433UI)
Nitrogen evaporator/sample concentrator
Analytical balance (Kern, model: 770-15)
Rotary shaker
Microcentrifuge
Heating block with shaker for 2 mL microcentrifuge tubes (HLC HTM 130, Haep Labor Consult)
Heating block for DURAN glass tubes (LiebischTM Monoblock)
Ball mill (Retsch MM400)
Optional: Mortar and pestle
Vortex mixer (Vortex-Genie 2)
Micropipettes
Acid resistant micropipette
Laboratory spatulas
Software
GC/MSD MassHunter with MSD ChemStation Data Analysis (G1701FA F.01.03.2357; Agilent Technologies)
Microsoft Excel 2019 (Microsoft)
Procedure
Plant growth
For individual experimental setup, we recommend plants to be grown according to your usual procedure. Pots with the various genotypes (3–4 pots, each with five plants per genotype) should be randomised, placed in the same tray, and grown together under the same conditions. As an example, the procedure to enable comparative pectin analysis on alcohol-insoluble residue (AIR) derived from leaves, as used in Guzha et al. (2022), is shown below.
Prepare semi-sterile soil by heating it in the oven for 8 h at 80 °C.
Transfer soil to 10 cm pots, water, and place 6–10 Arabidopsis seeds in the four corners and the centre of the pots. Cover with plastic dome.
After stratification at 4 °C for two days, transfer to growth cabinet with short-day conditions (8 h light and 16 h darkness) at 22 °C, a relative humidity of 65%, and a light intensity of 120–140 μmol m-2·s-1. Remove the plastic dome after 3–4 days.
Grow plants for 7–10 days, then thin out excess seedlings to leave five plants per pot (one plant per position).
Grow until plants reach the desired growth stage depending on your experimental setup, e.g., 6-week-old plants for analysis of leaf AIR.
A detailed description on plant growth and seed harvest for mucilage analysis is given in Dean et al. (2019) (see General note 1).
Preparation of alcohol insoluble residue (AIR)
The method used in this part of the protocol is adapted from Bethke et al. (2016) and involves the following steps:
Flash-freeze the organs of interest (leaves, stems, roots, whole rosettes) in liquid nitrogen. Soil debris attached to roots or stems can be gently washed off using sterile distilled Milli-Q water.
Note: To minimise starch contamination, leaf samples can be harvested after keeping the plants in the dark for 48 h, if the experimental setup allows (see General note 2).
Grind the frozen plant material using mortar and pestle or a ball mill. Keep the plant material frozen throughout grinding and make sure that the plant material is ground as fine as possible to minimize non–cell wall contamination. Pause point: The ground plant material can be stored at -80 °C.
Transfer up to 100 mg of ground plant material in a pre-cooled 2 mL microcentrifuge tube. Keep samples cooled before the addition of ethanol in the next step.
Add 1.5 mL of 70% (v/v) ethanol and vortex thoroughly.
Centrifuge at 18,000× g for 10 min at room temperature and remove supernatant.
Repeat steps B4 and B5 as described above.
Add 1 mL of the solvent mixture chloroform/methanol 1:1 (v/v) and vortex thoroughly. Caution: Work with chloroform-containing solutions under a fume hood.
Centrifuge at 18,000× g for 10 min at room temperature and remove supernatant. Caution: Removal of the chloroform-containing solution should be carried out under a fume hood.
Repeat steps B7 and B8 two times as described above. After the final washing step, the plant material should have a grey to off-white colour, as all chlorophyll-containing fractions have been removed. Carry out additional washes with chloroform/methanol 1:1 (v/v) if the plant material is still green.
Add 1 mL of acetone to the plant material and vortex thoroughly.
Centrifuge at 18,000× g for 10 min at room temperature and remove supernatant.
Air dry the resulting AIR at room temperature overnight. Keep samples in a dust-free environment to avoid contamination while drying.
To avoid starch contamination, it is possible to carry out an α-amylase digest (see General note 2).
Extraction and hydrolysis of pectin
Pectin analysis from AIR
Prepare at least four replicates for each genotype or growth condition. Weigh 2 mg of AIR for each replicate and place in a 2 mL microcentrifuge tube. Record the exact weight of AIR placed in the microcentrifuge tube. Use AIR from plants grown at the same time and under the same conditions.
Add 1.4 mL of water to the AIR.
Add two glass beads (5 mm diameter) and homogenise AIR further with a ball mill for 1.5 min at 30 Hz.
Incubate samples for 2 h at 90 °C under constant vigorous shaking (speed #4 on HLC HTM 130).
Remove samples from heating block and allow the debris to settle for 5 min. Do not centrifuge the samples.
Transfer 1 mL of the supernatant to a DURAN® glass culture tube with screw cap.
Dry the samples under a stream of N2 gas at 50 °C.
Note: The drying step might take a while due to the aqueous nature of the sample.
Add 300 μL of 2 M TFA to the dried samples. Caution: TFA is corrosive and attacks skin, eyes, and mucous membranes. Wear chemical-resistant gloves and safety goggles and work with TFA only under a fume hood.
Hydrolyse the samples for 1 h at 121 °C in a heating block for DURAN glass tubes.
After hydrolysis, evaporate the TFA under N2 stream.
Add 500 μL of water and 100 μL of the internal standard 0.05 mg/mL allo-inositol to the dried hydrolysis products. Pause point: For long-term storage, dissolved hydrolysis products can be kept at -20 °C for several months.
Sample preparation for subsequent measurements: dry 20 μL of the dissolved hydrolysis products under a stream of N2 gas.
Derivatise the dried samples with 15 μL of MOX (30 mg/mL in anhydrous pyridine) at room temperature overnight. Use freshly prepared MOX (see General note 3). Wear chemical-resistant gloves and work with pyridine and MOX solution only under a fume hood. Use solvent-resistant tips for pipetting.
After overnight derivatisation, add 30 μL of MSTFA and measure by GC–MS after at least 1 h incubation time at room temperature. Incubation time with MSTFA before GC–MS analysis should not exceed 6 h.
Pectin analysis from mucilage
Prepare at least four replicates of each genotype. Weigh 5 mg of seeds for each replicate and transfer them to a 2 mL microcentrifuge tube. Record the exact mass of prepared seeds.
Add 1.4 mL of water to the seeds and vortex for a few seconds.
Place samples on a rotary shaker for 2 h to extract mucilage.
Remove the samples from the shaker and allow the seeds to settle for 1–2 min. Do not centrifuge the samples.
Transfer 1 mL of the supernatant to a DURAN glass tube.
Dry the samples under a stream of N2 gas at 50 °C.
Note: The drying step might take a while due to the aqueous nature of the sample.
Add 300 μL of 2 M TFA to the dried samples.
Caution: TFA is corrosive and attacks skin, eyes, and mucous membranes. Wear chemical-resistant gloves and safety goggles and work with TFA only under a fume hood.
Hydrolyse the samples for 1 h at 121 °C in a heating block for DURAN glass tubes.
After hydrolysis, evaporate the TFA under N2 stream.
Add 500 μL of water and 100 μL of the internal standard 0.05 mg/mL allo-inositol to the dried hydrolysis products.
Pause point: For long-term storage, dissolved hydrolysis products can be kept at -20 °C for several months.
Sample preparation for subsequent measurements: dry 20 μL of the dissolved hydrolysis products under a stream of N2 gas.
Derivatise the dried samples with 15 μL of MOX (30 mg/mL in anhydrous pyridine) at room temperature overnight. Use freshly prepared MOX (see Notes). Wear chemical-resistant gloves and work with pyridine and MOX solution only under a fume hood. Use solvent-resistant tips for pipetting.
After overnight derivatisation, add 30 μL of MSTFA and measure by GC–MS after at least 1 h incubation time at room temperature. Incubation time with MSTFA before GC–MS analysis should not exceed 6 h.
GC–MS analysis
Preparation of standards for calibration
From the stock solutions of the monosaccharides, prepare standard combinations of defined molar amounts by combining aliquots of dilutions from the respective stock solutions with the correct volume. (For example, for a mix of 1 nmol arabinose and 1 nmol galactose, combine 4 μL each of 1/1,000 dilutions of the stock solutions of arabinose and galactose.) Add 5 μL of 0.05 mg/mL allo-inositol.
For less abundant pectic monosaccharides, prepare 3–5 different molar amount samples in the range of 1–5 nmol. For highly abundant pectic monosaccharides, prepare 3–5 different molar amount samples in the range of 20–100 nmol. Low or high abundance of the various monosaccharides depends on the sample; galacturonic acid will generally be highly abundant, whereas rhamnose is expected to be highly abundant in seed mucilage but less abundant in AIR from leaves. If the sample composition is unclear, it might be helpful to compare initial GC–MS results of a low and a high concentration of each standard with the sample result and then adjust the concentrations accordingly. You can combine different monosaccharide standards in the following combinations:
1. Xylose, fucose, mannose
2. Arabinose, galactose
3. Glucose, glucuronic acid
4. Rhamnose, galacturonic acid
Prepare all standard combinations and different molar amounts in triplicates.
Dry the prepared standard solutions under a stream of N2 gas.
Derivatise the dried standards with 15 μL of MOX (30 mg/mL in anhydrous pyridine) at room temperature overnight. Use freshly prepared MOX solution (see General note 3). Wear chemical-resistant gloves and work with pyridine and MOX solution only under a fume hood. Use solvent-resistant tips for pipetting.
After overnight derivatisation, add 30 μL of MSTFA and analyse on GC–MS after at least 1 h incubation time. Incubation time with MSTFA before GC–MS analysis should not exceed 6 h.
Monosaccharide analysis with GC-MS
Inject 1 μL of the derivatised sample with a split value of 10. Helium is used as carrier gas at a flow rate of 1 mL/min.
Use the following temperature gradient: 150 °C for 2 min, ramp to 250 °C at 5 K/min (20 min), ramp to 320 °C at 15 K/min (4.67 min), 320 °C for 3 min. The inlet temperature is set to 250 °C.
The transfer line temperature is set at 280 °C; ionisation is done with an electron energy of 70 eV and an ion source temperature of 230 °C. Mass spectra are recorded in an m/z range of 40–500.
Data processing and peak integration
Perform peak assignment and integration according to your usual procedure. Here, we describe exemplarily data processing and peak integration with MSD ChemStation Data Analysis.
Define compound list for the individual monosaccharides in the software MSD ChemStation Data Analysis. Use the GC–MS runs of the standards to determine retention times and characteristic fragment ions of the individual monosaccharides. Include the internal standard allo-inositol as additional compound.
Note: For most monosaccharides, two peaks will be detected (Figure 1), as derivatisation with MOX yields two stereoisomers (compare retention indices in Table 1). For quantification, select the analyte peak without interference from other monosaccharide-derived analyte peaks. If there is no interference for any analyte peak of an individual monosaccharide, select the largest one.
Figure 1. Example of the GC–MS chromatogram of monosaccharides from mucilage pectin as observed with MSD ChemStation Data Analysis.
The vertical axis shows detected signals, while the horizontal axis shows the retention time in minutes after injection. allo-Inositol was used as internal standard (4). Fucose and glucuronic acid were not detected in this sample; the respective analyte peaks would have retention times of ca. 8.9 and 9.1 min for fucose, and 12.8 and 13.6 min for glucuronic acid. The peak at 9.1 min does not have the correct mass spectrum when compared with an external fucose standard.
Table 1. GC–MS detection of individual monosaccharides. Possible target ions for quantification and the device-independent retention index are given in the middle columns. As an example, the retention times of the different monosaccharides as shown in Figure 1 are also given. Monosaccharide peaks that could not be detected in the respective sample are denoted as n.d.
m/z quantitation ion Retention index Retention time in Figure 1
allo-Inositol 318 1908 11.7 min
L-Arabinose 307 1677 7.8 min
D-Xylose 307 1662, 1671 7.5 min, 7.7 min
L-Fucose 277 1744, 1756 both peaks n.d.
L-Rhamnose 277 1735, 1742 8.7 min, 8.9 min
D-Galactose 319 1928, 1950 12.0 min, second peak n.d.
D-Glucose 319 1935, 1954 12.1 min, second peak n.d.
D-Mannose 319 1922, 1937 11.9 min, second peak n.d.
D-Galacturonic acid 333 1986, 2004 13.0 min, 13.3 min
D-Glucuronic acid 333 1977, 1992 both peaks n.d.
Define one characteristic target ion as quantitation signal for all compounds. See Table 1 for possible quantitation ions of the derivatised analytes of the indicated monosaccharides.
The software will integrate the peaks of the target ions belonging to the different monosaccharides in the user-defined compound list. After automatic integration, it is important to check that monosaccharide peaks are correctly assigned, and peak areas are accurately integrated. If necessary, correct peak assignment and integration.
Re-integrate the peak intensities after correction of peak assignments or integration. Export the values for further analysis in Excel.
Data analysis
The exported peak integration values can be further processed with Microsoft Excel or equivalent software. An example calculation to obtain the molar amount of xylose depicted in the sample of Figure 1 is displayed at the end of this section.
Analysis of standards
Paste the values obtained with MSD ChemStation Data Analysis into an Excel spreadsheet.
Normalise each peak area of the different monosaccharide peaks to the internal standard allo-inositol. To that end, calculate the ratio of the respective peak areas and the integrated peak area of allo-inositol.
Plot the normalised peak area vs. the molar amounts of the standard and perform a linear regression analysis on the plot. The R2 value should be close to 1.
Use the LINEST function in Excel to calculate the parameters of a linear correlation between peak area and molar amount of the individual monosaccharide standards with the least squares method. The function will calculate the values m and b of the following model:
Note down the respective values of m and b for each individual monosaccharide peak. These are required for sample analysis.
Sample analysis
As for the standards, paste the values obtained with MSD ChemStation Data Analysis into an Excel spreadsheet.
Normalise the peak areas of the different monosaccharides to the internal standard allo-inositol.
Calculate the initial normalisation by forming the ratio between the peak area of the individual monosaccharides and the peak area of the standard allo-inositol.
In the standards, a total of 0.25 μg of allo-inositol was added (5 μL × 0.05 mg/mL), whereas in the samples 5 μg of allo-inositol (100 μL × 0.05 mg/mL) were added to the hydrolysis product. To correct against these different amounts of the standard, multiply the values for the normalised peak area obtained above (step B2a) with a correction factor of 20 (5 μg/0.25 μg = 20).
Using the normalised peak area of the monosaccharides and the linear regression parameters m and b obtained above, you can now calculate the molar amounts of the individual monosaccharides with the following formula:
Normalise the obtained molar amount to the initial sample mass of AIR or seeds. You can use the normalised molar amounts to determine the relative composition of your pectin samples. Calculate average and standard deviations for the percentages of the different monosaccharide compounds. You can also normalise the contribution of the less abundant monosaccharides to one of the dominating monosaccharides (see for example Guzha et al., 2022).
The measured values, calculations, and calculated values for the second xylose peak of Figure 1 are shown below as an example (Table 2).
Table 2. Exemplary values for the calculation of molar amounts of xylose.
The first column highlights the concentrations used for the external standard of xylose, all prepared in triplicates. The second and third columns show the integrated peak values as obtained from MSD ChemStation Data Analysis for xylose and allo-inositol, respectively. For the normalised values (column four), the ratio of values for xylose to the values for allo-inositol was calculated. In the lower section of the table, the averaged values for each concentration of the xylose standard are shown (Data analysis steps A1 and A2).
External standard Peak area of xylose [AU] Peak area of allo-inositol [AU] Normalised values
1 nmol 20176173 89977965 0.22423
1 nmol 16685600 78745304 0.21189
1 nmol 15301979 76681503 0.19955
2 nmol 33336682 76622222 0.43507
2 nmol 34115172 73181461 0.46617
2 nmol 31752336 68480181 0.46367
3 nmol 46560441 60818809 0.76555
3 nmol 47484312 61529497 0.77173
3 nmol 52695435 70783534 0.74445
Average normalised values
1 mol 0.21189
2 mol 0.45497
3 mol 0.76058
With the LINEST function of Excel, the correlation between the concentration of the standards and the average normalised values was calculated, leading to the following parameters (Data analysis step A4):
m = 0.27434; b = -0.07287.
For the sample shown in Figure 1, the following peak areas were integrated:
Xylose peak 2: 4484042; allo-inositol: 100751631
This enabled the calculations shown below:
Normalised value according to B2a:
Corrected normalised value according to B2b:
Molar amount of xylose according to B3:
Validation of protocol
This protocol or parts of it have been used in the following research articles: Guzha et al. (2022). Cell wall–localized BETA-XYLOSIDASE4 contributes to immunity of Arabidopsis against Botrytis cinerea (Figure 8E, 9F, S3A, S8, and S13).
General notes and troubleshooting
For mucilage analysis, it is critical that the mother plants (wild type and mutants) are grown together at the same standard growth conditions. Water the plants until the final siliques turn yellow and begin to dry out. A detailed description of plant growth conditions for mucilage analysis can be found in Dean et al. (2019).
Grow several independent seed sets (including wild type) and perform independent mucilage analyses to detect variations caused by slightly different growth conditions.
Starch contamination in the sample will cause excessive glucose signals. For leaf samples, plastidial starch accumulation can be avoided by keeping the plants in the dark for 48 h before harvest. Alternatively, starch can be removed by an α-amylase digest followed by an ethanol extraction after AIR preparation (Pettolino et al., 2012).
The derivatisation solution of 30 mg/mL MOX in anhydrous pyridine should not be older than three days to limit contaminations with water.
Calibration values obtained from one set of standards can be used for an extended time, as long as the sample type stays the same. In our hands, calibration values could be used for ~100 sample analyses.
Acknowledgments
This protocol was derived from Guzha et al. (2022). We are grateful to Drs. George Haughn, Gillian Dean, Robert McGee and Krešimir Šola at the University of British Columbia for advice and helpful discussions. This work was supported by German Research Foundation (DFG, IS 273/10-1, IRTG 2172 PRoTECT (project number 273134146) to T.I), the Studienstiftung des Deutschen Volkes (stipend to P.S.), and a grant from the U.S. Department of Energy, Office of Science, BER program DE-SC0020325 to K.D.C.
Competing interests
The authors have no financial or non-financial competing interests.
References
Atmodjo, M. A., Hao, Z. and Mohnen, D. (2013). Evolving views of pectin biosynthesis. Annu Rev Plant Biol 64: 747-779.
Bethke, G., Thao, A., Xiong, G., Li, B., Soltis, N. E., Hatsugai, N., Hillmer, R. A., Katagiri, F., Kliebenstein, D. J., Pauly, M., et al. (2016). Pectin Biosynthesis Is Critical for Cell Wall Integrity and Immunity in Arabidopsis thaliana. Plant Cell 28(2): 537-556.
Bethke G. and Glazebrook J. (2019). Measuring Pectin Properties to Track Cell Wall Alterations During Plant-Pathogen Interactions. Methods Mol Biol 1991: 55-60.
Biswal A. K., Tan L., Atmodjo M. A., DeMartini J., Gelineo-Albersheim I., Hunt K., Black I. M., Mohanty S.S., Ryno D., Wyman C. E., et al. (2017). Comparison of four glycosyl residue composition methods for effectiveness in detecting sugars from cell walls of dicot and grass tissues. Biotechnol Biofuels 10: 182.
Dean, G. H., Sola, K., Unda, F., Mansfield, S. D. and Haughn, G. W. (2019). Analysis of Monosaccharides from Arabidopsis Seed Mucilage and Whole Seeds Using HPAEC-PAD. Bio Protoc 9(24): e3464.
Guzha, A., McGee, R., Scholz, P., Hartken, D., Lüdke, D., Bauer, K., Wenig, M., Zienkiewicz, K., Herrfurth, C., Feussner, I., et al. (2022). Cell wall-localized BETA-XYLOSIDASE4 contributes to immunity of Arabidopsis against Botrytis cinerea. Plant Physiol 189(3): 1794-1813.
Harholt, J., Suttangkakul, A. and Vibe Scheller, H. (2010). Biosynthesis of pectin. Plant Physiol 153(2): 384-395.
Höfte, H. and Voxeur, A. (2017). Plant cell walls. Curr Biol 27(17): R865-R870.
Pettolino, F. A., Walsh, C., Fincher, G. B. and Bacic, A. (2012). Determining the polysaccharide composition of plant cell walls. Nat Protoc 7(9): 1590-1607.
Shin, Y., Chane, A., Jung, M. and Lee, Y. (2021). Recent Advances in Understanding the Roles of Pectin as an Active Participant in Plant Signaling Networks. Plants (Basel) 10(8): 1712.
Vogel, J. P., Raab, T. K., Somerville, C. R. and Somerville, S. C. (2004). Mutations in PMR5 result in powdery mildew resistance and altered cell wall composition. Plant J 40(6): 968-978.
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Biochemistry > Carbohydrate > Polysaccharide
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Update Notice: Efficient Generation of Genome-wide Libraries for Protein–ligand Screens Using Gibson Assembly
TS Tamara Sternlieb
ML Mira Loock
MG Mengjin Gao
IC Igor Cestari
Published: May 20, 2023
DOI: 10.21769/BioProtoc.4747 Views: 298
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After official publication in Bio-protocol (https://bio-protocol.org/e4558), we would like to add an updated information to the Acknowledgments section: “This protocol is derived from the original research paper by Rhiannon Heslop et al. (2023).”
The updated version of the Acknowledgments section is “This work was funded by the Canadian Institutes of Health Research (CIHR PJT-175222, to IC), the Natural Sciences and Engineering Research Council of Canada (NSERC RGPIN-2019-05271, to IC), the Canada Foundation for Innovation (JELF 258389, to IC), and by McGill University (130251, to IC). We thank Andressa Lira and Sahil Sanghi for their technical support at the early stages of this work. This research was enabled in part by computational resources provided by Calcul Quebec (https://www.calculquebec.ca/en/) and Compute Canada (www.computecanada.ca). This protocol is derived from the original research paper by Rhiannon Heslop et al. (2023).
Accordingly, a new reference is added to the References list, “17. Heslop, R., Gao, M., Brito Lira, A., Sternlieb, T., Loock, M., Sanghi, S. R. and Cestari, I. (2023). Genome-Wide Libraries for Protozoan Pathogen Drug Target Screening Using Yeast Surface Display. ACS Infect. Dis: e2c00568.”
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Human Schwann Cells in vitro I. Nerve Tissue Processing, Pre-degeneration, Isolation, and Culturing of Primary Cells
GA Gabriela I. Aparicio
PM Paula V. Monje
Published: Vol 13, Iss 22, Nov 20, 2023
DOI: 10.21769/BioProtoc.4748 Views: 898
Reviewed by: Vivien J. Coulson-ThomasCarmen Melendez-VasquezHeleen van 't Spijker
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The authors used this protocol in Molecular Neurobiology Aug 2018
Abstract
This paper presents versatile protocols to prepare primary human Schwann cell (hSC) cultures from mature peripheral nervous system tissues, including fascicles from long spinal nerves, nerve roots, and ganglia. This protocol starts with a description of nerve tissue procurement, handling, and dissection to obtain tissue sections suitable for hSC isolation and culturing. A description follows on how to disintegrate the nerve tissue by delayed enzymatic dissociation, plate the initial cell suspensions on a two-dimensional substrate, and culture the primary hSCs. Each section contains detailed procedures, technical notes, and background information to aid investigators in understanding and managing all steps. Some general recommendations are made to optimize the recovery, growth, and purity of the hSC cultures irrespective of the tissue source. These recommendations include: (1) pre-culturing epineurium- and perineurium-free nerve fascicles under conditions of adherence or suspension depending on the size of the explants to facilitate the release of proliferative, in vitro–activated hSCs; (2) plating the initial cell suspensions as individual droplets on a laminin-coated substrate to expedite cell adhesion and thereby increase the recovery of viable cells; and (3) culturing the fascicles (pre-degeneration step) and the cells derived therefrom in mitogen- and serum-supplemented medium to accelerate hSC dedifferentiation and promote mitogenesis before and after tissue dissociation, respectively. The hSC cultures obtained as suggested in this protocol are suitable for assorted basic and translational research applications. With the appropriate adaptations, donor-relevant hSC cultures can be prepared using fresh or postmortem tissue biospecimens of a wide range of types and sizes.
Keywords: Human Schwann cells Ensheathing glia Peripheral nerve Tissue procurement Tissue culture Cell culture Pre-degeneration Mitogenic factors Adherent substrates Fibroblasts
Background
Schwann cells (SCs) are a heterogeneous group of axon-ensheathing cells in the peripheral nervous system of all vertebrate species (Jessen et al., 2015). These nerve-resident neuroglial cells can be isolated and expanded in vitro using standard cell culture techniques. Human SC (hSC) cultures can be prepared using any type of nerve or ganglion from developing and adult organ donors [reviewed recently in Monje (2020)]. Culturing hSCs is a lengthier and more labor-intensive process than culturing SCs from rodents and other experimental animals. However, the hSCs obtained from an initial harvest can be amplified substantially in vitro to generate large numbers of cells for a wide range of experimental approaches (Figure 1). Once established, the hSC cultures can be managed in a manner similar to adherent cell lines, though there are limitations regarding the expandability of individual stocks.
Figure 1. Scalable workflow for the preparation of nerve-derived human Schwann cell (hSC) cultures. The protocols described in this article and associated manuscripts (Monje, 2023a and 2023b) address the following basic procedures: (1) tissue procurement, dissection, and culture of fascicles (pre-degeneration); (2) enzymatic dissociation, isolation, and culture of primary hSCs; (3) derivation of established hSC cultures and amplification via serial passaging; (4) routine manipulations in vitro (e.g., purification, cryopreservation, labeling, and gene delivery); and (5) quality control of identity and bioactivity of the cells to be used in experimentation. Our protocols are adaptable; for instance, the number of hSCs can be scaled up by increasing the size of the tissue specimens used for cell isolation (primary cultures) and expanding the populations in medium containing mitogenic factors and serum (established cultures).
SC cultures from human tissues have facilitated numerous discoveries over nearly four decades [reviewed in Guest et al. (2013); Monje et al. (2021); Vallejo et al. (2022)]. These cultures are accurate in vitro models for studying neural development, differentiation, regeneration, electrophysiology, and toxicology in normal and disease states. They are also valuable for cell therapy development to repair damage caused by trauma and neurodegenerative disease. Indeed, the transplantation of cultured hSCs from a patient’s sural nerve has been implemented as a strategy to treat spinal cord and peripheral nerve injuries in USA-FDA-regulated clinical trials (Levi et al., 2016; Anderson et al., 2017; Khan et al., 2021).
Empirical data have shown that highly viable, expandable hSC cultures can be established from tissues provided by donors > 60 years of age [reviewed in Bunge et al. (2017)]. Biospecimens from live donors are not required. In fact, hSC cultures from postmortem tissues are phenotypically indistinguishable from those obtained from live donors (Boyer et al., 1994; Casella et al., 1996; Bastidas et al., 2017). Importantly, in vitro cultured SCs from adult nerves retain their ability to proliferate in response to axon contact, promote axonal growth, and form a myelin sheath (Morrissey et al., 1991), which are key functions of SCs during nerve development, maturation, and repair (Jessen et al., 2015).
Two main obstacles must be overcome to establish SC cultures from humans: (1) harvesting sufficient proliferative hSCs from the source tissue; and (2) expanding these primary hSCs sufficiently while maintaining low levels of fibroblast contamination (Morrissey et al., 1995; Peng et al., 2020). Processing adult biospecimens for cell isolation is more time demanding and challenging than doing so from developing (embryonic, neonatal) nerves. This is at least in part due to the presence of multiple layers of connective tissue (CT) and extracellular matrix (ECM), both within and around the SC-enriched fascicles. In addition, the highly elaborate cellular architecture of mature myelinating and ensheathing (non-myelinating or Remak) SCs, which are among the largest cells in the body, impose a hard-to-overcome technical barrier for disintegrating the tissue without disturbing the integrity of the cells. Although isolating hSCs immediately after nerve harvesting is feasible under certain conditions (Weiss et al., 2016), we strongly recommend pre-culturing the tissues without dissociation to achieve more consistent cell yields (Bunge et al., 2017; Chu et al., 2022). We also recommend selective immunological purification protocols to tackle the problem of fibroblast overgrowth, although other methods are available to enrich hSCs over non-glial cell populations.
In the following sections, we present generic protocols for preparing primary hSC cultures by including procedures for: (1) the dissection and harvesting of fascicles, roots, and ganglia from adult human donors; (2) the pre-degeneration of nerve segments, an intermediate step that involves the culturing of intact tissues to enrich the number of hSCs; (3) the disintegration of cultured tissues by proteolytic enzymes and the plating of initial cell suspensions on an adherent substrate; and (4) the growth and management of primary hSCs at the desired quality and quantity (Figures 1 and 2). Relevant notes on the procurement, storage, and handling of patient-derived biospecimens are included along with two different protocols for processing large and small tissue biospecimens. Various recommendations are made for managing the hSC cultures during the initial stages of growth. Supportive data on the properties of hSC cultures prepared according to these methods can be found in our published studies (Monje et al., 2006, 2008 and 2018). Our methodologies were developed using assorted tissue sources from deidentified donors. Importantly, the methods presented here are intended for non-clinical research only and differ substantially from those used in preclinical research (Bastidas et al., 2017) and clinical trials (Khan et al., 2021). However, it should be mentioned that in vitro cultured hSCs from nerves, skin, and ganglia are expected to exhibit fairly similar phenotypic and functional characteristics regardless of the tissue of origin and the mode of preparation (Stratton et al., 2017; Monje et al., 2018 and 2020; Chu et al., 2022).
Figure 2. Isolation and culture of primary human Schwann cell (hSCs). The diagram depicts the overall cell culture strategy, highlighting the timeline for each procedure and the main steps involved, as illustrated by the representative images. This workflow starts with the bioprocessing of peripheral nerve tissues (steps 1–3) and ends with the establishment of confluent cultures of primary hSCs ready to use (step 11). The entire process can take at least three weeks, depending mostly on the length of the pre-degeneration phase (step 4, tissue culture).
Materials and reagents
All materials, reagents, and solutions should be endotoxin-free and cell culture grade. Except for dissection tools that are cleaned and sterilized for each procedure, most laboratory ware can be acquired as disposable materials for single use. It is recommended to use commercially available cell culture grade water to prepare all solutions, buffers, and culture media. In the sections below, we have added products’ information for reference only. We have mainly used disposable cell culture–treated flasks and dishes from CorningTM, but products from other brands may be equally suitable.
Supplies and consumables
Dumont forceps #3, #4, and #5; straight tip shape (Fine Science Tools, catalog numbers: 11231-30, 11242-30, and 11251-10, respectively)
Spring scissors, angled to side (Fine Science Tools, catalog number: 15006-09)
Moria Dowell spring scissors, straight tip shape (Fine Science Tools, catalog number: 15372-62)
Disposable 5 mL serological pipettes, polystyrene, sterile (VWR, catalog number: 89130-896)
Disposable 10 mL serological pipettes, polystyrene, sterile (VWR, catalog number: 19221005)
Polystyrene Pasteur pipettes, sterile and individually wrapped (VWR, Argos Technology, catalog number: 10122-560)
Borosilicate glass Pasteur (transfer) pipettes (Corning, catalog number: 7095D-9) with attached rubber bulb
100 mm Petri dishes (Corning, catalog number: 351029)
Polypropylene conical-bottom centrifuge tubes, 15 and 50 mL (Corning, catalog numbers: 430791 and 430290, respectively)
Round-bottom centrifuge tubes with a snap cap, polypropylene, 15 mL (BD, catalog number: 352059)
Polystyrene cell culture dishes, 35, 60, or 100 mm (Corning, catalog numbers: 353001, 353002, and 353003, respectively)
24-well plate, flat bottom (Corning, catalog number: 3524)
Containers with wet ice
Media, supplements, and reagents for cell culture
Distilled water, cell culture grade (Thermo Fisher Scientific, Gibco, catalog number: 15-230-147)
Dulbecco’s phosphate-buffered saline (DPBS), pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14190)
Hank’s balanced salt solution (HBSS), formulated without calcium or magnesium and containing phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 14170-112)
Leibovitz’s L15 medium (Thermo Fisher Scientific, Gibco, catalog number: 11415064)
Dulbecco’s modified Eagle’s medium (DMEM) with high glucose and phenol red, pH 7.2 (Thermo Fisher Scientific, Gibco, catalog number: 11965092)
1,000× Gentamycin (Thermo Fisher Scientific, Gibco, catalog number: 15750-060)
De-complemented, gamma-irradiated fetal bovine serum (FBS) (HyClone, catalog number: SV 30014.03)
100× GlutaMAX supplement (Thermo Fisher Scientific, Gibco, catalog number: 35050061)
Forskolin (Sigma-Aldrich, catalog number: F68861)
Heregulin-β1177-244 (referred to as heregulin) (Preprotech, catalog number: G-100-03)
Laminin stock solution, consisting of a sterile 1 mg/mL laminin from Engelbreth-Holm-Swarm murine sarcoma basement membrane (Sigma-Aldrich, catalog number: L2020). Store in aliquots at -80 °C and use as described in Andersen and Monje (2018)
Poly-L-lysine (PLL) stock solution (Sigma, catalog number: P-2636). Prepare, store in aliquots at -80 °C, and use as described in Andersen and Monje (2018)
Matrigel growth factor reduced basement membrane matrix, phenol red-free (BD Biosciences Discovery Labware, catalog number: 356231)
Dispase II or neutral protease, ≥ 0.8 units/mg protein, lyophilized powder (Roche, catalog number: 165-859)
Collagenase Type I, ≥ 125 units per milligram dry weight, dialyzed, lyophilized powder (Worthington, CLS-1, catalog number: 4196)
Dissection medium (DM) (see Recipes)
Low proliferation medium (LP) (see Recipes)
High proliferation medium (HP) (see Recipes)
Laminin coating solution (see Recipes)
10× enzymatic solution (see Recipes)
Antibodies and fluorescent dyes
Anti-NGFR mouse IgG monoclonal antibody, produced in-house from the HB-8737 hybridoma cell line (reactivity: human/primate-specific NGFR; American Type Culture Collection, ATCC). See Ravelo et al. (2018) for a step-by-step description of NGFR immunostaining in live and fixed hSC cultures. (Optional) Use the rabbit monoclonal antibody EP1039Y (Abcam, catalog number: ab52987).
Anti-O4 mouse IgM monoclonal antibody, produced in-house from the O4 hybridoma cell line (reactivity: human/rat/mouse/pig/other; kindly provided by Dr. Melitta Schachner). See Ravelo et al. (2018) for a step-by-step description of O4 immunostaining in live hSC cultures. (Optional) Use a commercially available purified O4 antibody (Novus Biologicals, catalog number: NL637)
Hoechst 33342 (Sigma, catalog number: B2261), prepared in water at 1 mg/mL
Syto-24 green, fluorescent nucleic acid stain (Invitrogen, catalog number: S75559)
Propidium iodide (PI) nucleic acid stain (Sigma, catalog number: P4170) prepared in water at 1 mg/mL. See Ravelo et al. (2018) for a step-by-step description of viability assays using PI and Hoechst 33342 or Syto-24 green
FluoroMyelin red (or green), fluorescent myelin stain (Invitrogen, catalog number: F34652)
FM 4-64FX, fixable analog of FM 4-64 membrane stain (Invitrogen, catalog number: F34653)
4’,6-Diamidino-2-Phenylindole, dilactate (DAPI) (Invitrogen, catalog number: D3571), prepared in water at 1 mg/mL
Equipment
Stereomicroscope with an attached digital camera (Zeiss, Stemi 305/Axio cam ER C52)
Double gooseneck fiber optics with intensity control for cool white light illumination (DolanJenner)
NIGHTSEA® Fluorescence Viewing System equipped with blue and green barrier filters (Electron Microscopy Sciences)
Inverted phase contrast microscope with an attached digital camera (VWR V5MP)
Inverted fluorescence microscope with an attached digital camera (Olympus IX71)
Benchtop centrifuge (Beckman Coulter, model: Allegra X-I2R)
Cell counter for automated counting of cells in suspension (Bio-Rad, TC20 Automated cell counter). (Optional) Hemocytometer for manual cell counting
CO2 cell incubator set up at 37 °C and 8%–9% CO2 (Thermo Fisher Scientific, Forma Steri-Cycle)
Biosafety cabinet, BL2 level (Thermo Fisher Scientific, 1300 Series A2)
Germinator 500, bench-top sterilizer (Cell Point, catalog number: 5517)
Procedure
Protocol 1: Procurement and dissection of human nerve tissues
This protocol relies on the availability of nerves or ganglia harvested under aseptic conditions. Freshly collected tissue remains from patients undergoing therapeutic or plastic surgeries or diagnostic biopsies are well-suited and may be preferred for hSC culturing if made available promptly from local providers. Nerves harvested postmortem are also appropriate if collected and stored properly until arrival at the laboratory.
Most tissues that become available for research are procured from adult subjects. Embryonic and fetal tissues render highly proliferative hSC cultures (Boyer et al., 1994; Scarpini et al., 1988), but they may be difficult to obtain due to ethical concerns or other restrictions. The nerve biospecimens may not be limited to a particular type or size as long as the appearance is normal, and the anatomical layers are clearly distinguishable. The best results are obtained by using nerves (or ganglia) of sufficient length (> 1 cm) and caliber (> 2–3 mm) because they can be manipulated and visualized with ease under a standard stereomicroscope. The protocols below explain how to manually isolate individual segments (fascicles, roots, and ganglia) from the surrounding connective tissue using typical long segmental nerves (Protocol 1B) and dorsal root ganglia (DRG) with attached nerve roots (Protocol 1C). Removing as much ECM as possible from the outside of the nerve or ganglia can prevent the introduction of contaminating fibroblasts into the hSC cultures. The dissection protocols will have to be adapted for the isolation of hSCs from normal tissues enriched in nerve terminals, such as the gut or the skin, and pathological tissues known to contain hSCs, such as tumors from nerves or the skin.
Tissue procurement, handling, and storage
Collect and store the human nerve in a sterile container filled with an excess volume of storage medium as appropriate for the size of the specimen, e.g., 50 mL of medium for a specimen that is 5 cm long. Use a gentamycin-supplemented balanced solution, such as University of Wisconsin (UW)® cold storage solution, high-glucose DMEM, or L15 medium, for storage. It is important to minimize the time between the surgical removal of the tissue and its immersion in the storage medium. Maintain the container in wet ice (4 °C) until beginning the dissection procedure (see Note a).
Transfer the tissue to the laboratory and start the cleaning of the nerve as soon as possible. If the procedure cannot be started immediately, the tissue can be safely stored inside a 4 °C refrigerator (for more details on storage time and conditions, please see Recommendations and Troubleshooting).
Isolation of fascicles from typical mature nerves
Transfer the nerve into a sterile 100 or 150 mm Petri dish (depending on the size of the biospecimen) containing ice-cold DM (see Recipe 1) (Figure 3A).
Figure 3. Separation of nerve fascicles from connective tissue. Low magnification images of a whole nerve biopsy (tibial) as received from a surgical procedure and its dissection under a stereomicroscope. (A) A nerve fragment placed in a Petri dish. (B) Removal of externally attached tissues: connective tissue (CT) and blood vessels (BV). (C) Appearance of a clean nerve. (D and E) Sectioning of the nerve into smaller segments. (F and G) Fascicle separation from the surrounding connective tissue. (H) Final harvest. White arrow: fascicles to be transferred to a culture dish; yellow arrow: connective tissue to be discarded. (I and J) Selected fascicles inside a tissue culture dish ready to be cultured.
By working under the dissecting stereomicroscope illuminated with a cool light system, carefully remove the loosely attached collagen-enriched connective tissue, blood vessels (BV), fat, and muscle around the nerve using the appropriate combination of tools. For instance, for biospecimens > 2 cm in length, use blunt forceps or number 3 fine forceps to hold the nerve from one end, and spring scissors or forceps to remove the excess tissue around the nerve (Figure 3B) (see Note b).
Replace the DM (or transfer the nerves to new dishes) as frequently as needed to maintain an as-clean-as-possible solution during the whole cleaning procedure. Use ice-cold solutions and work fast to prevent an increase in temperature and consequent sample alteration. This procedure may be labor intensive depending on the length and condition of the nerve specimen. Nerves of a higher caliber, such as the adult human sciatic nerve, can contain adipose tissue surrounding and intermixing between the fascicles.
Once the nerves are cleared of loosely attached external material (Figure 3C), use sharp scissors (e.g., Metzenbaum scissors or spring scissors, depending on the caliber of the specimen) to cut the specimens transversally and generate smaller, more manageable segments. These segments can be ~1 cm and ~2–3 cm long for larger (e.g., sciatic) and lower caliber (e.g., tibial) nerves, respectively (Figure 3D and 3E). Ensure that the proximal (close to the spinal cord) and distal (close to the periphery) ends of the segments are appropriately oriented for expedited fascicle removal (see step 6).
Transfer the individual segments into a new dish containing DM and work with one segment at a time. Visually identify the individual fascicles (in cross section) protruding in between the surrounding connective tissue (whitish material) by exposing the sectioned area on the distal end of the fascicles (Figure 4A–4C).
Separate the fascicles from the collagenous connective tissue one by one by grabbing them from their distal end and pulling outward using #4 or #5 forceps while gently holding the proximal side of the nerve segment with another pair of forceps. Always pull the nerves out in the same direction while visually following their trajectory until they separate fully. If the tissue is maintained steady from the proximal end, all fascicles within one segment can be pulled out quickly and smoothly, thus leaving a carcass of collapsible connective tissue composed mostly of the epifascicular and interfascicular epineurial layers (Figure 3F–3H, 4B–4D, and 5G–5H) (see Note c).
Figure 4. Technique to separate nerve fascicles. Upper-left panel: nerve fascicles are a bundle of fibers [axon–Schwann cell (SC) units] enclosed by the perineurium layer. These bundles are organized into larger bundles surrounded by substantial extracellular matrix (ECM) and enclosed within a continuous membranous layer, the epineurium. Human SC (hSC) cultures are derived from mature myelinating (shown in the diagram) and non-myelinating/Remak (not shown) SCs within the fascicles that become proliferative during the tissue culture step. Upper-right panel: higher magnification views of individual fascicles protruding from the distal end of a nerve segment before (A) and at the time of being detached from the connective tissue layers (B, C). (D) Close-up views of a nerve fascicle exhibiting clearly defined Fontana bands (top) and the corresponding collapsible layers comprising the epineurium and possibly also the perineurium (bottom). Lower panels: (E and F) Low-magnification images of a human sciatic nerve (seen in cross section) labeled with DAPI (nuclei, blue) and FM 4-64FX (myelin, red). The brightfield image was combined with the fluorescence images (F) to visualize the complex organization of the nerve consisting of endoneurium (green line, E), perineurium (red line, E), and epineurium (black line, E). The fascicles (blue line, E–F) are surrounded by multilayered perineurial and epineurial membranes enriched in blood vessels (BV, orange circles). (i–iv) Zoom-in images to highlight myelinating SCs depicting myelin sheaths of various calibers. Section thickness was 20 µm.
(Optional) Discard the fascicles with indistinct boundaries, attached ECM, and/or perineurium unless they can be eliminated easily with fine forceps. These layers contain abundant non-glial cells and can introduce a higher-than-acceptable number of fibroblasts into the hSC cultures (see Note d and Figure 5G and 5H).
Proceed as described in Protocol 2A.
Figure 5. Characterization of nerve biospecimens suitable for cell culture. (A, B) Appearance of fascicles before and after removal of connective tissue (CT). (C–H) Fluorescent labeling of myelin (red) (C, D) and total cell nuclei (green) (F, H) in clean fibers and CT layers. The tissues were stained live using Fluoromyelin Red (D) and Syto-24 (F and H), which detect myelin and cell nuclei, respectively. Cells from the CT can contaminate the hSC cultures if not eliminated from the onset (G, H).
Isolation of nerve roots and ganglia from the adult DRG
Transfer the tissue into a 100 mm dish containing ice-cold DM. Carefully trim off the external connective tissue using a combination of spring scissors and fine forceps. Expose the body of the ganglia and the protruding nerve roots as shown in Figure 6A and 6B.
Transfer the partially cleaned DRG with its attached roots to a new 100 mm dish with ice-cold DM. Using spring scissors, cut away all remaining connective tissue capsule trying not to pinch the roots or the ganglia. Then, cut off the nerve roots by positioning the spring scissors as close as possible to the DRG body. The clean DRG has a creamy-yellowish, spongy appearance with a heart or football shape. The roots have a whiteish appearance (see Note d). If the roots or DRGs have bloody areas, they should be cut away together with any remaining roots, fat, or connective tissue (Figure 6C and 6D).
Transfer the roots and DRG bodies separately to new dishes containing DM. Use fine spring scissors or scalpels (with #11 blades, stainless steel) to slice the nerve roots into smaller 1 mm segments (Figure 6E and 6F). Use the same tools to cut the DRG bodies longitudinally into halves, then fourths, then eighths and so forth up until the segments are approximately 1 mm in diameter. Notice that the individual DRG neurons are observable at low (4×) magnification.
Proceed as described in Protocol 2B.
Notes:
The conditions of nerve harvesting, transfer, and storage for the biospecimens may not be controlled and standardized, but a record for each material should be maintained regarding all relevant variables that can affect the viability of the cells. If tissues are provided by a local surgeon, the surgical team can be provided with enough collection tubes, ready-to-use storage medium, and other materials for expedited tissue transfer.
Prepare an excess of sterile dissection tools for the fast replacement of instrumentation. It is good practice to clean and sterilize the forceps as frequently as possible using a microbead sterilizer (e.g., Germinator) positioned inside the biosafety cabinet. Confirm that the tips of the fine forceps do not become damaged while performing the procedure. Replace distorted forceps as frequently as needed.
Most fascicles inside the collagenous carcass are identifiable by visual observation under the stereomicroscope (Figure 4A and 4B). When attempting to isolate the fascicles, hold each nerve segment with fine forceps at the opposite end from where the fascicles are pulled. Then, grip and pull each fascicle up until no fascicles remain, as per visual inspection, in any of the segments. Note that the number and caliber of individual fascicles recovered from each nerve can vary substantially from donor to donor, even when the nerves are taken from comparable anatomical locations. For instance, the human sural nerve, a nerve from the lower leg, has a variable number of fascicles (usually 3–20 bundles with an average of 10 per nerve) of a range of calibers (Mizia et al., 2014).
The fascicles and nerve roots should exhibit a smooth surface with striated whitish bands (bands of Fontana) and no visibly attached connective tissue (Figures 4D and 5B). The interfascicular epineurium and/or the perineurium can be pulled away easily using #5 forceps if they remain attached to the isolated fascicles. The perineurium is a delicate semi-transparent tube (veil) that surrounds the fascicles, but its presence is not always obvious by inspection under the stereomicroscope.
Figure 6. In vitro pre-degeneration of small tissue biospecimens: adherent explants inside a Matrigel drop. (A–D) General view of tissue specimens [dorsal root ganglion (DRG) with attached roots] as obtained from a surgical procedure (A, B), after the removal of external connective tissue (C), and during dissection of the roots (D). (E–G) Harvest of clean nerve roots, sectioning into 1 mm segments (explants), and immersion into a drop of Matrigel inside a culture well. (H–L) Cultured explants, shown at five and eight days post plating (DIV: days in vitro), depict the outgrowth of cells that continue to migrate, proliferate, and cover the available surface in the following weeks. This method can be used for the culture of hSCs from nerve roots (E–M) or ganglia (not shown), with the consideration that the culture conditions do not support the survival of the DRG neurons. (M) NGFR-immunostaining was performed to reveal the cellular constituents of the outgrowth. The cells around the explant comprise NGFR+ human Schwann cells (hSCs, magenta) and NGFR- fibroblasts (FB), as indicated. The cell nuclei were labeled with DAPI (cyan). Tissues were courtesy of Jamie Bradbury.
Protocol 2: Culturing of undissociated tissues
Tissue culturing has been implemented for over three decades to enhance the yields and viability of cell suspensions resulting from enzymatic disintegration of the nerves (Morrissey et al., 1991). Whereas immediate dissociation is best suited for SC isolation from developing nerves, delayed dissociation with an intermediate step of culturing is preferred for adult tissues (Morrissey et al., 1991). This step is often referred to as pre-degeneration due to its resemblance to the process of Wallerian degeneration associated with injury-induced axonal loss and SC activation in vivo (Jessen and Arthur-Farraj, 2019). The culture of undissociated nerves is understood to facilitate the prompt dedifferentiation of the hSCs within their basal lamina tubes while receiving trophic and nutritional support from the culture medium. Early studies revealed that the culturing of human nerve fascicles led to less cellular damage after dissociation (Casella et al., 1996). Even though adult rat nerves can be dissociated immediately after harvesting to generate enough SCs for cell culture (Andersen et al., 2016; Andersen and Monje, 2018), immediate dissociation of adult human nerves often leads to poor cell viability or inconsistent cell yields.
In the following sections, we suggest using distinct pre-degeneration methods according to the size and type of explant tissue. Whereas free-floating explants (Protocol 2A) are most suitable for large tissue specimens (nerves > 1 cm in length), adherent explants (Protocol 2B) are recommended for smaller-sized specimens (e.g., nerve roots or ganglionic tissue). In the former method, individual nerve fascicles are allowed to pre-degenerate while suspended in the culture medium. In the latter method, small pieces of tissue (1–3 mm in length) are immersed inside a droplet of Matrigel to stably attach them to the surface of a culture dish or well. Once the Matrigel matrix becomes a solid gel, culture medium can be flooded around the explant for continued culture. This method renders a population of mixed adherent cell types that migrate out from the explant, including hSCs (Figure 6H–6M).
Free-floating method
Starting with a harvest of nerve fascicles (Figure 3I–3J) and using a sterile transfer pipette, completely aspirate the dissection medium from the periphery of the dish without disturbing the fascicles.
Transfer the fascicles to a new culture dish, well, or flask (according to the volume of tissue) containing HP medium (see Recipe 3) by gently grabbing all the fascicles together using blunt forceps or aspirating them using a pre-wet, wide-bore, glass pipette.
Incubate the fascicles at 37 °C in an 8% CO2 incubator, ensuring they are immersed in medium.
Replace the HP medium on average every three days or as needed, e.g., when the medium turns yellowish (Figure 7A), for the entire pre-degeneration period. Aspirate the medium around the fascicles using a transfer pipette, avoiding the fascicles, and add an equivalent volume of new medium. Cells within the explants are highly active and acidification of the medium is expected (see Notes a and b).
Figure 7. In vitro pre-degeneration of large nerve biospecimens: floating explants. (A–G) Floating (non-attached) spinal nerves (as indicated) are shown at low (A and B) and high magnification (C–G) under white (C, E), blue (D, F), and green light (G) illumination (NIGHTSEA® stereomicroscope). The fascicles were pre-degenerated in HP medium inside a 100 mm dish. In C and D, degenerated fibers are shown next to degenerated connective tissue (CT) from the corresponding nerve segment to reveal the cellular enrichment within the fibers after staining live tissue with Syto-24 (nuclei, green). Pre-degenerated fibers exhibit scattered myelin debris, as shown by staining with FM 4-64FX (G, red). Note that the fibers rather than the CT are filled with cells, indicating that human Schwann cells (hSCs) rather than non-glial cells survive and multiply under our pre-degeneration conditions. The tissue-resident hSCs are highly active metabolically, especially within the first two weeks of culture, leading to quick acidification of the medium, which turns yellow (A). This acidification results from secreted metabolic products and should be distinguished from microbial growth by appropriate tests. The number of dead cells (PI positive) in these samples was negligible (not shown). Tissues shown in this figure were courtesy of Michael Murphy and Kristen Wanczyk.
Adherent explant method
Place 30–40 µL of cold Matrigel matrix in the center of a culture well (e.g., 24-well plate) or a dish placed on ice (see Note c).
Immediately transfer a 1 mm long piece of nerve or ganglia (Protocol 1C) into the matrix using fine forceps, ensuring the matrix embeds the tissue thoroughly (Figure 6G).
Immediately transfer the dish to the CO2 incubator.
Once the Matrigel becomes solid and the tissue explant is secured, use a transfer pipette to gently add culture medium around the explant to fill the dish.
Observe regularly and perform media changes as explained in the free-floating method (see Note d).
Notes:
The length of the pre-degeneration phase can range from one to five weeks. However, we have found that hSCs can remain alive within undissociated fascicles for a much longer time (>3 months) (unpublished). An important control is to assess the content and viability of cells inside pre-degenerated nerves before attempting an enzymatic dissociation (Figure 7D and 7F).
Reducing the size of the fascicles to increase the surface area and more efficiently expose the cells to soluble media components is recommended. By doing so, we expect to expedite the diffusion of the growth factors contained in the medium to act directly on the hSCs to promote their survival and multiplication.
Matrigel may not be needed for secure adhesion of the explants, but it is highly recommended to prevent accidental tissue loss during media changes. Tissue explants substantially shrink during culture and detachment can be problematic when dealing with smaller samples.
After the tissue explant is attached to the Matrigel, the hSCs start to come out of the explant into the surrounding Matrigel, and then into the substrate of the dish, usually within the first week (Figure 6J and 6L). Use precoated dishes with PLL (or equivalent) and laminin to favor the growth of adherent cells in a monolayer (Figure 6H and 6I). The cultures obtained in this manner contain a substantial proportion of fibroblasts (Figure 6M). Attachment of nerve explants to tissue culture plastic was initially used to reduce fibroblast contamination (Morrissey et al., 1991). Here, we use Matrigel as physical support to retain small tissue explants and prevent them from being lost during media changes rather than to eliminate fibroblasts.
Protocol 3: Nerve tissue dissociation, cell plating, and growth
Harvesting sufficient cells from nerve tissues is the most critical step in the culture workflow. Early empirical observations showed that enzymatic dissociation, as opposed to mechanical dissociation, is more appropriate to disorganize the adult nerve tissue (Casella et al., 1996), and that prolonged incubation with a combination of collagenase and neutral protease leads to more consistent cell yields (Casella et al., 1996; Casella et al., 2000). We have confirmed these observations and suggest proceeding with nerve tissue dissociation essentially as shown in the Casella et al. (1996 and 2000) papers (Protocol 3A) with a few modifications for the seeding and growth of the cells. A simple drop plating method (Protocol 3B) where cells are seeded in the form of small-volume discrete droplets onto dishes sequentially coated with PLL and laminin is recommended to increase cell recovery and expedite cell attachment. After testing the effect of various growth factors on hSC proliferation (Monje et al., 2018), we recommend using medium supplemented with serum, heregulin, and forskolin from the onset, as this formulation induces hSC mitogenesis as soon as 1–2 days post plating and maintains the hSCs in an active proliferative state during the initial stages (Protocol 3C) and for several passages. It is expected that the cells plated under these conditions reach confluence within 5–10 days depending on the initial plating density.
Enzymatic dissociation of pre-cultured tissues
Prepare the pre-degenerated fascicles for dissociation by cutting the fibers into smaller 1 cm segments, if needed. Before proceeding with the next steps, inspect the tissue under a dissecting or phase contrast microscope to ensure no connective tissue or debris are attached (Figure 8A).
Figure 8. Isolation, plating, and analysis of human Schwann cells (hSC)-enriched suspensions. (A–D) Enzymatic dissociation of nerve fascicles. (E) Drop plating on a Poly-L-lysine (PLL)-laminin substrate. (F and G) Testing for cell viability and myelin contamination right after tissue digestion. The total cell nuclei were labeled with Hoechst (blue), dead cells were labeled with PI (red), and myelin was labeled with Fluoromyelin (FM, green).
Remove the culture medium and wash the pre-degenerated fibers once with a large volume of DMEM before adding the 1× enzymatic solution (see Recipe 5). For instance, use 1.5 mL of enzymatic solution to digest the fibers harvested from a 1 cm sural nerve biopsy placed in a 3 cm culture dish. Scale up the volume of enzymatic cocktail to match the volume of tissue and the dimensions of the dish to ensure that the tissue is fully covered by enzymatic solution.
Transfer the fascicles to an incubator set at 37 °C and 8% of CO2 for overnight dissociation (15–18 h) without agitation. Use phase contrast microscopy to monitor the progression of the digestion at regular time points (Figure 8B and 8C) (see Note a).
The next day, transfer the digested explants into a 15 mL snap-cap tube (round-bottom tube) containing 5–10 mL of DMEM medium with 40% FBS to rapidly neutralize the action of the enzymes. Rinse the dish and collect the explants using LP medium (see Recipe 2) until there is nothing left in the dish.
Centrifuge the tissues at 200× g for 8 min at 4 °C to collect the particulate material. Discard the enzymatic solution.
Remove the supernatant manually using a sterile transfer pipette and add 5 mL of LP medium. Alternatively, aspiration can be used with care to avoid disturbing the pellet.
Gently resuspend the cells by pipetting up and down with a wide end bore glass pipette until the tissue disintegrates as observed by visual inspection. Then, add LP medium to a final volume of 15 mL. This suspension should look as shown in Figure 8D. Do not overdo the mechanical disintegration, as excessive passing of the cells through a pipette can increase cell death. Consider that large undigested tissues, if still present, can be removed in subsequent steps.
Repeat step 5 and 6 to create a suspension ready for plating and remove traces of the enzymatic solution. Resuspend the cell pellet gently in the desired volume of HP medium using a narrow-end glass transfer pipette. The cells are ready to be plated as described in Protocol 3B.
(Optional) Set aside a 50–100 µL aliquot of this cell suspension to determine cell counts, viability, and myelin content using nucleic acid- and myelin-specific fluorochromes (Figure 11). For live/dead nuclear labeling, use a combination of the following: (1) Syto-24 (total cells, green) and PI (dead cells, red) (not shown); or (2) Hoescht (total cells, blue under UV light) and PI (red) (Figure 8F). For myelin staining, use Fluoromyelin (red or green) or FM 4-64FX (red) alone or in combination with a nuclear stain (Figure 8G).
Plating of the initial cell suspensions
Confirm under a phase contrast microscope that a refined suspension containing single cells (or small clumps) was obtained in the prior step (Figure 8F and 8G) (see Note b).
Plate the cell suspensions in small drops directly on the surface of a PLL-laminin-coated 100 mm dish. Do so by placing discrete 30–50 µL drops in a regular pattern, as shown in Figure 2 and Figure 8E. The plating density may be difficult to estimate at this stage. For a reference, plate 106 cells in a 100 mm dish considering that one dish can accommodate 20–30 drops (see Note c).
(Optional) Set aside a 100–200 µL aliquot of this cell suspension for plating droplets in a PLL-laminin-coated chamber-slide or multi-well dish. These adherent cells are to be used for quality control analysis of cell viability, purity, and myelin content (Figures 10 and 11).
Carefully transfer the culture dishes to a CO2 incubator for at least 15–18 h (see Note d).
Culture of primary hSCs
The following day, use phase contrast microscopy to confirm that the cells have attached properly. Gently and smoothly fill each 100 mm dish with HP medium to cover the surface of the dish (8–10 mL) without disturbing the cells.
Proceed with normal medium changes by refreshing the HP medium every three days on average. Change the medium slowly without creating a flow of liquid onto the cells as they detach easily at this stage.
(Optional) Include additional washes with 37 °C DMEM or L15 to remove extracellular myelin debris and dead cells, preferably after the second day of plating (Figure 9A and 9B).
Once the cells reach a high enough density (or confluency), they can be used in quality control assessments (Figure 10) or experimentation. Confluent cultures can be considered passage-zero (P0) primary hSCs (see Note e).
Figure 9. Growth of a drop-plated human Schwann cell (hSC) culture at passage-zero. (A–C) Cell attachment and expansion right after drop plating. Observing abundant myelin debris in the medium (A) and inside the cells (Figure 10) is a normal and expected feature of these early cultures [within three days in vitro (DIV)]. Loosely attached myelin granules (A, extracellular myelin) can be easily removed with media changes or additional washes (B). (D–F) Appearance of the monolayer outgrowth of a nearly confluent hSC culture at 5 DIV. Images were taken within the periphery (D) and the center (E, F) of an individual drop. Cells migrating out of the area initially circumscribed by the drop are denoted by the dotted line and the arrow (D). Cell alignment (E, F) may or may not be evident at this stage.
Figure 10. Analysis of a drop-plated human Schwann cell (hSC) culture at passage-zero. (A, B) Phase contrast microscopy showing cell morphology and intracellular myelin content in the cytoplasm of hSCs. Myelin persists for several days. However, granules of larger diameter (B) are no longer appreciated five to seven days post plating due to intracellular degradation (C). (F) Fluorescence microscopy revealing myelin-loaded hSCs using a myelin stain (FM 4-64FX, Myelin, red) in combination with O4 immunolabeling. The hSCs are NGFR+ (see Figure 2) and O4+ (green, E and F) at this stage [5 to 7 days in vitro (DIV)]. O4 staining declines with prolonged culture whereas NGFR persists (Monje et al., 2018). Nuclei were labeled with DAPI (blue).
Notes:
Check the progression of tissue dissociation by phase contrast microscopy 1–2 h after adding the enzymatic cocktail to appreciate the action of the enzymes (Figure 8B and 8C). It is possible to observe individual cells in suspension being released into the medium quite early after the addition of the enzymes. However, the smooth and progressive disassembly of adult nerve tissue is generally achieved after prolonged (15–18 h) incubation. Introducing mechanical steps or agitation is not recommended because it can negatively affect cell viability.
The preparations that result from enzymatic dissociation consist of single cells and cells in small clumps. A variable proportion of undigested tissues, dead cells, and myelin debris of varied granularities are also expected. To prevent further stress on the cells, proceed with the plating even if cellular aggregates and undigested fibers are observed.
Plating the suspensions as small droplets allows SCs to rapidly attach to the surface and separate easily from floating myelin debris (Andersen and Monje, 2018). This method is preferred over plating the cells directly in a large volume of medium (traditional method).
Maintain the cells inside a humidified CO2 incubator set at 37 °C with 8%–9% CO2. Monitor the cultures within 3 h post plating only to verify prompt adhesion. Do not disturb the cultures otherwise. The cells adhere and extend processes quickly onto a PLL-laminin substrate. The morphology of the cells is variable in the initial stages and detach easily if disturbed. Cell yields and purity usually differ in independent isolation experiments.
These cells can be replated and sub-cultured for several passages. Accompanying papers provide information on managing and analyzing both primary and expanded hSCs (Monje, 2023a and 2023b).
Recommendations and troubleshooting
Performing daily microscopic observations of the cultured tissues and cells and introducing proper controls are essential throughout the protocol. Primary hSCs are variable and delicate cultures until they become established. For this reason, this section includes useful conceptual and technical information to help investigators make rational use of the materials and procedures as well as trace and correct problems as they arise (Figure 11).
Figure 11. Suggested controls at each step during the culturing of nerve tissues and cells. The diagram depicts an action plan for performing sterility controls (left) and other assessments (right) to rule out microbial contamination and confirm the purity of the primary cell cultures, respectively.
Tissue procurement and use. The protocols for tissue harvesting, transfer, and use in laboratory research must be reviewed and approved by the appropriate institutional committees. Tissues are often furnished to researchers by local medical facilities, organ procurement centers, and tissue banks under proper authorization and transfer agreements that follow institutional guidelines for protecting donor-relevant information. Most biospecimens are deidentified before transfer. However, it is recommended that the recipient laboratory is given access to pertinent information to assess factors impacting the hSC cultures and the operators. Such information may include the date and time of tissue extraction, patients’ demographics, medical condition (or cause of death), and the presence of communicable diseases. Ideally, this information should be made available prior to unpacking the tissue in the laboratory to allow the investigative team to make an informed decision on the suitability of the tissues for cell culture, and/or the need to take additional precautions. Evidence of cancer, peripheral neuropathy, or chronic disease affecting the nerve, such as insulin-dependent diabetes, may be considered an exclusion factor for the derivation of hSC cultures unless the intention is to investigate the properties of hSCs under these conditions. The investigators must decide if they will process tissues from patients who have tested positive for blood-borne pathogens or other communicable diseases. We routinely exclude tissues if we become aware that the donors have tested positive for HIV, hepatitis B, or hepatitis C. Samples from donors that tested positive for other viruses may be processed for cell culturing after being quarantined and labeled appropriately.
Ideally, the culture of the fascicles should be initiated within 24 h following nerve harvesting or 24 h postmortem time. However, the culturing can be delayed for several days if the tissues are stored appropriately at 4 °C. A simple live/dead test (Figures 8 and 11) can suffice to assess both cell content and viability prior to tissue processing. It has been shown that short intervals (< 1 week) of cold storage do not substantially impair the viability and function of SCs obtained from adult human and rat nerves. If the cold storage period is extended to three weeks, the viability declines; however, some cells survive and can be isolated for culturing (Levi et al., 1994). In general, we suggest not to exclude tissues based on time of collection from the body (for autopsy specimens) or the cold storage period (for fresh or autopsy specimens) without empirical verification of cell viability.
Biosafety. Importantly, all human tissues and cells should be managed under the assumption that unknown pathogens are present. Universal precautions should be used, and personnel should be educated in the handling of biohazardous materials. Operators should follow institutional guidelines to work at the appropriate biosafety level and limit occupational exposure to biohazardous materials. Human cells and tissues should be managed inside a certified BL2 biosafety cabinet. Procedures that minimize the creation of aerosols and the transport of potentially hazardous materials should be implemented. Fluids and materials in contact with the human cells/tissues should be labeled, inactivated by autoclaving, and disposed of as biohazards.
Sterility. Implement best practices in the handling of the tissues and cells to maintain sterility. Examine the cultures daily by phase contrast microscopy using low (4×–10×) and high (40×) power objectives to assess the overall health and progression of the cultures and to rule out gross microbial contamination. Routine sterility testing is recommended throughout the main steps but mostly at the onset of culturing (Figure 11). Microbial contamination of the source tissue prior to arrival to the laboratory and/or during the dissection procedure is a likely cause of contamination of the cell cultures. Once the cultures become contaminated, they should be discarded. We have a simple method in place for microbial testing using liquid cultures in Luria-Bertani (LB) broth. For this, we inoculate 10 mL of LB medium with approximately 50 µL aliquots of the cell/tissue supernatants from the media or buffers used for tissue storage, dissection, pre-degeneration, and culturing along with aliquots of the unused media stocks. These LB cultures, grown at 37 °C for at least five days, can allow the source of microbial contamination to be traced before it arises in the cell/tissue cultures. This method is useful but has limitations. For more reliable testing of banked cell cultures, we outsource samples to an analytical lab that conducts testing for the presence of bacteria (aerobic, anaerobic), fungi, and mycoplasma (culture-based methods). We do not usually conduct routine mycoplasma testing in primary cultures used in basic in vitro cell research unless the cells are banked. Detection by fluorescence microscopy and PCR (commercial kits) are useful for such purposes.
Culture media formulation and use. Adding heregulin and forskolin is recommended but not essential for the growth and survival of primary hSCs. Medium containing FBS at 10%–15% can be used for plating the cell suspensions when chemical mitogens are unavailable or undesirable experimentally, with the caveat that these conditions do not lead to substantial hSC proliferation; instead, they exacerbate the propagation of fibroblasts. Researchers may optimize their culture media formulation. Serum-free or low-serum media have been used by other groups (Haastert et al., 2007; Aghayan et al., 2012; Weiss et al., 2018). We do not generally recommend the removal of serum unless necessary experimentally.
Regarding the feeding schedule, we recommend feeding twice a week on average using 10 mL of medium for a 100 mm dish, or the volume that matches the size of the dish. To feed the cultures, remove the medium carefully from the side and gently add new medium (previously warmed at a 37 °C water bath) to the side to prevent turbulence on the surface, as this can lead to cell detachment. To prevent alkalization of the media, do not leave the cells and the culture media stocks open in the biosafety cabinet for long periods. The viability of cultured hSCs is impaired when exposed to alkaline media or buffers, even for short periods. Use properly calibrated solutions and follow best methodological practices to prevent sudden pH changes. If available, use phenol red–containing media to visually monitor and correct fluctuations in pH. One consideration is that phenol red can have potential estrogenic effects (Berthois et al., 1986) and redox activity (Morgan et al., 2019) and may be avoided in cell culture. Nevertheless, our empirical observations have shown that hSCs can be cultured equally well in the presence or absence of phenol red.
Substrate requirements. Freshly plated and passage-zero hSC cultures are prone to detachment, which can be reduced by (re)plating the hSCs on PLL-laminin. Prepare PLL-laminin-coated dishes as suggested in this protocol and use them preferably within 1–3 days post coating. We cannot recommend commercial dishes that provide a good substrate for hSCs (empirical data); however, other substrates (or matrices) may be used if hSC adhesion and proliferation are confirmed empirically (Vleggeert-Lankamp et al., 2004).
Myelin and other impurities. The myelin content in intact and pre-degenerated nerves varies substantially from donor to donor and even from fascicle to fascicle within a nerve (Figure 4E and 4F). Use HP rather than LP medium for more efficient myelin clearance by hSCs within the fascicle explants (pre-degeneration) and after plating (Casella et al., 1996). While the presence of myelin can be reduced over time, it is rarely eliminated during the pre-degeneration phase, even with prolonged incubation (>1 month). This is a distinctive feature of nerves obtained from humans (Aparicio and Monje, unpublished).
Myelin interferes with the visualization of cells after enzymatic digestion of the fascicles. Notice that standard phase contrast microscopy is inaccurate for this purpose. A practical way to estimate the number of cells in freshly isolated preparations is to stain them with vital fluorophores and count fluorescent nuclei instead of whole cells using a Neubauer hemocytometer or an appropriate automated device. The discrimination between live/dead cells is best accomplished by co-staining the preparations with a combination of green (e.g., Syto-24) and red (e.g., PI) fluorophores to prevent the interference of myelin autofluorescence in the UV channel. The Hoescht/PI combination is useful in plated preparations where the myelin floats (Figure 8F and 8G).
In addition to myelin debris, the initial cell suspensions prepared directly from tissues contain abundant dead cells, membranous vesicles, and other impurities intermixed with the cells. Passage-zero hSC cultures may contain myelin debris floating in the medium or loosely attached to the surface of the cells (Figure 9). A proportion of the hSCs contain cytoplasmic myelin granules of various sizes that are degraded quickly after the hSCs are plated on a substrate (Figure 9A–9C, Figure 10A–10C). Intracellular and extracellular myelin from the original tissue is drastically reduced approximately one week post plating. However, smaller-sized myelin granules and lipid droplets may persist in the cytoplasm of hSCs for some time. The extent of myelin contamination in each hSC preparation is variable, as determined by the type of nerve and the level of pre-degeneration. Nevertheless, the presence of myelin does not impair the viability or the proliferation of the cultured hSCs. A trained experimentalist can easily discriminate myelin contamination from microbial growth. If this discrimination is doubtful, perform appropriate sterility tests.
Management of tissues and cultured cells. The enzymatic dissociation of the nerve fibers is the most important step that affects cell yields and viability. The temporal course of dissociation must be controlled for each preparation, particularly if mature nerves with fully developed connective tissue layers are used. Increasing the concentration of the enzymes and teasing the fibers with fine forceps, as described in Andersen and Monje (2018) are two distinct ways to disaggregate the tissue more expeditiously with the caveat of increased cell death. Experimenters should always optimize the dissociation step by testing the viability of the cells throughout the course of digestion (Figure 8B and 8C).
Primary hSCs can be plated in their final dishes for direct experimentation. However, these cells proliferate at a high rate in the presence of HP medium (with a duplication time of < 2 days) and the appearance of the cultures changes daily. If cells are plated as drops, we recommend using the cells in experimentation once they reach confluency within the surface area initially delimited by each individual drop (Figure 9D and 9E). Over-confluent cultures may show areas with frequent SC clusters. Use the hSC cultures as soon as signs of migration and clustering in local areas become evident. If the drops are plated initially in a 100 mm plate, these cells can be re-plated into a new 100 mm plate, or the equivalent surface area of a multi-well dish, for recovery. One limitation of our method is the variable cell yields per isolation experiment (see Donor variability). hSC purity is usually high (>80%) under these conditions.
Identification and analysis of cultured cells. Passage-zero hSCs are highly active and migratory. Even though normal hSCs tend to grow in monolayer, the primary cells overlap their processes and rarely align with one another to form fingerprint-like patterns on the surface, as is expected of established SC cultures from rodents and humans. SCs and fibroblasts are not easily distinguishable at this stage. Therefore, researchers should use immunological methods to confirm the composition of the cultures. Additional relevant controls are to estimate the proportion of live/dead cells (Figure 8) and the content of myelin debris (Figure 10) by fluorescence microscopy. We have previously reported detailed methods for purity and viability assessments (Ravelo et al., 2018; Peng et al., 2020); thus, here we have simply provided examples of typical results in Figure 2 (NGFR staining), Figure 10 (O4 staining), and Figure 8 (viability). Researchers may consider using image analysis software such as ImageJ to quantitatively assess purity, viability, and myelin contamination from adherent cells or cells in suspension. If intracellular myelin is confirmed in a proportion of the cells, it can be used as a criterion for hSC identification, as myelin engulfment is a specific property of the SCs. Fibroblasts do not normally engulf myelin fragments, and macrophages are rarely seen under these experimental conditions.
Minimal manipulation. Primary cells are considered more reliable model systems than established cultures or cell lines because they more closely reflect the phenotype present in the original tissue. However, maintaining this phenotype in vitro is a challenge. It has been shown that the characteristics of hSCs change with passaging, but the extent to which in vitro–cultured hSCs diverge from those in the nerves is unknown thus far. Therefore, limit the time of exposure to mitogenic factors and the rounds of expansion if the expectation is to generate a product with as little manipulation as possible. The protocols described herein have favored consistency in cell isolation and growth over time in vitro and exposure to mitogens. Other protocols emphasize minimal manipulation to produce hSCs from normal nerves and schwannomas (Dilwali et al., 2014).
Donor variability. One general caveat of patient-derived cultures is that the quality of the populations varies from donor to donor and batch to batch for reasons that are seldom clear to the investigators. This variability may be difficult to control but cannot be ignored when evaluating outcomes from in vitro and in vivo experiments. The heterogeneity of hSC cultures and the divergence from the cells of origin may be problematic if the cultured cells are intended to represent endogenous hSCs from patients. Factors as diverse as the donors' genetic background and medical history, the conditions of tissue procurement, and cell type selection in culture may contribute to batch variability [reviewed in Monje (2020)]. Having accurate records of the history of the tissues and the cells may help explain variation. If possible, derive cultures from different donors and use those to simultaneously compare the responses of cells from different batches before making generalizable conclusions on the effect of experimental variables.
Recipes
Dissection medium (DM)
Reagent Final concentration Amount
L15 medium n/a 499.5 mL
Gentamycin (1,000×) 1× 0.5 mL
Total 500 mL
Low proliferation medium (LP)
Reagent [Stock concentration] Final concentration Amount
DMEM (or DMEM/F12) n/a 445 mL
FBS (100%) 10% 50 mL
GlutaMAX (100×) 1× 5 mL
Gentamycin (1,000×) 1× 0.5 mL
Total n/a 500 mL
Note: Adjust the pH of this and other culture media to be 7 or lower (up to 6), sterilize by filtering through a 0.22 µm filter, and maintain at 4 °C.
High proliferation medium (HP)
Reagent [Stock concentration] Final concentration Amount
Low proliferation medium n/a 500 mL
Heregulin-β1177-244 (25 µM) 10 nM 200 µL
Forskolin (15 mM) 2 µM 69.25 µL
Total 500 mL
Note: Use this medium preferably within a month of preparation and re-filter if needed. Do not freeze.
Laminin coating solution
Reagent [Stock concentration] Final concentration Amount
Laminin (1 mg/mL) 15.6 µg/µL 78 µL
DPBS n/a 5 mL
Total 5 mL
Note: This recipe coats one 100 mm cell culture dish (growth area = 78 cm2). We use 1 µg of laminin/cm2 of coating surface for the growth of hSCs. See our paper for the PLL-laminin coating protocol (Ravelo et al., 2018).
10× enzymatic solution
Reagent [Stock concentration] Final concentration Amount
Dispase II (powder) 25 mg/mL 125 mg
Type I collagenase (powder) 5 mg/mL 25 mg
DMEM n/a 5 mL
Total 5 mL
Note: Prepare the 10× enzymatic solution as indicated above and sterilize by filtration using a 0.22 µm filter. Aliquot and store this solution at -80 °C for up to one year. To prepare the 1× working solution thaw the 10× aliquots on ice and use DMEM to bring the concentration of enzymes to 1×. Do not add antibiotics.
Acknowledgments
We recognize Ketty Bacallao and Natalia Andersen for expert technical assistance, and Patrick Wood and Mary Bunge for their valuable mentoring. Michael Murphy, Kristen Wanczyk, Jamie Bradbury, Greg Gerhardt, Craig van Horne, and George Quintero contributed tissue specimens via institutional Materials Transfer Agreements. Former colleagues from the University of Miami (UM), Yelena Pressman, Linda White, Anna Gomez, Gagani Athauda, Peggy Bates, Aisha Khan, Adriana Brooks, Risset Silvera, Maxwell Donaldson, and James Guest are kindly acknowledged for their valuable input over many years of shared work. The protocols described here were developed and optimized during a >10-year period supported by funding (to P.V.M.) from the National Institutes of Health NIH-NINDS (NS084326), the Craig H Neilsen Foundation, The Miami Project to Cure Paralysis and the Buoniconti Foundation from the University of Miami, and the Indiana State Department of Health (grants 33997 and 43547). G.I.A received support from the International Society for Neurochemistry-Committee for Aid and Education in Neurochemistry (ISN-CAEN), the Fulbright, Bunge & Born, Williams Foundations (program 2020–2021), and CONICET-Argentina. P.V.M. and G.I.A. received support from the Department of Neurosurgery at the University of Kentucky. The contents of this article are the authors' responsibility and do not necessarily represent the official views of the funding agencies. We are grateful to Lingxiao Deng for fruitful scientific discussions and lab support. We thank Beth Ansel for critical review, Valeria Nogueira for illustrations, and Louise Pay for English editing. We are greatly indebted to the generosity of the anonymous patients and their families for donating tissues for research.
Competing interests
P.V.M. is the founder of GliaBio LLC, a consulting company focusing on glial cell research. The authors declare that the research described here was conducted without commercial or financial relationships that could be construed as a potential conflict of interest.
Author’s contributions: P.V.M.: conceptualization, experimentation, data acquisition and analysis, manuscript writing, figure preparation, funding. G.I.A.: experimentation, data acquisition, manuscript writing, figure preparation.
Ethical considerations
Experimentation with human tissues and cells was deemed to constitute non-human subjects research by the Human Subjects Research Offices of the University of Miami (2003–2018) and Indiana University (2018–2022). Procedures were further reviewed and approved by the Institutional Biosafety Committee of Indiana University. Human tissues were acquired from multiple sources, as follows. Michael Murphy and Jamie Bradbury (Indiana University) contributed deidentified nerves and ganglia from surgical remains. Craig van Horne (University of Kentucky) contributed deidentified sural nerve biopsies from clinical trial participants. The National Disease Research Interchange (NDRI) provided cadaveric sural and sciatic nerves under an institutional agreement with P.V.M while affiliated to U.M. Protocols for tissue procurement and sharing by the provider scientists were approved by the respective institutional review boards. Experiments were conducted using non-pathological tissues from males and females as made available from the provider scientists or organ procurement centers without regard to the gender or age of the donors. Requests for information and materials (cells, antibodies, and other materials) should be directed to P.V.M. Investigators are encouraged to contact P.V.M. for feedback on the reported protocols.
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The authors used this protocol in eLIFE Oct 2022
Abstract
Determining the oligomeric state of membrane proteins is critical for understanding their function. However, traditional ex situ methods like clear native gel electrophoresis can disrupt protein subunit interactions during sample preparation. In situ methods such as stepwise photobleaching have limitations due to high expression levels and limitations of optical resolution in microscopy. Super-resolution microscopy techniques such as single-molecule localization microscopy (SMLM) have the potential to overcome these limitations, but the stochastic nature of signals can lead to miscounting due to over-expression, background noise, and temporal separation of signals. Additionally, this technique has limited application due to the limited selection of fluorescent labels and the demanding control of laser power. To address these issues, we developed a dual color colocalization (DCC) strategy that offers higher tolerance to background noise and simplifies data acquisition and processing for high-throughput and reliable counting. The DCC strategy was used to determine the oligomeric states of membrane proteins of the SLC17 and SLC26 family with SMLM, providing a robust and efficient method for studying protein interactions.
Graphical overview
(A) Illustration of the principle for determining the oligomeric state of protein complexes with dual color colocalization–single-molecule localization microscopy (DCC-SMLM). In the inset, as an example, a dimeric protein (brown) is labeled with a marker (M) and an indicator fluorescent protein (F) on each of its two subunits. The overall probability of detecting the dimer with SMLM, as denoted by R, the colocalization ratio, is equal to the ratio of the number of colocalized marker and indicator clusters (NMF) to that of the marker clusters (NM). The plot shows the linear relationship of the oligomeric state (n) vs. the natural logarithm of 1 subtracted by the colocalization ratio, supplemented by the equation of the fitting curve, in which p denotes the recall rate of the indicator fluorescent protein (F). (B) The workflow diagram shows the procedures of DCC-SMLM (Locs: localizations; COM: coefficient of mismatch; LCA: lateral chromatic aberration).
Keywords: Single-molecule localization microscopy Super resolution microscopy Quaternary structure Membrane proteins PAmCherry Photoactivated localization microscopy
Background
The composition of the quaternary structure of a protein tells us how this protein is operating and is therefore critical for the understanding of the molecular mechanisms behind their function. The experimental determination of protein oligomeric states is nontrivial. Several methods or strategies have been developed in the past decades, such as native gel electrophoresis (Schägger and von Jagow, 1991), stepwise photobleaching (Ulbrich and Isacoff, 2007), and single-molecule localization microscopy (SMLM) (Sengupta et al., 2011; Lee et al., 2012). Ex situ methods, such as native gel electrophoresis, rely on the extraction and purification of proteins for further investigation (Schägger and von Jagow, 1991). The detergents and mechanical forces exerted on protein complexes during these steps can, however, result in a loss of weaker interactions between subunits, creating a bias in the observed results. In response to these limitations, in situ methods, primarily centered on fluorescence microscopy, have been increasingly used in the past years. Often, this is in the form of stepwise photobleaching, where protein subunits are labeled with fluorophores and subsequently bleached by strong excitation (Ulbrich and Isacoff, 2007). This results in a stepwise loss of fluorescence for every bleached fluorophore. Due to the spatial limitations of light microscopy, however, this method only works if the density of expressed proteins is low. This mostly limits the application of this method to the use of Xenopus laevis oocytes, which are not a faithful representation of the cytosolic environment in mammalian cells.
As a rather elegant strategy to implement super resolution microscopy technique, SMLM temporally separates the emissions from individual molecules to avoid overlapping signals from multiple emitters in the same diffraction limited spot. Fitting of these isolated signals can increase the resolution by up to tenfold. To achieve homogeneous labeling, the fluorophores are usually genetically encoded photoactivatable fluorescent proteins fused to the protein of interest. Thus, this type of SMLM is also called photoactivated localization microscopy (Betzig et al., 2006).
Despite the achievable resolution and genetic labeling, it is still not possible to count the number of proteins within a sample by the number of fluorescent signals observed. For various reasons, not all fluorescent proteins will generate a signal during experiments (i.e., the recall rate is lower than 1). The distribution of the number of counts per protein complex is described by a binomial distribution, defined by the recall rate (Nan et al., 2013; Durisic et al., 2014). This strategy faces numerous challenges, primarily in that it demands a high recall rate (at least 0.5) of the fluorescent proteins. There are only a few options of suitable fluorescent proteins reported previously [e.g., PAmCherry (Durisic et al., 2014) and mEos3.2 (Zhang et al., 2012)]. Emission events from each fluorescent protein can extend over several frames and may be discontinuous. It is therefore imperative to develop algorithms that assign emission events to individual proteins within the complex. Furthermore, the binomial distributions of signals are often spuriously affected by a significant amount of background signals that are almost indistinguishable from true signals. We recently published a novel strategy, dual color colocalization (DCC) (Tan et al., 2022), circumventing these problems, as illustrated in the Graphical overview. The DCC strategy uses two covalently linked fluorescent proteins, serving as the marker (M) and the indicator (F), fused to the protein of interest. The marker is used to select the fraction of protein complexes that will be used for counting, while the indicator will be used to determine the oligomeric state (n) of the protein complex. This is done by experimental determination of the detection probability (R) of the protein complex, as given by the colocalization of the two fluorescent proteins:
(Equation 1)
NMF is the number of protein complexes detected via both M and F, i.e., the colocalized protein complexes. NM is the overall number of clusters that show fluorescence of M, regardless of whether they are colocalized with F or not, and p is the recall rate of the indicator F. In this way, we disregard the complicated temporal separation (except to improve resolution based on SMLM) and assignment of emission events to individual proteins. This simplifies the imaging procedure and the data processing and builds a direct connection between the detection probability of the protein complex and its oligomeric state (Equation 1).
Since our method relies on a cumulative probability instead of the probability densities of multiple detected states in a binomial distribution, the DCC strategy does not demand a high recall rate, thereby broadening the spectrum of usable fluorophores. For the marker protein, we choose the bright mVenus, given that the corresponding filter setting leads to a very low noise level, which is essential for the faithfulness of the result.
Overall, this dual-color strategy greatly increases the signal-to-noise ratio in comparison with the single-color counting strategy. As we considered that the two adjacently linked fluorescent proteins may interfere with one another, or be truncated simultaneously, we elaborated our model by introducing another parameter, the coefficient of modification denoted by m, resulting in the following formula:
(Equation 2)
In this protocol, we present the best practices that we established during the development of our method. We used DCC to determine the in situ oligomeric states of the vGlut family as monomers and SLC26 family as dimers. The DCC strategy has been proven to be an efficient improvement of the conventional strategy using SMLM to determine protein oligomeric state.
Materials and reagents
Filter for solutions, 0.22 μm (e.g., Corning® 1,000 mL Bottle Top Vacuum Filter, catalog number: 431174)
Sterile cell culture Petri dishes [Sigma-Aldrich, catalog number: D7804 (35 mm) and catalog number: Z755923 (100 mm)]
Low background coverslips, 25 mm, No. 1 (VWR, catalog number: 631-1584)
Microscope slides, 75 mm × 25 mm, plain (VWR, catalog number: 48300-026)
HEK293T cells (Sigma-Aldrich, catalog number: 96121229-1VL)
Coplin slide staining jar (glass) for 75 mm × 25 mm slides (DWK Life Sciences, catalog number: UX-48585-20), used for cleaning and storage of coverslips
HEK293T cell culture medium made from:
DMEM (Thermo Fisher Scientific, catalog number: 10564011)
Supplemented with 50 U/mL penicillin-streptomycin (Thermo Fisher Scientific, catalog number: 15140122)
Supplemented with 10% fetal bovine serum (FBS) (Thermo Fisher Scientific, catalog number: 16140071)
Trypsin-EDTA, 0.05% (Thermo Fisher Scientific, catalog number: 25300062)
Lipofectamine 2000 (Life Technologies, catalog number: 11668019)
OptiMEM for cell transfection (Thermo Fisher Scientific, catalog number: 31985088)
Hydrogen peroxide, 30%, stabilized (Merck, catalog number: 108597)
Methanol, gradient grade for liquid chromatography (Merck, catalog number: MX0486)
Sulfuric acid, 95%–97% (Sigma-Aldrich, catalog number: 1.00731)
Sodium hydroxide (NaOH) pellets (Merck, catalog number: 106469)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S5886)
Disodium phosphate (Na2HPO4) (Sigma-Aldrich, catalog number: S5136)
Potassium dihydrogen phosphate (KH2PO4) (Sigma-Aldrich, catalog number: P5655)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P5405)
TetraSpeck fluorescent bead solution, 100 nm diameter (Thermo Fisher Scientific, catalog number: T7279)
pH indicator paper (broad range from acidic to basic; no particular supplier is required)
Phosphate-buffered saline (PBS) (see Recipes)
Equipment
Cell culture incubator with a constant temperature at 37 °C and 5% CO2
Laminar hood with an aspiration pump and a microscope for standard biosafety level 1 cell culture operations
Personal safety equipment for handling Piranha solution, including lab coat, acid-resistant gloves, and protective goggles
Glassware for preparing buffers and Piranha solution, including 100 mL and 1 L glass beakers, and 10 and 20 mL transfer pipettes.
Hot plate with magnetic stirrer for preparing solutions
Chemical fume hood for preparing and using Piranha solution
Total internal reflection fluorescence (TIRF) microscope capable of recording SMLM images in at least two separate color channels (plus activation 405 nm laser if photoactivatable proteins such as PAmCherry are used). Please consult with your local experts, if necessary, to determine the suitability of any available microscope. As a guide, the microscope we used in our original publication (Tan et al., 2022) was outfitted with the following components (Tang et al., 2015):
An ApoN 60× Oil TIRF objective (NA 1.49; Olympus)
Three laser lines:
i. 405 nm diode laser (Cube 405-100C; Coherent) for the activation of PAmCherry
ii. Argon-ion laser, tuned for 514 nm (Coherent, catalog number: Innova 70C) for imaging of mVenus
iii. 561 nm diode laser (Coherent, catalog number: Sapphire 561-200 CDRH-CP) for imaging of PAmCherry
An acousto-optic tunable filter (AOTF nC-VIS-TN 1001; AA Opto-Electronic, Orsay, France) to control the intensity of the 514 and 561 nm laser beams reaching the objective
Two EMCCD cameras (Andor iXon DU897E; Andor, Belfast, UK; one for each color channel) cooled down to -75 °C with liquid nitrogen to record emissions from mVenus and PAmCherry, respectively, using a pixel resolution of 512 × 512 pixels
Multi-channel dichroic mirror (e.g., 442/514/561 nm lasers BrightLine® triple-edge laser-flat dichroic beam splitter, catalog number: Di01-R442/514/561-25x36, Semrock)
Appropriate emission filters [e.g., 485/537/627 nm BrightLine® triple-band bandpass filter (Semrock, catalog number: FF01-485/537/627-25) and 525/50 nm BrightLine® single-band bandpass filter (Semrock, catalog number: FF03-525/50-25) for mVenus; 609/57 nm BrightLine® single-band bandpass filter (Semrock, catalog number: FF01-609/57-25) for PAmCherry]
A recording chamber that holds the coverslip in place without movement during the recording. Furthermore, the chamber must allow for filling with 1–2 mL of imaging buffer (i.e., PBS) on top. (This can be custom-built, or commercially available chambers such as the MS-518 series by ALA Scientific or the 20 series chambers by Warner Instruments can be used.)
Software
Data acquisition system for microscope control and imaging. We used a custom-built program for the original publication that is not available for other microscope systems. In general, however, suitable microscopes come with appropriate software installed. Please consult with your local experts for training in using the microscope and control software, if necessary
Fiji (https://imagej.net/ij/download.html)
Python 3 with Jupyter Notebook (https://jupyter.org/), Pandas (https://pandas.pydata.org/), NumPy (https://numpy.org/), SciPy (https://scipy.org/), Matplotlib (https://matplotlib.org/), and Pillow (https://pillow.readthedocs.io/en/stable/index.html). To use Python and these packages, we recommend using Anaconda (https://www.anaconda.com/) to manage the packages and solve dependencies
SNSMIL (http://english.nanoctr.cas.cn/dai/software/201505/t20150504_146857.html (Tang et al., 2015) or SMAP (https://github.com/jries/SMAP (Ries, 2020) for localization extraction
DCC-SMLM library (https://github.com/GabStoelting/DCC-SMLM (Tan et al., 2022)
Procedure
Sample preparation
Preparation of plasmids
Clone the cDNA of the proteins of interest (POI) and the standards into the vector plasmid pcDNA3 or pcDNA5/FRT/TO. The proteins must be labeled with two fluorophores—in our case, mVenus and PAmCherry. We chose to fuse the coding sequence of mVenus and PAmCherry to the C-terminus of the coding region of the POI or the protein standard to simplify the procedure and because we had prior knowledge that the function of the proteins would remain largely intact. C-terminal linkage of the fluorophores, however, is not a requirement of our method. Fluorophores may be linked to the N-terminus or embedded within the protein itself. It may be necessary to perform functional experiments to assess whether the function is impaired by linkage to fluorescent proteins such as mVenus and PAmCherry. If possible, it may be possible to link one fluorophore to the N-terminus and the other fluorophore to the C-terminus of the protein. This would eliminate the need to consider unwanted truncation of the fluorophores and thus the need for the parameter m (Equation 2).
We used barttin as the monomeric, the ClC-K and ClC-2 as the dimeric, EAAT2 as the trimeric, and Kcnj2 as the tetrameric standards. However, other plasma membrane proteins can also be tested and used as the standard. Some important criteria for choosing the standard proteins include: (1) the protein should be predominantly in the plasma membrane; (2) the protein should be expressed well in the chosen model cell line; and (3) the protein should have a mostly homogeneous oligomeric state, but not aggregate with others into super complexes or dissociate into individual subunits. We used a linker between the POI or standard protein and the fluorescent protein tag consisting of a flexible hydrophilic sequence with 10–30 amino acid residues. The linker should not be too long since it may increase the chance of protein cleavage. For more details about the standard proteins and the linker sequences, please refer to Tan et al. (2022).
Preparation of coverslips
Place the coverslips (25 mm) in the Coplin slide staining jar, standing vertically and separate from each other, so that both sides can be immersed in Piranha solution (see below).
Determine the volume of Piranha solution that is needed to completely immerse the coverslips in the Coplin jar. With the Coplin jar and coverslips that we specify in Materials and Reagents, 32 mL is sufficient.
Note: Piranha solution is hazardous and can be life-threatening if not handled properly! It is advised to consult the safety department of your institution and get training and permission before use. All procedures involving Piranha solution must be performed under a fume hood, and the experimental personnel should wear protective equipment, including lab coat, acid resistant gloves, and goggles. It is very important to keep the area free of any organic solutions, such as acetic acid and ethanol, since mixing Piranha solution with organic solutions is strongly exothermic and may lead to an explosion.
Put a cleaned and dry glass beaker with an acid-resistant stirring bar on a magnetic stirring plate. Do not use heat since preparation of Piranha solution is already strongly exothermic. Starting from this step, the operation must be performed under a chemical fume hood.
With a clean glass pipette, add three volumes (3/4 of the total needed volume of Piranha solution) of concentrated sulfuric acid into the beaker. For 32 mL of Piranha solution, three volumes equal 24 mL.
Turn on stirring at a low speed, so liquid does not spill out of the beaker.
With a glass pipette, slowly add one volume (1/4 of the total needed volume of Piranha solution) of 30% hydrogen peroxide solution to the sulfuric acid in the beaker while slowly stirring it. For 32 mL of Piranha solution, one volume equals 8 mL.
Note: Heat very quickly builds up and the beaker becomes hot, so use caution when handling it.
Keep the solution stirring for approximately 3 min to fully mix up and stabilize.
Pour the Piranha solution into the Coplin jar with coverslips. Before pouring, attach a magnet under the beaker to hold the stirring bar, so it will not fall out.
Cover the Coplin jar with the lid but do not seal it since gas will be generated in the cleaning process.
Note: Sealing the lid can lead to an explosion.
Leave the filled jar under the laminar hood for five days.
To handle the residual Piranha solution left in the beaker, use a glass pipette under the laminar hood to slowly add 50 mL of water into the beaker while stirring it on the magnetic plate. While adding water, attach the tip of the glass pipette to the inner wall of the beaker to avoid any splash of liquid out of the beaker.
Slowly add 5 M sodium hydroxide solution to neutralize the diluted residue Piranha solution while keeping it stirred. Use a pH indicator paper to confirm neutrality.
Pour the neutralized solution into the sink.
When the cleaning is done after five days, prepare a large beaker full of crushed ice with a stirring bar on the bottom. The size of this beaker should be at least 30 times the volume of the Piranha solution.
Put the beaker on the magnetic plate under the laminar hood. Do not use heat.
Carefully pour the Piranha solution onto the ice in the beaker while retaining the coverslips in the Coplin jar.
Start stirring slowly.
When the ice is melted, carefully add NaOH pellets to the solution to neutralize it while stirring.
Discard the solution according to institutional guidelines when it is pH neural, stabilized, and cooled down.
Rinse the coverslips in the Coplin jar with double-distilled water 20 times. Collect the waste from the first three rinses into a beaker and neutralize with NaOH solution before discarding.
For storage, immerse the cleaned coverslips in double-distilled water in the same jar with the lid sealed with parafilm. Keep refrigerated.
Cell culture and passage of HEK293T cells
Culture HEK293T cells in culture medium (DMEM supplemented with 50 U/mL penicillin-streptomycin and 10% FBS) in 10 cm Petri dishes at 37 °C in the cell culture incubator supplemented with 5% CO2 (Tan et al., 2017). Passage them at least once a week. Refresh the medium every three days. To passage the cells, follow the procedures below:
Culture the stock cells with 10 mL of medium in 10 cm Petri dishes. Aspirate the medium from the dish.
Wash the cells with 10 mL of PBS three times. Aspirate the solution.
Add 1 mL of trypsin solution warmed up to 37 °C in advance to the dish. Evenly spread the solution on the bottom of the dish.
Incubate the dish on a 37 °C heating plate for 1 min.
Shake the dish on the desk to fully detach the cells from the dish bottom. Check under the cell culture microscope if cells are detached.
Add 9 mL of culture medium (pre-warmed to 37 °C) to the dish. Split the cells by resuspending the cell suspension 10 times to get single cells. Examine the cell suspension under the microscope and check if cells are separated.
Add 1 mL of cell suspension into a new 10 cm dish with 10 mL of fresh medium pre-warmed to 37 °C. Gently shake the dish to evenly distribute the cells.
Place the dish into a cell culture incubator. Passage the cells ten times until replacing with newly thawed cells.
Cell transfection
To prepare cells for transfection, use cleaned and sterilized tweezers to transfer a cleaned 25 mm coverslip into a 3.5 cm Petri dish. The coverslips have become hydrophilic during treatment with Piranha solution, so no further surface coating is required for HEK293T cells.
Add 2 mL of warm culture medium to the dish.
Dispense 0.1 mL of cell suspension onto the coverslip prepared as described above for cell passaging. Swirl the dish carefully to distribute the cells evenly. The volume of the cell suspension should be chosen such that the cells are sparsely distributed on the dish bottom and separate from one another after they settle down.
Keep the cells in the incubator for 6 h to let them settle down.
Prepare two autoclaved 1.5 mL microcentrifuge tubes and add 100 μL of OptiMEM to each of them. Add 1 μg of plasmid of interest to one tube and 3 μL of Lipofectamine 2000 to the other one.
Incubate both tubes separately at room temperature for 5 min.
Slowly transfer the DNA-OptiMEM mixture to the Lipofectamine 2000–containing tube drop by drop.
Incubate the mixture at room temperature for 20 min.
Drop by drop, add the mixture to the 3.5 cm Petri dish of cells prepared 6 h before.
Incubate the cells in the incubator overnight.
Cell fixation
After the overnight incubation with the transfection mixture, aspirate the culture medium.
Wash the cells gently with 2 mL of PBS five times. To avoid introducing impurities, the PBS solution should be filtered with a 0.22 μm sterile filter during preparation, stored in a cleaned and autoclaved glass bottle, and refrigerated.
Add 2 mL of -20 °C methanol to the dish and incubate at -20 °C for 5 min.
Remove the methanol and wash five times with 2 mL of PBS.
Keep the cells in PBS.
Imaging of the standard proteins and the proteins of interest
Note: The following steps are specific to each individual microscope/control software combination. We give example values and settings for the microscope used in Tang et al. (2015) and Tan et al. (2022). The proper values for your setup (in particular for the camera and laser) will have to be locally determined and adjusted. Please discuss with your appropriate local experts and technicians, if necessary.
After fixation, take the coverslip with cells from the Petri dish and mount in a recording chamber. Our chamber was custom-made, holding up to 2 mL of solution and being placed on the stage of the microscope, stabilized with magnets.
To track the sample drift and for chromatic aberration corrections, dilute 1 μL of TetraSpeck fluorescent bead solution (vortexed) with 0.5 mL of PBS and then transfer it to the imaging chamber containing the coverslip with the fixed cells.
Leave the sample in a light-proof box for 1 h in the imaging room. This allows the beads to settle and attach to the coverslip (Figure 1A and 1B) and the temperature to reach the lab temperature (22 °C), minimizing sample drift due to thermal shift.
Note: The lab temperature should be monitored and maintained by air conditioning at a constant temperature (in our case, 22 °C) to avoid thermal drift and to guarantee the optimal performance of the setup.
Figure 1. Representative image for the appropriate density of fiducial markers and PAmCherry blinking events. (A) A total of 12 beads (marked by green arrow heads) are attached to the coverslip around the cell after the pre-imaging incubation, shown as bright spots. Cells with beads on top are usually not used for imaging because the bright emissions from the beads interfere with the fluorescent proteins. Scale bar, 5 μm. (B) Gold indicates the area covered by the cell, while the red arrow heads indicate the blinking events recorded within the cell area on the PAmCherry channel during a single frame. Very bright spots around the cell are emissions from the fluorescent beads seen in A. Scale bars, 5 μm.
For the same purpose, add 2 mL of PBS for use as imaging buffer to a 1.5 mL microcentrifuge tube and let it warm up to the lab temperature.
During the bead incubation period, switch on the imaging system to warm it up for at least half an hour.
Align the lasers and adjust the laser intensities at the sample plane according to the manufacturer’s instructions. Laser intensities must be chosen so that individual blinking events are spatially separated to allow for the extraction of super-resolution information. On the other hand, recording durations should allow for the activation and bleaching of all fluorescent proteins present in the sample. As a guide, for our experiments, we measured an intensity of 4.4 mW for the 514 nm laser and 5.4 mW for the 561 nm laser in the sample plane. We increased the intensity of the 405 nm laser during PAmCherry imaging slowly from 3.0 µW to 4.8 mW, so that each frame contained several spatially separated blinking events.
Aspirate the bead solution from the coverslip and rinse it gently with 1 mL of PBS pre-warmed to the lab temperature.
Remove the solution and gently add 1 mL of PBS pre-warmed to the lab temperature.
Put the imaging chamber onto the microscope stage. Let it stabilize for 15 min. To lower the background signal level and therefore increase the signal-to-noise ratio, either darken the lab or, even better, cover the stage area.
Set up the parameters for the camera to have 50 ms exposure and 85.59 ms as the duration between frames, including read-out of data from the camera. The frame duration affects the number of frames that a blinking event covers. Therefore, one must tweak the minimal number of localizations parameter for the clustering analysis if different values are chosen.
In widefield mode, use the green imaging channel (for mVenus) to choose a cell with moderate-to-low expression level of the transfected plasmid.
Move the cell to the center of the view. Around the cell there should be 3–6 beads seen as bright spots in both color channels. Trajectories of the individual beads during the acquisition will be used for the sample drift correction.
In widefield mode, turn on the 561 nm laser for approximately 3,000 frames to bleach impurities within the sample. Background signals will be significantly reduced while PAmCherry molecules are largely unaffected without activation by the 405 nm laser.
Adjust the axial position of the sample stage to bring the beads into focus, as determined by reaching the maximal fluorescence from the beads.
With the white light and the 561 nm laser on, take a transmission image to record the location and shape of the cell and beads.
Switch to TIRF mode and record 4,000 frames with the 514 nm laser for mVenus. Usually, all mVenus molecules should be bleached at the end of recording. In case there are still many blinking events at the end, recording can be continued for another 2,000 frames. If this extended period still does not exhaust blinking events, it indicates either that the expression level is too high, or the laser power is too low. With the emission filter settings and the cleaning procedure, there should be very few blinking events on the green channel from the impurities outside of the cell. This is critical for the DCC algorithm to work.
With the same TIRF settings, record 6,000–12,000 frames with the 561 nm laser for PAmCherry, with 405 nm activation laser intensity increasing from the minimal to the maximal laser power (Section B.4), allowing the emission events to be sparsely distributed (as shown in Figure 1B) and all PAmCherry molecules to be bleached at the end of the recording. If 12,000 frames cannot exhaust all the blinking events within the cell, it may indicate that the expression level is too high.
Adjust the axial position of the sample stage and bring it to focus judging from the intensity of the beads. If the offset is larger than 100 nm, then the recording is considered invalid due to large z-axis drift and must be discarded.
To record beads for the lateral chromatic aberration (LCA) correction using the fitted LCA protocol with the same sample, choose a region on the coverslip that does not contain any transfected cells but only beads (> 10, the more the better as long as they do not overlap with one another). In widefield mode, record 50 frames for each of the green and red channels as a single recording. Slightly move the sample to change the position of the beads and record another 50 frames. Repeat this moving and recording procedure numerous times to extract information from beads in at least 100 different positions, evenly distributed across the whole field of view.
Preparation of a bead sample
To record beads for the lateral chromatic aberration correction using the regional LCA protocol, prepare a bead-only sample for the SMLM recording. This sample can be stored in the dark at room temperature and lasts for several experiments.
Clean a microscope slide (75 mm × 25 mm) as for the coverslips with Piranha solution and let it air dry before use.
Thoroughly vortex the multi-color TetraSpeck fluorescent bead solution for 1 min.
Add 5 μL of bead solution into 400 μL of double-distilled water in a 1.5 mL Eppendorf tube and then vortex to mix it.
Add the diluted bead solution to a cleaned coverslip (25 mm) and let it air dry at room temperature.
After drying, place the coverslip on a microscope slide with the side of beads facing down towards the slide.
Dip a micropipette tip in nail polish and place 4–5 tiny drops on the edge of the coverslip to glue the coverslip onto the slide.
Once the nail polish is dry, the bead sample is ready for use.
Store the sample in the dark at room temperature until next imaging.
Imaging of the bead sample
Note: The following steps are specific to each individual microscope/control software combination. The exact values (in particular for the camera and laser) will have to be locally determined and adjusted for your setup. Please discuss with your appropriate local experts and technicians.
Launch the microscope and align the lasers as for the cell sample.
Mount the bead sample slide on a custom-made chamber that can stick to the microscope stage with magnets. The coverslip should be facing towards the objective when the slide is on the microscope stage.
Place the sample on the microscope stage and let it stabilize for approximately 20 min.
Set up the acquisition parameters so that pixel size, exposure time, and frame rates are the same as those for cell sample imaging. The excitation laser intensities can be reduced to avoid over-exposure since beads are much brighter than fluorescent proteins.
In widefield mode, record the bead sample for 50 frames each in the green and red channels as a single recording.
Slightly move the sample and repeat the imaging if the density of the beads is too low. Usually, a total of 500 beads or 500 positions from fewer beads in the field of view can be sufficient, as shown in Figure 2.
Figure 2. Example of bead recordings used for the lateral chromatic aberration correction. Left: A total of 18 individual bead recordings are combined to occupy the whole camera view (~41 × 41 μm). Each white spot indicates a bead recorded on the PAmCherry channel, spanning 50 frames. Scale bar, 5 μm. Right: Enlarged area defined by the red square in the left image. Scale bars, 1 μm.
Extraction of localizations
Using SNSMIL software:
We used SNSMIL (Tang et al., 2015) to localize the emission events in the recorded videos (image frames). This software was developed using recordings taken with our microscope setup and is therefore particularly optimized for our use case.
Alternatively, we have used the newer software SMAP (Ries, 2020) to extract localizations from the fluorescence signals in the recorded videos. This software uses improved algorithms and may work better with other microscopes. In our hands, we did not find a general advantage of using either SMAP or SNSMIL.
Data analysis
The DCC-SMLM algorithms have been integrated into a Python library. To use this library and the example of the analysis as explained below, a working knowledge of the Python programming language is required. For the examples given below, you also need to have some familiarity with Jupyter Notebooks. The protocol below should be studied while simultaneously working with the indicated files as stated in the beginning of each step. We have added a flowchart that may serve as a guide to the necessary steps and files used during analysis (Figure 3).
Figure 3. Flowchart for the different steps used during analysis. Green boxes indicate necessary microscope recordings and yellow boxes indicate analysis steps with the mentioned example Jupyter notebook scripts. (A) Two-color recordings of fluorescent TetraSpeck beads are used to determine chromatic aberration (CA). For this, at least 500 beads spread across the whole field of view should be used. It is not required to record 500 separate beads, but the same sample may be moved in between recordings, resulting in multiple recordings that can be stacked together to increase the number of recorded positions. The CA parameters are saved in a .csv file and will be used by the scripts in the other steps (purple arrow). (B) Colocalization ratios of proteins with known oligomeric state must be individually determined. In our experience, at least 12 recordings per protein should be taken. The ratios are then used to calibrate the values of p and m in the DCC-SMLM model (Equation 2). These values will be used in the next step (brown arrow). (C) Colocalization ratios of proteins with unknown oligomeric states are recorded and individually analyzed. These analyses are then used to determine the most likely oligomeric state of the protein of interest.
Installation of the DCC-SMLM library
Please make sure that you have Python 3 and the Jupyter Notebook installed. If not, we recommend downloading the free version of the Anaconda distribution for your operating system from https://www.anaconda.com. This distribution contains all required tools and libraries to use our analysis scripts.
Download our library from https://github.com/GabStoelting/DCC-SMLM into a separate folder. This may be done directly from the GitHub website, using command line, or GUI git tools.
If you want to use the library for your own project, we currently recommend copying the DCCSMLM.py file to a new project folder. The example notebooks may be copied and later modified as well.
Determination of the chromatic aberration (Figure 3A)
Note: Open the “CA_example.ipynb” example in Jupyter Notebook and work through the code from top to bottom. The explanations below explain the DCC-SMLM library specific functions as they occur in the Notebook from top to bottom.
Load all extracted localizations from the two-color bead recordings (see Procedure C) in the .SciH5 file from a processed SNSMIL extraction or the .mat output file from SMAP.
Identify bead signals using the find_clusters function for each color channel. The parameters are:
i. intensity_threshold, which defines the minimum fluorescence intensity per localization in arbitrary units.
ii. min_samples, which sets the minimum number of frames the bead must be recorded in.
iii. eps, which is the epsilon parameter for the underlying DBSCAN clustering, being a measure for the maximal distance between localizations within a cluster.
iv. save_column, which is the name of the column of the internal data structure that will be used to assign each bead a specific ID number.
The chromatic aberration between the same bead identified in the green and red color channels is calculated using the determine_chromatic_aberration function. In principle, this function identifies the closest bead in one color channel relative to another and measures the deviation. The parameters are:
i. The first parameter is the bead dataset in the green channel.
ii. The second parameter is the bead dataset in the red channel.
iii. The third parameter is the name of the save_column set in the previous step.
iv. Lastly, a cutoff can be specified that will limit the range for the search of beads in the opposite color (distance_cutoff).
As shown in the literature, chromatic aberration varies linearly relative to the optical point of the microscope. The measured distance of the same bead in each color channel can therefore be plotted as a function of the x- or y-coordinate. A plot of the data from all available beads can be fit with a linear function and used to calculate the necessary correction for every coordinate within the field of view.
The fit parameters can be saved and used for correction in other analyses below.
Analysis of a single recording (Figure 3B and 3C)
Note: Open the “SciH5_example.ipynb” or “SMAP_example.ipynb” examples in Jupyter Notebook and work through the code from top to bottom. The explanations below explain the DCC-SMLM library specific functions as they occur in the Notebook from top to bottom.
Load the .SciH5 or .mat file as described above using the load_channel function. If you previously determined and saved the parameters of the linear fit for the chromatic aberration correction, you can pass the ca_file parameter to that .csv file. This will automatically correct the coordinates of all localizations within the channel.
Identify the beads in the recording for determination and correction of sample drift. For this, the find_clusters method is again used as described above for the bead sample. Beads can usually be separated by their high fluorescence intensity relative to the signals emitted by fluorescent proteins.
The extract_drift function will extract the shift of the position of each bead from one frame to the next. Missing data is interpolated as described in our original publication. The mean drift is saved in the drift variable of each color channel. The parameters of this function are:
i. The first parameter contains the localizations of the beads.
ii. The second parameter contains the name of the column in which the bead IDs were assigned.
Clusters of emissions by fluorescent proteins are identified using the find_clusters function. The intensity_threshold and min_samples parameters should typically be much lower than for identifying fluorescent beads.
The colocalization ratio can be calculated using the get_colocalization_ratio function. The parameters are:
i. The names of the first color channel.
ii. The name of the second color channel.
iii. Name of the bead ID.
iv. A distance threshold is set using the distance_cutoff parameter.
v. Regions of interest can be defined by providing a list of rectangles in the format (x, y, width, height).
The resulting colocalization ratio can be saved for further analysis.
Analysis of calibration proteins (Figure 3B)
Note: Open the “Calibration_example.ipynb” example in Jupyter Notebook and work through the code from top to bottom. The explanations below explain the DCC-SMLM library specific functions as they occur in the Notebook from top to bottom.
Load the file containing the combined colocalization ratios of several recordings of calibration proteins with known oligomeric state as described in the previous step. Use the DCCReferenceProteins function for this.
i. The first parameter is the file name of the .csv file containing the data.
ii. The second parameter contains the name of the column that contains the colocalization ratios.
iii. The third defines the name of the column containing the known oligomeric state corresponding to the measured ratio.
The reference_bootstrap function determines the values of p and m from the calibration dataset. The first parameter defines the number of bootstraps performed; save_result = True will save the values of p and m to the DCCSMLM object as well as the confidence intervals.
The determined values for m and p can be saved for use in the analysis of proteins with unknown oligomeric state.
Estimation of the oligomeric state of proteins of interest (Figure 3C)
Note: Open the “Example_POI_analysis.ipynb” example in Jupyter Notebook and work through the code from top to bottom. The explanations below explain the DCC-SMLM library specific functions as they occur in the Notebook from top to bottom.
First, load the table with the reference proteins as stated in step 4.
Load a file containing the colocalization ratios of several recordings of proteins of interest using the DCCProteinOfInterest function. Give the name of the .csv file as first and the name of the column containing the colocalization ratios as second parameter.
Now, determine the coefficient of mismatch (COM) as described in our original publication. In summary, the COM is proportional to the difference of the observed colocalization ratio to the expected ratio as calculated from the calibrated values of m and p. The com_bootstrap method calculates the COM for a bootstrapped sample to also give statistical information. The first parameter defines the number of bootstrap samples, and the second parameter contains the object containing the reference data as determined in the previous step. Setting reference_bootstrap = True will also bootstrap the reference (calibration) dataset as described above to consider the propagation of the errors of the calibration dataset as well as of the protein of interest.
Plot the COM. The smallest value corresponds to the most likely oligomeric state. The bootstrap permutation will also give an estimation of the uncertainty of this value.
Additionally, an analysis using a Kolmogorov-Smirnov test can be performed. For this method, the distribution of the colocalization ratios of the protein of interest is compared against the distribution of the calibration proteins. For this, we implemented the KS function.
Recipes
PBS
8 g of NaCl, 0.2 g of KCl, 1.15 g of Na2HPO4·2H2O, 0.2 g of KH2PO4
Dissolve in 500 mL of double-distilled water and fill up to 1 L. Adjust pH to 7.3–7.4 and filter the solution with the sterile filter. Store in an autoclaved glass bottle at 4 °C.
Acknowledgments
Development of this protocol was supported by the Deutsche Forschungsgemeinschaft (FA 301/15–1 to Ch.F.) as part of Research Unit FOR 5046, project P4. We acknowledge the original research paper (Tan et al., 2022), from which this protocol is derived, as the primary reference when additional information is needed.
Competing interests
The authors declare no competing financial or non-financial interests.
Ethical considerations
No human or animal tissues or samples are involved in this protocol.
References
Betzig, E., Patterson, G. H., Sougrat, R., Lindwasser, O. W., Olenych, S., Bonifacino, J. S., Davidson, M. W., Lippincott-Schwartz, J. and Hess, H. F. (2006). Imaging Intracellular Fluorescent Proteins at Nanometer Resolution. Science 313(5793): 1642-1645.
Durisic, N., Laparra-Cuervo, L., Sandoval-Álvarez, Ã., Borbely, J. and Lakadamyali, M. (2014). Single-molecule evaluation of fluorescent protein photoactivation efficiency using an in vivo nanotemplate. Nat Methods 11(2): 156-162.
Lee, S. H., Shin, J., Lee, A. and Bustamante, C. (2012). Counting single photoactivatable fluorescent molecules by photoactivated localization microscopy (PALM). Proc Natl Acad Sci U S A 109(43): 17436-17441.
Nan, X., Collisson, E. A., Lewis, S., Huang, J., Tamgüney, T. M., Liphardt, J. T., McCormick, F., Gray, J. W. and Chu, S. (2013). Single-molecule superresolution imaging allows quantitative analysis of RAF multimer formation and signaling. Proc Natl Acad Sci U S A 110(46): 18519-18524.
Ries, J. (2020). SMAP: a modular super-resolution microscopy analysis platform for SMLM data. Nat Methods 17(9): 870-872.
Schägger, H. and von Jagow, G. (1991). Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem 199(2): 223-231.
Sengupta, P., Jovanovic-Talisman, T., Skoko, D., Renz, M., Veatch, S. and Lippincott-Schwartz, J. (2011). Probing protein heterogeneity in the plasma membrane using PALM and pair correlation analysis. Nat Methods 8(11): 969-975.
Tan, H., Bungert-Plümke, S., Fahlke, C. and Stölting, G. (2017). Reduced Membrane Insertion of CLC-K by V33L Barttin Results in Loss of Hearing, but Leaves Kidney Function Intact. Front Physiol 8: e00269.
Tan, H., Bungert-Plümke, S., Kortzak, D., Fahlke, C. and Stölting, G. (2022). Determination of oligomeric states of proteins via dual-color colocalization with single molecule localization microscopy. Elife 11: e76631.
Tang, Y., Dai, L., Zhang, X., Li, J., Hendriks, J., Fan, X., Gruteser, N., Meisenberg, A., Baumann, A., Katranidis, A., et al. (2015). SNSMIL, a real-time single molecule identification and localization algorithm for super-resolution fluorescence microscopy. Sci Rep 5(1): 11073.
Ulbrich, M. and Isacoff, E. (2007). Subunit counting in membrane-bound proteins. Nat Methods 4(4): 319-321.
Zhang, M., Chang, H., Zhang, Y., Yu, J., Wu, L., Ji, W., Chen, J., Liu, B., Lu, J., Liu, Y., et al. (2012). Rational design of true monomeric and bright photoactivatable fluorescent proteins. Nat Methods 9(7): 727-729.
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4,750 | https://bio-protocol.org/en/bpdetail?id=4750&type=0 | # Bio-Protocol Content
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Peer-reviewed
Ex vivo Culture and Contractile Force Measurements of Non-human Primate Heart Slices
CP Christine M. Poch
AD Andreas Dendorfer
KL Karl-Ludwig Laugwitz
AM Alessandra Moretti
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4750 Views: 371
Reviewed by: Ralph Thomas BoettcherDavide BottaFarah Haque
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Original Research Article:
The authors used this protocol in Nature Cell Biology May 2022
Abstract
Cardiovascular diseases are the leading cause of death and morbidity worldwide. Patient mortality has been successfully reduced by nearly half in the last four decades, mainly due to advances in minimally invasive surgery techniques and interventional cardiology methods. However, a major hurdle is still the translational gap between preclinical findings and the conversion into effective therapies, which is partly due to the use of model systems that fail to recapitulate key aspects of human physiology and disease. Large animal models such as pigs and non-human primates are highly valuable because they closely resemble humans but are costly and time intensive. Here, we provide a method for long-term ex vivo culture of non-human primate (NHP) myocardial tissue that offers a powerful alternative for a wide range of applications including electrophysiology studies, drug screening, and gene function analyses.
Graphical overview
Keywords: Non-human primate Native myocardium Ex vivo culture Contractile force measurement Animal model Long-term culture
Background
Cells isolated from animal or human hearts by enzymatic dissociation have provided fundamental insights into the electrophysiology, contractile function, and transcriptional profiles of cardiomyocytes (Liu et al., 2021). However, cells cultured in 2D lack the native environment of the 3D myocardium, which includes intra- and intercellular interactions and syncytial properties that are determinant factors in studying cells, genes, and drug modulations in cardiac pathologies. It is also known that functional and structural preservation of native myocardial tissue ex vivo is dependent on the application of both mechanical preload and electrical stimulation (Brandenburger et al. 2012; Watson et al., 2017; Perbellini et al., 2018; Qiao et al., 2019). Recently, Fischer et al. (2019) established and commercialized a biomimetic ex vivo culture system taking these parameters into account to maintain intact native human myocardial slices in culture for up to four months (Fischer et al., 2019; www.invitrosys.com). This novel technology enables human disease modeling with high temporal and spatial resolution in assessing cellular behavior after pharmacologic or genetic interventions. A drawback of this method is that human tissue is not readily available, with samples generally originating from end-stage heart failure patients with a wide range of comorbidities. Moreover, the patients’ age, genetic background, and medication can be confounding factors during ex vivo analyses. To enable standardized investigational studies, we adapted this system to the culture of non-human primate (NHP) heart slices.
NHP hearts are an excellent surrogate to address basic research questions due to their genetic, metabolic, and physiologic similarities to humans and the ability to control diet, environment, and breeding (Cox et al. 2017). As described in the step-by-step protocol below, left ventricular transmural sections of native NHP hearts are cut into thin tissue slices (300 μm) that are subsequently subjected to physiological preload and electrical stimulation. Under biomimetic culture conditions, a continuous readout of contractile performance and electrophysiologic behavior such as excitability, force-frequency relationship, and effective refractory period can be reached for up to three weeks. Furthermore, as we recently reported, this system can be used as a platform to study critical steps of cardiac regeneration processes in single-cell resolution demonstrating that time- and resource-consuming in vivo studies in animals can be partially bridged (Poch et al. 2022).
Materials and reagents
35 mm Petri dish (Corning Life Sciences, catalog number: CLS430165)
50 mL capped Falcon tube (Corning Life Sciences, catalog number: 352070)
Medium 199, Earle’s salts (M199) (Gibco, catalog number: 11150059)
Insulin-transferrin-selenium (ITS) (Gibco, catalog number: 41400045)
Penicillin-streptomycin (P/S) (Gibco, catalog number: 15140122)
β-Mercaptoethanol (Bio-Rad, catalog number: 1610747)
Histoacryl tissue glue (Braun, catalog number: 04929052)
PET plastic triangles (InVitroSys, Germany)
Isopropyl alcohol (Fischar, catalog number: 08819076)
NaCl (Sigma, catalog number: S5886-1kg)
KCl (Merck, catalog number: 1.04933.0500)
MgCl2·6H2O (AppliChem, catalog number: A1036,0500)
NaH2PO4·H2O (Merck, catalog number: 1.06346.0500)
Glucose·H2O (AppliChem, catalog number: A3730,0500)
CaC12·2H2O (Merck, catalog number: 2382.1000)
BDM (2,3-Butanedione monoxime) (Sigma-Aldrich, catalog number: B0753)
HEPES (AppliChem, catalog number: A 1069,0500)
Filtration unit (Steritop Quick Release, Millipore, S2GPTOSRE)
Agarose low melt (Carl Roth, catalog number: 6351.2)
Cutting buffer (see Recipes)
4% agarose preparation (see Recipes)
Culture medium M199 (see Recipes)
Equipment
Vibratome VT1200S (Leica Biosystems, Germany)
Vibrocheck device (Leica Biosystems, Germany)
MyoDish 1 Tissue Culture System (InVitroSys, Germany)
Incubator (37 °C, 5% CO2) (Thermo Scientific, catalog number: 4110)
Water bath (37 °C)
Razor blade (Wilkinson Sword)
Scalpel (Feather, catalog number: 00636494)
Tweezer (Mmobil®, catalog numbers: ESD-15 and ESD-10)
Software
MyoDish Software (InVitroSys, www.invitrosys.com)
MyoDish Data File Converter v1.1 (inVitroSys, www.invitrosys.com)
LabChart Reader (v8.1.24, www.adinstruments.com)
Microsoft Excel 2019 (www.microsoft.com)
GraphPad Prism 9 (www.graphpad.com)
Procedure
Preparation and assembly of MyoDish culture chambers
Prepare the cutting buffer, 4% agarose, and M199 medium as listed in the Recipes section below.
Sterilize electrodes of MyoDish Culture system (autoclave or place in isopropyl alcohol).
Prepare plastic triangles by detaching them from the holding sheet. Sterilize by autoclaving or submersion in isopropyl alcohol.
Note: Triangles can be purchased from InVitroSys or manually prepared from a foil of polyethylene terephthalate (PET, 125 μm thick).
Clean chambers with isopropyl alcohol and let them dry under a laminar flow hood.
Assemble chambers and electrodes according to manufacturer’s instructions (www.invitrosys.com; Figure 1).
Figure 1. Schematic of assembled MyoDish culture chambers. Electrodes for field stimulation and plastic triangles attached to myocardial slice for anchorage onto spring and holding wires. NHP: non-human primate.
Cutting of tissue slices
Warm M199 culture medium in the water bath (37 °C), add 2.4 mL of medium into each chamber, and place them on the rocker platform in the 37 °C incubator.
Fill 10 mL of 4% agarose into a syringe capped with a Luer lock cap and place it into the water bath until equilibrated to 37 °C.
Turn on the vibratome (placed under laminar flow hood) and reduce Z-distortion of the razor blade using the Vibrocheck device. Sterilize with isopropyl alcohol all parts that may possibly come in contact with the tissue or cutting solution.
The freshly explanted NHP heart needs to be processed immediately after explantation. Cut out a mid-ventricular, approximately 2 cm × 3 cm transmural muscle piece.
Note: Fiber orientation is not ideal if the tissue is excised too apically (make sure to cut mid-ventricular; cutting plane indicated on Figure 2A).
If heart samples cannot be cut and taken into culture immediately (e.g., tissue needs to be shipped), store mid-ventricular samples in a 50 mL capped Falcon tube filled with cutting buffer on wet ice. Delivery should be on the same day.
Place transmural NHP left ventricular tissue blocks into a 35 mm Petri dish containing cold cutting buffer.
Note: The tissue must be continuously stored in cold cutting buffer (4 °C) to avoid hypercontraction and to keep the tissue alive.
Remove endocardial and trabecular layers using sterilized scissors or scalpel. Choose a region without fibrous tissue and of vital color. Orient the tissue according to morphology (uniform fiber orientation, vital color) and trim edges with a razor blade to obtain a tissue block of approximately 1.5 cm × 1 cm in size (Figure 2B and 2C). Fiber orientation is best visible at tissue edges; if it is not visible by eye, use a light microscope equipped with a 10× or 20× lens.
Transfer freshly trimmed tissue blocks into a dry 35 mm Petri dish with the epicardium facing down; the bottom should be dry and stick to the dish.
Figure 2. Processing of non-human primate (NHP) heart. A) Schematic of native NHP heart with indicated, mid-ventricular cutting plane on the left ventricle. B) Mid-ventricular transmural left ventricular (LV) section of ~2 cm × 3 cm. Epicardium facing down and endocardial and trabecular layers facing upward. C) Trimmed LV cube (~1.5 cm × 1 cm) after removal of endocardial and trabecular layers. Uniform, diagonal fiber orientation is visible. Epicardium facing down. D) Trimmed tissue block embedded in 4% agarose within 35 mm Petri dish. Epicardium facing down.
Embed tissue pieces in preheated (37 °C) 4% agarose (up to two pieces in one 35 mm dish) and let solidify for approximately 3–5 min (Figure 2D). Remove excessive agarose and trim to rectangular shape.
Note: Leave a minimum of approximately 5 mm of agarose around the tissue to ensure stability during cutting process.
Attach the prepared tissue-agarose blocks to the vibratome cutting tray (epicardium facing down; Figure 3) by using histoacryl tissue glue.
Note: Make sure that all edges are well attached to the surface.
Place the vibratome cutting tray into the cutting bath filled with 4 °C cutting buffer. Tissue edges should be parallel to the razor blade (Figure 3).
Place wet ice into the outside tub of the vibratome (ice bath) to make sure that the tissue is continuously cooled to 4 °C during the cutting process (Figure 3).
Move the razor blade towards the tissue, set the borders, and start cutting with a vibration amplitude of 1.3 mm at a speed of 0.07 mm/s and thickness of 300 μm.
Note: Do not increase the cutting speed once the razor blade reaches the tissue. Discard the first slices in which the tissue is not evenly cut.
Figure 3. Image of vibratome cutting. A) Overview of vibratome prepared for cutting. B) Zoom into boxed area showing vibratome bath filled with cutting buffer surrounded by an ice bath (filled with wet ice). Myocardial tissue block in 4% agarose glued to vibratome cutting tray. Razor blade adjusted parallel to tissue block edges.
Transfer one slice from the vibratome bath to the lid of a Petri dish using tweezers. Detach the slice from agarose with tweezers and make sure to only touch the tissue at the borders.
Remove any residual cutting buffer from the surface with a pipette (do not touch the tissue directly) to prepare the slice to be attached to plastic triangles.
Pipette a stripe of approximately 10 μL of histoacryl glue to the surface of a Petri dish, dip the edge of a plastic triangle into it, and quickly press it onto the edge of the tissue slice. Make sure to attach the triangle perpendicular to the orientation of the muscle fibers (Figure 4A and 4B). If fiber orientation is not visible by eye, use a light microscope for magnification (Figure 4C).
Attach the second plastic triangle in a similar manner on the opposite site of the tissue. Afterwards, trim the tissue on the edges to the size of the triangle.
Repeat the procedure with the other freshly cut tissue slices in the vibratome bath. Make sure to always glue the triangles according to fiber direction.
Note: Slices with attached plastic triangles can be stored in cold cutting buffer up to 1 h before moving them to culture dishes.
Figure 4. Myocardial slice with attached plastic triangles rectangular to muscle fiber orientation. A) schematic and B) representative image of myocardial slice within MyoDish culture chamber with C) 25-fold magnification to visualize fiber orientation. Arrows indicate fiber orientation.
Note: Plastic triangles are equipped with a pinhole to be anchored on spring and holding wires.
Transfer to biomimetic culture chambers
Take an assembled MyoDish culture chamber containing 2.4 mL of M199 medium from the incubator. Hook the slices onto the spring and holding wires with tweezers. Start with the attachment to the upper spring wire. The second triangle is pinned onto the lower holding wire after adjustment of the correct distance according to the individual length of the tissue (Figure 5).
Figure 5. Image of biomimetic MyoDish culture chamber with 2.4 mL of M199 medium. The non-human primate (NHP) myocardial slice is pinned on the upper spring and holding wires before adjusting the preload.
Submerge the slice into the medium and stretch the tissue until it is properly aligned.
Cover the chamber with a lid and transfer it to the incubator. Make sure to have a constant temperature of 37 °C in the incubator, as temperature changes can influence viability.
Carefully increase the diastolic tension by increasing the tissue stretch until a mere deflection of contraction is detectable (= baseline).
Adjust the diastolic force to a physiologic preload of 1 mN by progressively increasing the tissue stretch from baseline to 1,000 units. Sharp contraction signals should be detectable immediately. This adjustment of the preload should be repeated after the first 48 h in culture.
Start electrical stimulation with 50 mA at a frequency of 1 Hz.
Start rocking of slices with a target frequency of 60 rpm.
Start data recording by entering the storage location and file name and choose autosave settings (set it to 24 h). Detailed information on technical settings can be found on the manufacturer’s homepage (www.invitrosys.com).
Medium change
Perform medium change every other day by leaving a residual volume of 0.6 mL of old medium in the dish followed by addition of 1.8 mL of fresh, prewarmed M199 medium. Ensure that the fresh medium remains heated to 37 °C during the whole medium exchange procedure.
Exchange medium in all culture chambers one after the other and make sure you place the chamber immediately back into the incubator. Ensure that the atmosphere and temperature in the incubator are not changing significantly during this process, as this is a critical step for successful long-term cultures.
Data analysis
Save files with raw data of measurements in one folder.
Start MyoDish Data File Converter v1.1 and convert files into Axon binary format (.abf).
Select LabChart Reader as the default program for .abf files and open .abf files.
LabChart Reader displays all selected channels among each other in different colors. Set all channels into “autoscale” (command in top bar).
Adjust the timeline displayed at the bottom right (how many measurements per screen pixel). Adjust the timescale by clicking right on the time axis (e.g., display in seconds from start of the file). In the display settings (Setup menu) deselect “Lines between blocks.” This way, a full view of contractile force during the whole culture period, as well as analysis of single days, is possible (Figure 6).
Measure the contraction force by determination of the amplitude. Define one column of the Data Pad as “cyclic measurements” and assign it to the calculation of “average cyclic height.” Choose “peak detection” in the setup panel of the column. Mark a registration period of interest in the data tracing and choose “Add to Data Pad.” The average value of all beat amplitudes within this time period will be displayed in the most recent line of the Data Pad.
Copy the Data Pad to a spreadsheet program (e.g., Microsoft Excel) and calculate contraction force by dividing the arbitrary units by the calibration value supplied with the specific chamber (approximately 1,000 AU/mN).
Export the table of contractile force in mN of any desired days (e.g., to GraphPad Prism) for statistical analysis. A minimum of six patches is usually chosen for statistical comparison using repeated measures ANOVA.
Figure 6. Representative contractile force trace of native non-human primate (NHP) heart slice. Readout of the entire culture period (top) or certain timepoints (bottom) can be generated with LabChart Reader (adapted from Poch et al., 2022).
Recipes
Cutting buffer
pH 7.4, adjust with 1 M NaOH. Filter and store up to three months at 4 °C.
Reagent Concentration (g/L) mM
NaCl
KCl
MgCl2
NaH2PO4
Glucose
CaCl2
BDM (2,3-Butanedione)
HEPES
8
0.4
0.2
0.046
2
0.13
3
1.2
136
5.4
1
0.33
10
0.9
30
5
4% agarose preparation
Dissolve agarose at 80 °C for 15 min. Store up to three months at 4 °C. Add 625 μL of HCl for 1 L.
Reagent Concentration (g/L) mM
NaCl
KCl
MgCl2
NaH2PO4
CaCl2
BDM (2,3-Butanedione)
HEPES
8
0.4
0.2
0.046
0.13
3
1.2
136
5.4
1
0.33
0.9
30
5
Culture medium M199
50 mL for 16 slices. Filter, store up to one week at 4 °C.
Reagent Amount
M199
ITS
β-Mercaptoethanol (14 M) 49 mL
500 μL (1:100)
50 μM; mix 1.8 μL of 14M β-Mercaptoethanol stock solution with 0.5 mL of medium. Add 50 μL (= 1:1,000) to M199 culture medium.
Acknowledgments
This work was supported by grants from: European Research Council (ERC) 788381 (to A.M.); the German Research Foundation, Transregio Research Unit 152 (to A.M., K.-L.L.) and 267 (to A.M., K.-L.L.) and DZHK (German Centre for Cardiovascular Research).
We acknowledge the original research papers that this protocol was derived from Poch et al. (2022) and Fischer et al. (2019).
Competing interests
A.D. holds a patent on the technology of biomimetic cultivation and is co-founder and shareholder of InVitroSys GmbH. The remaining authors declare no competing interests.
References
Brandenburger, M., Wenzel, J., Bogdan, R., Richardt, D., Nguemo, F., Reppel, M., Hescheler, J., Terlau, H. and Dendorfer, A. (2012). Organotypic slice culture from human adult ventricular myocardium. Cardiovasc Res 93(1): 50-59.
Cox, L. A., Olivier, M., Spradling-Reeves, K., Karere, G. M., Comuzzie, A. G. and VandeBerg, J. L. (2017). Nonhuman Primates and Translational Research-Cardiovascular Disease. ILAR J 58(2): 235-250.
Fischer, C., Milting, H., Fein, E., Reiser, E., Lu, K., Seidel, T., Schinner, C., Schwarzmayr, T., Schramm, R., Tomasi, R., et al. (2019). Long-term functional and structural preservation of precision-cut human myocardium under continuous electromechanical stimulation in vitro. Nat Commun 10(1): 117.
Liu, H., Bersell, K. and Kühn, B. (2021). Isolation and Characterization of Intact Cardiomyocytes from Frozen and Fresh Human Myocardium and Mouse Hearts. Methods Mol Biol 2158: 199-210.
Perbellini, F., Watson, S. A., Scigliano, M., Alayoubi, S., Tkach, S., Bardi, I., Quaife, N., Kane, C., Dufton, N. P., Simon, A., et al. (2018). Investigation of cardiac fibroblasts using myocardial slices. Cardiovasc Res 114(1): 77-89.
Poch, C. M., Foo, K. S., De Angelis, M. T., Jennbacken, K., Santamaria, G., Bahr, A., Wang, Q. D., Reiter, F., Hornaschewitz, N., Zawada, D., et al. (2022). Migratory and anti-fibrotic programmes define the regenerative potential of human cardiac progenitors. Nat Cell Biol 24(5): 659-671.
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4,751 | https://bio-protocol.org/en/bpdetail?id=4751&type=0 | # Bio-Protocol Content
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Protein Structure Predictions, Atomic Model Building, and Validation Using a Cryo-EM Density Map from Hepatitis B Virus Spherical Subviral Particle
ND Nadia DiNunno *
EB Emily N. Bianchini *
HL Haitao Liu
JW Joseph Che-Yen Wang
(*contributed equally to this work)
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4751 Views: 893
Reviewed by: Prashanth N SuravajhalaAnuj KumarHarpreet Singh
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Original Research Article:
The authors used this protocol in Science Advances Aug 2022
Abstract
Hepatitis B virus (HBV) infection is a global public health concern. During chronic infection, the HBV small-surface antigen is expressed in large excess as non-infectious spherical subviral particles (SVPs), which possess strong immunogenicity. To date, attempts at understanding the structure of HBV spherical SVP have been restricted to 12–30 Å with contradictory conclusions regarding its architecture. We have used cryo-electron microscopy (cryo-EM) and 3D image reconstruction to solve the HBV spherical SVP to 6.3 Å. Here, we present an extended protocol on combining AlphaFold2 prediction with a moderate-resolution cryo-EM density map to build a reliable 3D model. This protocol utilizes multiple software packages that are routinely used in the cryo-EM community. The workflow includes 3D model prediction, model evaluation, rigid-body fitting, flexible fitting, real-space refinement, model validation, and model adjustment. Finally, the described protocol can also be applied to high-resolution cryo-EM datasets (2–4 Å).
Keywords: HBV HBsAg SVP Cryo-EM AlphaFold2 Model building
Background
Hepatitis B surface antigen (HBsAg) is both a constituent of the viral envelope and a well-established serological marker for hepatitis B virus (HBV) infection. It exists in three forms (S stands for small, M for middle, and L for large) that share identical C-termini but different N-termini due to in-frame translation from different start codons. Naturally, all three types of HBsAg are incorporated into the envelope of virions in various amounts. Additionally, HBsAg can assemble into non-infectious subviral particles (SVP), either of filamentous or spherical morphology (Heermann et al., 1984; Ganem and Prince, 2004; Gerlich, 2013). The abundance of spherical SVP in hepatitis B carriers is expected to be 1,000–10,000 times higher than that of infectious virions. Despite this, the role of this large surplus of spherical SVP is unknown (Bruns et al., 1998; Chai et al., 2008; Gerlich, 2013; Hu and Liu, 2017). Among these particles, the S-HBsAg is the most abundant protein constituent.
The HBV SVP has many biomedical applications. It is currently used in a licensed vaccine for preventing hepatitis B infection in newborns. When inserted with a foreign antigenic epitope, the chimeric SVP becomes a platform to induce immune response against medically relevant sequences. This method has been used in the development of the malaria vaccine (Guerra Mendoza et al., 2019; Schuerman, 2019; Ho et al., 2020). Furthermore, HBsAg and SVP also possess immunoinhibitory functions (Fang et al., 2015; S. Liu et al., 2015; Tout et al., 2018; Ho et al., 2020; Kim et al., 2020; Megahed et al., 2020). Recently, a new antiviral strategy involving blocking of the assembly and release of SVPs was found to achieve functional control of HBV infection (Mijočević et al., 2019; Vaillant, 2019). Despite having many roles in biomedical applications and hepatitis B infection, there is no detailed structural information on HBsAg or how it assembles into SVP.
To understand the structural relationship between HBsAg and SVP, we purified spherical SVP from patient serum and investigated the structure using cryo-EM. Since the purification steps and cryo-EM structural determination were described elsewhere (H. Liu et al., 2022), this protocol focuses on the procedure for building the atomic model for S-HBsAg by a combination of AlphaFold2 prediction and a 6.3 Å cryo-EM density map of the spherical SVP from HBV genotype E (H. Liu et al., 2022). We began by using AlphaFold2 to predict the 3D models from HBV S-HBsAg sequence. We then evaluated the result of the prediction using a cryo-EM density map and selected the best model for additional analysis. Finally, we combined knowledge from the literature and used flexible fitting to obtain a 3D model for S-HBsAg. This process utilized AlphaFold2, UCSF ChimeraX (Pettersen et al., 2021), ISOLDE (Croll, 2018), PHENIX, MolProbity (Chen et al., 2010), and Coot to generate the Protein Data Bank (PDB) model and to validate the result. This protocol was used to build a 3D model into a moderate-resolution cryo-EM density map (5–7 Å); it can also be applied to experimental data when a high-resolution cryo-EM map (2–4 Å) is available.
Materials and reagents
EMD-26117 (https://www.ebi.ac.uk/emdb/EMD-26117)
HBV S protein sequence (https://www.ncbi.nlm.nih.gov/protein/1035343445)
Equipment
Linux workstation with dedicated GPU or MacBook (with 64 GB RAM).
Software
AlphaFold2 via ColabFold v1.5.2 (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb)
UCSF ChimeraX v1.5 (https://www.cgl.ucsf.edu/chimerax/)
ISOLDE (https://isolde.cimr.cam.ac.uk/)
PHENIX v1.20.1-4487 (https://phenix-online.org/)
MolProbity (http://molprobity.biochem.duke.edu/)
Coot v0.9.8.2 (https://www2.mrc-lmb.cam.ac.uk/personal/pemsley/coot/)
Note: The selection of software for structural modeling is largely a matter of personal preference. ISOLDE, PHENIX, and Coot can all be used for flexible structure modeling. ISOLDE and Coot provide direct visualization during the modeling process, allowing users to make objective decisions by dragging atoms or residues to specific locations. PHENIX, on the other hand, refines structures based on comprehensive checkups. All software can be used individually or in combination for this purpose. However, we have found that using ISOLDE at the beginning provides quick access to make large structural changes, fix Ramachandran and rotamer outliers, and reduce clashes. Additionally, ISOLDE allows users to "pin" part of the protein model at the original location while making new changes to another part of the structure, which is particularly helpful for regions where the density is ambiguous, such as flexible loops. Once an initial model is generated, PHENIX can be used to fine-tune the structure and fix structural outliers, such as residues with poor bond lengths and angles. Finally, we use Coot in a back-and-forth manner with information obtained from MolProbity to refine the final structure until we achieve a satisfactory result (Figure 1).
Figure 1. Protocol workflow
Procedure
Alphafold2 prediction
AlphaFold2 (Jumper et al., 2021) is available in the ColabFold advanced notebook format at the following website (Mirdita et al., 2022) for easy access: https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb.
Note: In this protocol, we used the newest version of ColabFold. The original paper was run using version v1.1.
The full surface protein sequence of HBV (residues 1–399) can be accessed at NCBI GenBank with Protein ID: ANJ02935. The S-HBsAg protein sequence starts at position 174–399.
Note: You can prepare the partial sequence using the following steps:
Bold font is either input field or button in the software; bold and italic font is the input command.
Go to https://www.ncbi.nlm.nih.gov/protein/ANJ02935 to obtain the HBV large S protein sequence.
Click the FASTA link and copy the sequence.
Go to https://www.bioinformatics.nl/cgi-bin/emboss/extractseq to extract partial sequence.
Paste the copied protein sequence under Input section (Figure 2A).
Go to Required section and input 174-399 under Regions to extract (Figure 2A).
Click the Run extractseq button at the bottom.
Copy only the sequence part for the next step (Figure 2B).
Figure 2. Using extractseq to extract region of interest from the full-length protein sequence. Protein sequence of interest can be obtained at NCBI protein database. (A) Copy the full-length protein sequence in the FASTA format and paste to extractseq web server. Input the region of interest to extract in the field. (B) Result from the extractseq.
Paste the S-HBsAg sequence into query_sequence field at the top region of the page (Figure 3A).
Notes:
For this prediction, we did not consider the oligomeric state of S-HBsAg.
The name for this prediction can be added at the jobname field.
We used only default options in the dropdown menus through the ColabFold notebook.
We did not use template (template_mode: none) or amber (unchecked).
From the menu bar at the top, click Runtime > Run All to initiate the prediction (Figure 3B).
Note: You will see a warning message saying that this notebook was not authored by Google. Please click Run anyway to continue.
Figure 3. Structural prediction using Alphafold2. Alphafold2 prediction can be accessed via ColabFold notebook. (A) The necessary fields to be filled for running Alphafold2 structure prediction. (B) Clicking at Run all will execute all steps automatically. Detailed instructions can be found at the bottom of the notebook.
Once the job finishes, download the results (five output models) in a .zip format and inspect them in ChimeraX.
Notes:
Note that the predicted 3D model is displayed under Display 3D structure section. It shows the highest ranked model by default and can be changed from the dropdown menu rank_num. Areas of the output models predicted with poor confidence are colored in red; areas with high confidence are shown in blue.
All predicted 3D models consist of four helical regions: two long straight helices, one V-shaped helix, and one U-shaped helix. In these five predicted models, the helical regions all have higher confidence. The loop regions that connect two helices were more disordered (Figure S4 in H. Liu et al., 2022).
Because ColabFold webpage can only display one model at a time, we used ChimeraX to visualize all models at once to explicitly compare protein regions that are variant or consistent across them all. Combined with consideration of local confidence scores, this information can aid model interpretation following prediction (Figure 4).
Figure 4. Inspection of the Alphafold2 results. Structural prediction results showed that all models contain similar structural features: four helical regions. Helix 1 and helix 2 are straight, helix 3 has a V shape, and helix 4 has a U shape. The loop regions connected between helices showed substantial differences in all models.
In ChimeraX, we used matchmaker to quickly align all five models together. We found that helical regions had similar structures between all models. On the other hand, the loop region that connects preceding and following helices had distinct connecting angles from model to model. As a result, the secondary structural elements were virtually identical in five predicted models, but the tertiary structural organizations were different.
Proceed to build and refine the best model using a combination of Coot, PHENIX, and ISOLDE.
Model evaluation and rigid-body fitting
Select one model that has the highest similarity to the subunit within the cryo-EM density map of HBV spherical SVP. As the cryo-EM map was refined to 6.3 Å, the helical regions in the cryo-EM map are identifiable by rendering the map at the higher density level. Then, compare the predicted PDB model to the high-density region of the cryo-EM map and select the best model to proceed. To facilitate this process, segment a portion of the density map that contains a unique structural feature (Figure S3 in H. Liu et al., 2022).
Open ChimeraX and open the cryo-EM density map of HBV SVP via command: open emdb:26117 (Figure S2E in H. Liu et al., 2022).
Under Volume Viewer, change the step to 1 and the density Level to 0.013 to render the whole structure.
At the menu bar, select Tool > Volume > Hide Dust.
Set Size limit to 8.4 and click Hide Dust (Figure 5).
Notes:
Inspect the whole particle structure by rotating the map at different angles.
We observed 24 protrusions projected from the particle surface.
For this protocol, we changed the display to a white background. You can do this by going to the menu Presets > Publication 1. We also changed the display to a soft lighting by going to the Graphics tab and clicking Soft.
Figure 5. Structural visualization and analysis in ChimeraX. Hepatitis B virus (HBV) spherical subviral particle rendered in ChimeraX. Inset shows a low pass filtered, computationally segmented dimer density.
Under Volume Viewer, change the density level to 0.021.
Notes:
Inspect the volume to identify the repeating density (subunits).
We found 48 V-shaped densities arranged in parallel on the surface of the particle. At one end of the V-shaped density, it connects to a curved U-shaped density. At the other end of the V-shaped density, it is linked to a long, straight helical density. Following the straight density is another helical density, tilted in a distinct direction. The four helical densities identified from the cryo-EM density map resembles the structural organization we observed earlier in the AlphaFold2 prediction.
By changing the density level back-and-forth, we also observed two V-shaped densities connected to one protrusion.
Taking it all together, we found four connected helical densities organized into one repeated element. Two of the elements connected to one protrusion, indicating that this may be a dimer. In total, 48 repeated elements assembled into a spherical SVP.
If the repeating unit in the structure can be identified, the segmentation tool can be used to isolate only one subunit from the cryo-EM map. This can be done by selecting the menu Volume > Segment Map. In this protocol, we segmented a dimer as the density was unable to be separated to a single subunit state (Figure 5, inset).
Manually move all five predicted PDB models into the corresponding region in the cryo-EM map using the Right Mouse function.
Notes:
We used the V-shaped density as the marker and moved the PDB model into the density.
Move one model at a time using Move model and Rotate model under Right Mouse tab.
When the V-shape helix was roughly covered by the V-shape density from the cryo-EM map, we used an auto-function in ChimeraX to get better precision. This was done by using Tools > Volume Data > Fit in Map function (see result in Figure 6A).
Figure 6. Rigid-body fitting in ChimeraX. (A) Result of rigid-body fitting using the best predicted model into the cryo-EM density map. (B) ChimeraX window saving the resulting model into a new file.
Select each model from the drop-down menu and click Fit button.
Notes:
Steps B6–B8 are called rigid-body fitting.
Not every model can fit into the cryo-EM map very well.
Inspect the fitting result and pick the best fitting model for the subsequent model building.
Save the best PDB model in relation to the cryo-EM map. Go to menu File > Save… (Figure 6B).Notes:
Give a name for the PDB model at File name field.
Select PDB from the Files of type drop-down menu.
Select only the best model in the Save model list.
Check Save relative to model and select the cryo-EM map from the drop-down menu (Figure 6B).
Turn off other models and level only the best fitting model.
Flexible fitting using ISOLDE in ChimeraX (v1.5)
Once the model is selected and moved to the corresponding location of the cryo-EM map using rigid-body movement, work on the regions of the PDB model that fall outside the cryo-EM density using flexible fitting. The goal is to make the backbone of the protein fit into the cryo-EM density, as shown on Figure 7A.
Figure 7. Flexible fitting in ISOLDE. (A) The result of flexible fitting of two subunits into a segmented dimer density. (B) Validation window that is used to fix problematic residues. (C) Ramachandran plot of the protein model.
At the menu, select Tools > General > ISOLDE.
At the Working on field at the top of the window, make sure the best fitting model from above is selected.
At the General tab, under Add map(s) to working model, select the cryo-EM map in the drop-down menu of From loaded volume. This will associate the PDB model to the cryo-EM map and render the map with mesh density around the PDB model.
At Mask and spotlight settings, change Spotlight radius and put 15.0.
Add hydrogens to the PDB by typing addh at the Command field. Output will show that H has been added. Hydrogens can then be hidden by using command hide HC for ease of visualization.
Use the mouse to select one of the helical regions in the PDB model to start with.
Initialize flexible fitting of the model into the map by clicking Start simulation (blue triangle) at the ISOLDE tab in the ChimeraX main window or run the command isolde sim start sel.
Once the simulation starts, use the right mouse button to drag the selected residues into the cryo-EM density. If this does not work, click Tug selection at the ISOLDE tab and use the right mouse button to drag again.
When a satisfactory result is obtained, at Simulation Runtime drop the temperature to 0 K for a few seconds to allow the adjusted structure to settle at the thermal equilibrium and click at the green Stop button. Do this for all the remaining regions of the structure.
Notes:
If a window pops up regarding specifying the disulfide bonds in the metadata when selecting the best PDB model to work with, please select No as we do not have sufficient resolution to resolve the disulfide bond.
For this protocol, keep most parameters unchanged. If customized changes are desired, please refer to each parameter on the ISOLDE website.
If there is a pop-up error stating that there are too many clashes, stop simulation (red stop button to undo changes) and fix individual residues.
If working with helices, it is important to restrain the selection residues to alpha helix. This can be done under the Restraints tab at the Secondary Structure section.
Move the residues outside the cryo-EM map into the density by right-clicking. You can tug just a few residues or the whole selected region.
At the moderate resolution (5–7 Å), only a few residues with bulky side chains can be used as anchors for modeling the rest of residues with higher confidence. Reference from the literature regarding specific mutations or important interactions should be considered at this point to generate a reliable model (Figure S7B in H. Liu et al., 2022).
During modeling, it may be helpful to “pin” the antigenic and cytosolic loops at fixed locations. Because of the low local resolution and high flexibility at these two loop regions, we were unable to model the interactions confidently.
Under Validate tab, try to fix Peptide Bond geometry, Ramachandran Plot, Rotamers, and Clashes (Figure 7B).
Notes:
For Peptide bond geometry, click on the residue in the list, start the simulation, and change it under Rebuild tab or move/rotate it.
For Ramachandran Plot, click on the problematic residue in the plot for a close-up view. Each residue is represented by a circle (Figure 7C). The problematic residues are colored from white (allowed region) to red (outlier). Start the simulation; most times, this will fix the residue. If not, try to move or rotate the residue to fix it. Fix as many outliers as possible.
For Rotamers, work with the outlier residue (red color residue) in the list first. Start the simulation, go to ISOLDE tab, and click at the Preview next icon to change the rotamer. For bulky side chain, make sure the side chain is still inside the cryo-EM density and that it does not cause a new clash with neighboring residues after you change it. Click Set coords once the result is satisfying. The residues in red (outliers) should all be corrected. Others (disfavored residues) should also be corrected, as much as possible. Clicking the Update button will refresh the list.
For Clashes, start with the biggest clashes first by clicking on the residue in the list and inspecting the region of the residue that is involved in the clash. If the clash is caused by the side chain, change it to other possible rotamers. If the clash is involved in the carbon backbone, move or rotate the residue to resolve the clash.
At moderate resolution, we can only fix clashes at the regions that have good cryo-EM densities, e.g., the helical region. At the loop or disordered regions, it becomes more difficult to fix clashes; thus, we did not attempt to fix clashes at these regions of the model.
Prioritize fixing clashes when going in between Rotamers and Clashes.
Once the residue is successfully corrected, click Update list to remove it from the list.
Note that some bond lengths and bond angles in ISOLDE cannot be fully corrected. They need to be refined or fixed in the next steps.
Save the corrected PDB model into a new file.
Because the building block of spherical HBV SVP is a dimer, we should take the PDB model generated from the steps above and flexibly fit it into the neighboring subunit that forms the same protrusion.Notes:
Open the file saved in step C11.
Move the model into the density of the neighboring subunit using rigid-body fitting.
Flexibly adjust the model with ISOLDE.
Combine two PDB models into one file by using command combine #PDB1 #PDB2 (PDB1 should be the ID number for the PDB in step C11 and PDB2 should be the ID number from this step).
Real-space refinement of flexible fitting result using PHENIX (v1.20.1-4487)
Even though it is possible to manually adjust the model using ISOLDE in ChimeraX, it is better to further refine and optimize the results using the algorithm that considers the chemical principle. To do so, use PHENIX and supply the PDB model (dimer) and the cryo-EM map. For the best results, the segmented dimer cryo-EM map should be used here. To prepare files for PHENIX and Coot, the coordinate system needs to be adjusted.
In ChimeraX, open both the cryo-EM map (from section B) and the combined PDB file (from section C).
At the menu, go to Volume > Map Coordinates. Change Original index to 0. The cryo-EM map should be off from the PDB model.
Perform rigid-body fitting by moving the PDB model into the cryo-EM map. Be cautious and make sure that the subunit is correctly fit into the density. Once this is done, save the PDB model into a new file.
Start PHENIX, add a New project, name Project ID, and pick a folder for Project directory. The sequence in FASTA format can also be added, but it is not required.
Go to Cryo-EM: Map analysis, symmetry, manipulation task at the right-hand side, click Refinement, and select Real-space refinement.
Specify Job title.
Note that it does not have to be unique, but new files will be created using this as part of the file name.
At Input/Output tab, click Add file button to add files (PDB model and cryo-EM map). Check the box for ignore symmetry conflicts and enter map resolution 5.5 here.
In Refinement Settings, make sure that minimization_global, morphing, simulated_annealing, adp, occupancy, and ngh_flips are selected. Hit Run (Figure 8).
Figure 8. Setup window of PHENIX real-space refinement
Notes:
Rigid_body and local_grid_search can be unchecked to save time as the model was fitted to an approximate location in the earlier steps.
Other parameters can be used as default or changed to tune up the process.
If running with multiple CPU, specify the number of processors in Nproc box under Other Options.
Completed files have “real_space_refined” as a suffix in .pdb format.
Any bad bond length and angle should be corrected here.
MolProbity (v4.5, http://molprobity.biochem.duke.edu/)
We used MolProbity to validate the modeling result. Although PHENIX has a built-in validation tool from MolProbity, we found that the report created from the MolProbity website was better illustrated and allowed easy sorting.
Open a web browser and go to the MolProbity website at http://molprobity.biochem.duke.edu/.
Use Choose File to upload the resulting PDB model from PHENIX. Select PDB coords for the Type and click Upload.
Select Add hydrogens and choose the Asn/Gln/His flips method. Select Electron-cloud x-H for x-H bond-length and click Start adding H.
Notes:
If residue flips are suggested, a list of changes are shown. Check the explanation and the before and after flip result to decide if the changes are leading to a better result.
If changes are made here, click Regenerate H, applying only selected flips.
A new corrected PDB file will be downloaded.
Select Analyze all-atom contacts and geometry.
Use default options and click Run programs to perform these analyses.
A summary of data statistics will be shown. Click at Multi-criterion chart to see a full list.
Notes:
The goal is to fix as many outliers in the residues as possible. For loop and disordered regions, it should be difficult. On the other hand, if the cryo-EM density map can be resolved to better than 3 Å, all statistics in the table should be able to correct to the green color.
After ISOLDE and PHENIX, we should see only some residues remaining to be corrected. We will move to Coot to fix problematic residues.
Coot (v0.9.8.2)
Go to File > Open Coordinates and Open Map to open the PDB model and cryo-EM map, respectively.
Notes:
Read the PDB model with added hydrogen atoms.
Click the Display Manager button beneath the menu bar to change the display settings. We normally colored each chain in the PDB model with different colors by selecting Bonds (Colour by Chain) in the drop-down menu in the Molecules section. Add the plasmids into competent cells.
Go to the right-hand side menu bar and select Refine/Regularize Control… (R/RC) to open a dialog window. Select Use Torsion Restraints and Ramachandran Restrains under For Regularization and Refinement. Click Estimate to calculate the Weight Matrix for cryo-EM map. Click OK to finish.
For Ramachandran outliers, go to Validate > Ramachandran plot. Identify and correct all outliers in the plot.
Notes:
Look at the Ramachandran plot to identify the outliers; they are colored in red (Figure 9A).
Figure 9. Validation tools in Coot. (A) Ramachandran plot shows outliers and the summary of all residues. (B) Rotamer analysis shows outlier residues (red color) and residues in the allowed region.
Select the outlier by clicking on the red colored symbol. The viewer will then bring a close-up view at the selected residues.
Fix the outlier residue by selecting one preceding and one following residue by using Real Space Refine Zone at the right-hand side menu bar. As an example, if Ile 68 is an outlier, first click Real Space Refine Zone button and then click at Pro 67 and Cys 69.
A refinement result window pops up after it attempts to refine these residues. Check the refinement result at the top of the pop-up window. If all criteria are green, accept the refinement result. If one or more categories are red or yellow, reject refinement and change it again.
On some occasions, clicking Flip This Peptide or Flip Next Peptide in the pop-up window can resolve the problem.
Use the list from MolProbity to easily go through the unfavored residues as well.
For HBV SVP, we attempted to fix the helical region only.
For Bond lengths and Bond angles outliers identified in MolProbity, depending on the location of the bond, these can be corrected by using Read Space Refine Zone and clicking at the same residue twice to fix it. In some scenarios, it requires selecting preceding or following residues to fix it.
For Rotamer outliers, go to Validate > Rotamer Analysis and compare with MolProbity results (Figure 9B). Click on the poor rotamer residue (red and orange color) in the histogram plot for a close-up view in the 3D model. Click on Rotamers… at the right-hand side menu bar, click on the desired residue, and toggle between different rotamers. Select the rotamer closest in structure to the original conformation or within the cryo-EM density.
Notes:
Do not cause a new clash to neighboring atoms.
Continue until all red and orange rotamers have been repaired in Unusual Rotamer Graphs (histogram plot) window.
Save intermediate PDBs often without overwriting older versions.
Repeatedly alternate between Coot and MolProbity until all categories assessed in MolProbity validation appear green.
At the intermediate resolution, the information is insufficient to build a reliable atomic model directly from the cryo-EM density. Therefore, the side chain orientation and the atom locations were adjusted locally according to the published literature on the protein–protein interactions at this step.
Download the final PDB model without hydrogen atoms for PDB deposition.
Acknowledgments
The research was supported by the startup fund from The Pennsylvania State University College of Medicine, NIH grant R21-AI1641191, and R37AI043453. We thank Dr. A. Zlotnick for insightful discussion. This protocol was derived from the original research study published by H. Liu et al. (2022).
Competing interests
The authors declare no competing interests.
References
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Chai, N., Chang, H. E., Nicolas, E., Han, Z., Jarnik, M. and Taylor, J. (2008). Properties of subviral particles of hepatitis B virus. J Virol 82(16): 7812-7817.
Chen, V. B., Arendall 3rd, W. B., Headd, J. J., Keedy, D. A., Immormino, R. M., Kapral, G. J., Murray, L. W., Richardson, J. S. and Richardson, D. C. (2010). MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr D Biol Crystallogr 66(Pt 1):12-21.
Croll, T. I. (2018). ISOLDE: a physically realistic environment for model building into low-resolution electron-density maps. Acta Crystallogr D Struct Biol 74(Pt 6): 519-530.
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Ganem, D. and Prince, A. M. (2004). Hepatitis B virus infection--natural history and clinical consequences. N Engl J Med 350(11): 1118-1129.
Gerlich, W. H. (2013). Medical virology of hepatitis B: how it began and where we are now. Virol J 10: 239.
Guerra Mendoza, Y., Garric, E., Leach, A., Lievens, M., Ofori-Anyinam, O., Pirçon, J. Y., Stegmann, J. U., Vandoolaeghe, P., Otieno, L., Otieno, W., et al. (2019). Safety profile of the RTS,S/AS01 malaria vaccine in infants and children: additional data from a phase III randomized controlled trial in sub-Saharan Africa. Hum Vaccin Immunother 15(10): 2386-2398.
Heermann, K. H., Goldmann, U., Schwartz, W., Seyffarth, T., Baumgarten, H. and Gerlich, W. H. (1984). Large surface proteins of hepatitis B virus containing the pre-s sequence. J Virol 52(2): 396-402.
Ho, J. K., Jeevan-Raj, B. and Netter, H. J. (2020). Hepatitis B Virus (HBV) Subviral Particles as Protective Vaccines and Vaccine Platforms. Viruses 12(2): 126.
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Jumper, J., Evans, R., Pritzel, A., Green, T., Figurnov, M., Ronneberger, O., Tunyasuvunakool, K., Bates, R., Žídek, A., Potapenko, A., et al. (2021). Highly accurate protein structure prediction with AlphaFold. Nature 596(7873): 583-589.
Kim, J. H., Ghosh, A., Ayithan, N., Romani, S., Khanam, A., Park, J. J., Rijnbrand, R., Tang, L., Sofia, M. J., Kottilil, S., et al. (2020). Publisher Correction: Circulating serum HBsAg level is a biomarker for HBV-specific T and B cell responses in chronic hepatitis B patients. Sci Rep 10(1): 5947.
Liu, H., Hong, X., Xi, J., Menne, S., Hu, J. and Wang, J. C. (2022). Cryo-EM structures of human hepatitis B and woodchuck hepatitis virus small spherical subviral particles. Sci Adv 8(31): eabo4184.
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© 2023 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 > Microbe-host interactions > Virus
Biophysics > Electron cryotomography > 3D image reconstruction
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In the original version of the protocol, (https://bio-protocol.org/e1913), the formulas for MDA, SOD, POD, CAT, and GST calculation were incorrect. In all the mentioned formulas, Cp should be replaced with (Cp × V).
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After publishing the protocol (https://bio-protocol.org/e3533), we realized that there is a typo in the Recipes. In section 3, under the scopolamine working solution, we mentioned adding 950 µl of scopolamine stock solution to 50 µl of 0.9% sterile saline; it should be 50 µl of scopolamine stock solution and 950 µl of 0.9% sterile saline.
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4,754 | https://bio-protocol.org/en/bpdetail?id=4754&type=0 | # Bio-Protocol Content
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Visualizing Loss of Plasma Membrane Lipid Asymmetry Using Annexin V Staining
JB Julia F. Baum *
HU Huriye D. Uzun *
TP Thomas Günther Pomorski
(*contributed equally to this work)
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4754 Views: 1126
Reviewed by: Alexandros AlexandratosVishal NehruEVANGELOS THEODOROUTakashi Nishina
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Original Research Article:
The authors used this protocol in Journal of Cell Science Mar 2022
Abstract
Loss of plasma membrane lipid asymmetry contributes to many cellular functions and responses, including apoptosis, blood coagulation, and cell fusion. In this protocol, we describe the use of fluorescently labeled annexin V to detect loss of lipid asymmetry in the plasma membrane of adherent living cells by fluorescence microscopy. The approach provides a simple, sensitive, and reproducible method to detect changes in lipid asymmetry but is limited by low sample throughput. The protocol can also be adapted to other fluorescently labeled lipid-binding proteins or peptide probes. To validate the lipid binding properties of such probes, we additionally describe here the preparation and use of giant unilamellar vesicles as simple model membrane systems that have a size comparable to cells.
Key features
• Monitoring loss of lipid asymmetry in the plasma membrane via confocal microscopy.
• Protocol can be applied to any type of cell that is adherent in culture, including primary cells.
• Assay can be adapted to other fluorescently labeled lipid-binding proteins or peptide probes.
• Giant unilamellar vesicles serve as a tool to validate the lipid binding properties of such probes.
Graphical overview
Imaging the binding of fluorescent annexin V to adherent mammalian cells and giant vesicles by confocal microscopy. Annexin V labeling is a useful method for detecting a loss of plasma membrane lipid asymmetry in cells (top image, red); DAPI can be used to identify nuclei (top image, blue). Giant vesicles are used as a tool to validate the lipid binding properties of annexin V to anionic lipids (lower image, red).
Keywords: Confocal microscopy Giant vesicle Lipid asymmetry Lipid-binding protein Mammalian cells Plasma membrane
Background
A characteristic feature of many biological membranes is that their phospholipids are asymmetrically distributed across the lipid bilayer, a phenomenon known as transbilayer lipid asymmetry. A prominent example is the plasma membrane of animal cells, in which the phospholipids phosphatidylcholine and sphingomyelin are concentrated in the exoplasmic leaflet, whereas the aminophospholipids phosphatidylserine (PS) and phosphatidylethanolamine (PE) are restricted to the cytosolic leaflet (van Meer et al., 2008). Transbilayer lipid asymmetry is essential for several vital cellular functions, including the regulation of membrane protein activity, signaling, and vesicle formation in the secretory and endocytic pathways (Sprong et al., 2001; Ewers and Helenius, 2011; van Meer, 2011; Sebastian et al., 2012). In animals, loss of transbilayer lipid asymmetry has been linked to processes such as blood coagulation (Lentz, 2003; Jackson, 2011), cell adhesion (Schlegel et al., 1985; Malhotra et al., 1996; Wautier et al., 2011), macrophage recognition (Krahling et al., 1999), apoptosis (Bevers and Williamson, 2016), and myotube formation (van den Eijnde et al., 2001). The establishment and regulation of lipid asymmetry are therefore crucial for cells, and several membrane proteins have evolved to fulfill the function of cross-bilayer phospholipid transporters, comprising lipid flippases, floppases, and scramblases (Hankins et al., 2015; Ristovski et al., 2021).
Several methods have been developed to analyze the loss of phospholipid asymmetry in the plasma membrane of eukaryotic cells. These include chemical approaches using e.g., trinitrobenzene sulfonic acid or fluorescamine, which covalently react with amino groups of lipids and proteins (Marinetti et al., 1976; Pomorski et al., 2003). As the probes are membrane impermeant, only aminophospholipids exposed to the cell surface are modified and can then be detected by thin-layer chromatography or mass spectrometry. However, this approach is not suitable for live-cell imaging. More recent methods are based on fluorescently labeled lipid-binding proteins that can be added to the cells. One example is the PS-specific probe lactadherin, which binds to PS with a nanomolar affinity and without the need for cofactors (Waehrens et al., 2009). Another example is annexin V, a member of the annexin family of Ca2+-dependent, non-covalent lipid-binding proteins. Annexin V binds negatively charged lipids with relatively high affinity and is used extensively for the detection of exofacial PS by flow cytometry or microscopy (Koopman et al., 1994; Vermes et al., 1995; Tait et al., 2004). A new generation of fluorescent probes is based on cyclic peptides that successfully mimic the function of lipid-binding proteins and benefit from their small size, ease of labeling, and cofactor-free PS recognition (Hanshaw et al., 2005; DiVittorio et al., 2006; Zheng et al., 2011).
In this protocol, we describe the use of fluorescently labeled lipid-binding protein sensors to detect the loss of lipid asymmetry in living cells by fluorescence microscopy, exemplified on mouse C2C12 wild-type myoblasts and corresponding knockout cells lacking the P4-ATPase flippase subunit CDC50A (also known as TMEM30A). Deletion of CDC50A results in loss of the aminophospholipid flippase activity and constitutive loss of plasma membrane lipid asymmetry (Grifell-Junyent et al., 2022). The approach is illustrated using commercially available annexin V conjugated to Alexa Fluor 568, but other fluorescently labeled lipid-binding proteins or peptide probes can also be used.
To validate the specificity and sensitivity of such lipid binding probes, we also describe here the use of giant unilamellar vesicles (GUVs) as simple model membrane systems. One of the major advantages of using GUVs as model membrane systems is their similarity in size to cells. This allows GUVs to be observed directly under the microscope, making them a convenient and accessible tool for lipid-binding studies. By preparing GUVs with defined lipid compositions, the specificity and sensitivity of lipid-binding probes can be evaluated and their accuracy and reproducibility in live cell experiments can be ensured (Weingärtner et al., 2012; Chandra and Datta, 2022). GUVs with defined lipid compositions can be prepared by various methods, including swelling, PVA or agarose swelling, and electroformation using indium tin oxide glass slides and droplet transfer methods (Angelova and Dimitrov, 1986; Weinberger et al., 2013; Bhatia et al., 2015; Shimane and Kuruma, 2022). In this study, we describe the swelling method due to its simplicity. For alternative preparation methods, the reader is referred to other bio-protocols (Parigoris et al., 2020; Mathiassen and Pomorski, 2022). Our protocol provides a reliable and efficient method for detecting loss of lipid asymmetry in living cells and can be adapted for use with a variety of lipid-binding proteins.
Materials and reagents
Mammalian cell culture
In this study, we used mouse myoblast cells (C2C12; cell number: ACC 565, DSMZ Braunschweig, Germany) and the corresponding knockout cells lacking CDC50A (Grifell-Junyent et al., 2022) that were cultured in growth medium (see Recipe 1). Optimal culture media and conditions may differ for other cell lines.
Basal cell culture medium for growth (e.g., high glucose DMEM, without pyruvate; Sigma-Aldrich, catalog number: D5796), store at 4 °C
Ethanol absolute ≥ 99.8% (VWR, catalog number: 20821.321)
Ethylene glycol-bis(2-aminoethylether)-N,N,N′,N′-tetraacetic acid (EGTA) (e.g., Sigma-Aldrich, catalog number: E4378)
Fetal bovine serum (FBS), heat inactivated before use (e.g., Capricorn Scientific, catalog number: FBS-11A), store at -20 °C
Hanks’ balanced salt solution, Ca2+ and Mg2+ free (HBSS) (e.g., Sigma-Aldrich, catalog number: H6648), store at 4 °C
35 mm polymer bottom dishes (e.g., Ibidi, catalog number: 81156)
1.5 mL microcentrifuge tubes (Sarstedt, catalog number: 72.690.001)
Penicillin-streptomycin, 100× solution (e.g., Sigma-Aldrich, catalog number: P4333), store at -20 °C
Pipette controller (e.g., accu-jet pro, Brand, catalog number: 263 00)
Polypropylene tubes, 15 mL capacity (e.g., Falcon tubes, Sarstedt, catalog numbers: 62.554.502 and 62.547.254)
Sterile serological pipettes (e.g., Serological pipettes of 5, 10, and 25 mL; Sarstedt, catalog numbers: 86.1253.001, 86.1254.001, and 86.1685.001)
Sterile culture vessels T-75 flasks (e.g., Sarstedt, catalog number: 83.3911)
Trypsin-EDTA solution (e.g., Sigma-Aldrich, catalog number: T3924), store at -20 °C
Trypan Blue solution, 0.4% (e.g., Thermo Fischer Scientific, catalog number: 15250061)
Growth medium (see Recipe 1)
For annexin V labeling
Annexin V conjugated to Alexa Fluor 568 (e.g., Roche, catalog number: A13202), store at 4 °C
4’,6-Diamidino-2-phenyl-indol-dihydrochlorid (DAPI) (e.g., Sigma-Aldrich, catalog number: D9542)
Dead cell staining reagents, e.g., SYTOX Blue (Thermo Scientific, catalog number: S34857)
Ice
Tyrode’s balanced salt solution with Ca2+ (TBSS + Ca2+; see Recipe 2), store at 4 °C
Tyrode’s balanced salt solution without Ca2+ (TBSS - Ca2+; see Recipe 3), store at 4 °C
DAPI stock solution (1 mg/mL) (see Recipe 4)
Note: This procedure has also been successfully performed using FITC-labeled lactadherin (e.g., Haematologic Technologies, catalog number: BLAC-FITC).
For the preparation of giant unilamellar vesicles (GUVs)
1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) (Avanti® Polar Lipids, catalog number: 850375)
1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) (Avanti® Polar Lipids, catalog number: 850725)
1,2-dioleoyl-sn-glycero-3-phospho-L-serine (sodium salt) (DOPS) (Avanti® Polar Lipids, catalog number: 840035)
Calcium chloride (CaCl2) (Grüssing, catalog number: 10043-52-4)
Chloroform, ethanol-stabilized and certified for absence of HCl (Sigma-Aldrich, catalog number: 32211-M)
Detergent/soap
Ethanol, 70% (Sigma-Aldrich, catalog number: 64-17-5)
Glucose (Duchefa Biochemie, catalog number: G0802.5000)
HEPES (Carl Roth, catalog number: 7365-45-9)
Ice
Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma-Aldrich, catalog number: 7791-18-6)
Methanol ≥ 99.8% (VWR, catalog number: 67-56-1)
Potassium hydroxide (KOH) (Sigma-Aldrich, catalog number: 1310-58-3)
Potassium chloride (KCl) (Merck, catalog number: 7447-40-7)
Sodium chloride (NaCl) (Carl Roth, catalog number: 7647-14-5)
Sucrose (Duchefa Biochemie, catalog number: S0809.5000)
Lipid stock in chloroform (see Recipe 5)
Swelling buffer (320 mM sucrose) (see Recipe 6)
Binding buffer (see Recipe 7)
Note: Store or keep all reagents at room temperature, except indicated items. All buffers are prepared the day before and stored at 4 °C.
Equipment
General
Analytical balance (e.g., Sartorius Entris-I II, 220 g/0.1 mg; Buch Holm, catalog number: 4669128)
Computer with monitor (e.g., DELL U2415)
Confocal laser scanning microscope (e.g., Leica TCS SP8 confocal laser scanning microscope, Leitz, Wetzlar, Germany, equipped with a 63×/1.20 water objective)
Eppendorf Research® plus pipettes P20, P200, P1000 (Eppendorf, catalog numbers: 3123000039, 3123000055, 3123000063)
Eppendorf tube, 2 mL (Merck, catalog number: EP0030120094)
Freezers -20 °C and -80 °C
Magnetic stirrer (e.g., IKAMAG®, DREHZAHL ELECTRONIC, IKA, Staufen im Breisgau, Germany)
Magnets
pH meter (pH-Meter 761 Calimatic, Knick, Berlin, Germany)
Pipette tips 10, 200, 1,000 μL (Sarstedt, catalog numbers: 70.760.002, 70.3030.020, 70.3050.020)
Refrigerator (5 °C)
Water distillation system
For cell culture
Autoclave sterilizer (e.g., Systec VX-65, Systec, Linden, Germany)
Biological safety cabinet certified for handling biological materials (e.g., Herasafe KSP Class II Biological Safety Cabinets, Thermo Fisher Scientific)
Centrifuge with rotor for 15 and 50 mL polypropylene tubes (e.g., Eppendorf 5810 R; Wesseling, Germany)
Incubator with humidity and gas control to maintain 37 °C and 95% humidity in an atmosphere of 5% CO2 in air (e.g., Binder, Tuttlingen, Germany)
Inverted phase contrast microscope equipped with a 10× objective (HI PLAN I 10×/0.22 PH1; Leica DMi1, Mannheim, Germany)
Neubauer counting chamber (improved dark lines, 0.1 mm) and cover glasses (20 mm × 26 mm × 0.4 mm)
Water bath (e.g., WPE45 Memmert, Schwabach, Germany) for mammalian cells and for NBD-lipid labeling (Julabo CORIO C-BT5, catalog number: 9011305)
For preparation of GUVs
Cover glass slides (26 mm × 76 mm, #1.5, Thermo Fisher Scientific, Life Technologies Corporation Eugene)
Flow cabinet to work with organic solvents
Glass beads, 3 mm (Merck, catalog number: 104015)
Glass desiccator Boro 3.3 with a socket in the lid, 20 cm, including stopcock (Brand, catalog number: 65238)
Glass pipettes (e.g., graduated pipettes BLAUBRAND® Type 3 Class AS, 10 mL, graduation: 10 mL; Carl Roth, catalog number: HXT8.1)
Glass slide (Thermo Scientific, microscope slides 76 mm × 26 mm, catalog number: MEZ 101026)
Glass vials (Rotilabo® screw neck ND8 vials, brown/white glass, 1.5 mL; Carl Roth, catalog number: KE30.1) with screw caps (without a borehole, without septum, PP, black, ND8; Carl Roth, catalog number: KE39.1)
Glass tubes (Carl Roth, catalog number: DURAN C208.1)
Hamilton 700 Series syringes 25, 100, 1,000 μL (Hamilton Company, Nevada, USA)
High vacuum grease (DOW CORNING, 65201 Wiesbaden, made in USA, Artwork Nr. 0315)
Ice bucket (e.g., Magic Touch 2TM ice bucket with lid; Sigma-Aldrich, catalog number: BAM168072002)
O-ring (28 mm × 1 mm, Nanion Technologies, München)
Parafilm (PARAFILM® M; Sigma-Aldrich, catalog number: P7793-1EA)
Rotavapor® R-100 Evaporator with I-100 Controller and V-100 vacuum pump (Flawil, Switzerland)
Scissors
Ultra-violet/ozone probe and surface decontamination unit (e.g., Novascan Technologies Inc., Boone, IA, USA)
Vortex mixer (e.g., Vortex Genie 2 Scientific Industries Inc., catalog number: SI-0236)
Vacuum Pump V-100 with Interface I-100 (Buchi, catalog numbers: 11593636 and 11593655D)
Wipes (Precision Wipes, KIMTECH Science, Kimberly-Clark® Professional, catalog number: 7552)
Software
ImageJ (Wayne, Rasband, S., U. S. National Institutes of Health, Bethesda, Maryland, USA, https://imagej.nih.gov/ij/index.html , version v.153q)
Leica Application Suite AF (LAS AF, Leitz, Wetzlar, Germany)
Microsoft Excel (Microsoft Corporation, 2018)
PowerPoint (Microsoft Corporation, 2018)
OriginPro (OriginLab, 2023)
Procedure
The following procedure outlines four main steps: (A) preparation of mammalian cells, (B) cell counting, (C) annexin V staining of adherent cells, and (D) preparation of GUVs. The last step was originally applied to study the binding specificity of annexin V to different membrane lipid compositions (Weingärtner et al., 2012; Grifell-Junyent et al., 2022). We recommend this analysis before applying new probes in cell studies.
Preparation of mammalian cells
Grow adherent cells in sterile culture vessels (T-75 flask) in growth medium (see Recipe 1) in a tissue culture incubator (37 °C, 5% CO2, 95% humidity) until they reach ~60%–70% confluency.
Caution: C2C12 cells will differentiate if grown too confluent and start to fuse. Differentiation and fusion are accompanied by transcriptional changes, which can lead to different results.
Aspirate and discard growth media with sterile serological pipette.
Wash cells twice with 5 mL of HBSS (Ca2+ and Mg2+ free, pre-warmed at 37 °C) using a sterile serological pipette.
Add 1.5 mL of trypsin-EDTA solution (pre-warmed at 37 °C) using a sterile serological pipette and incubate the T-75 flasks in a tissue culture incubator (37 °C, 5% CO2, 95% humidity). Tilt the vessel back and forth a few times to make sure the thin layer of trypsin is evenly spread.
After 5 min, check for detachment by gently tilting vessel and/or observing under the microscope. If all cells have not detached in 5 min, incubate an additional 1–2 min and check again. Continue to incubate and check as necessary, only until cells are no longer attached to the plate surface.
Caution: Avoid prolonged incubation period with trypsin-EDTA solution.
Stop trypsinization by adding 7.5 mL of growth medium (pre-warmed at 37 °C, see Recipe 1) to the cell suspension.
Transfer the cell suspension into a 15 mL Falcon tube and set aside 100 μL in a 1.5 mL microcentrifuge tube for cell counting, e.g., using the hemocytometer (see section B).
Centrifuge cells in the 15 mL Falcon tube at 300× g for 5 min at room temperature and discard the supernatant to remove the trypsin-EDTA containing medium from the cells.
Add 10 mL of fresh growth medium (pre-warmed at 37 °C, see Recipe 1) to the cell pellet and re-suspend completely by gently pipetting up and down using a serological pipette.
Caution: Cells in suspension settle quickly. After counting, we recommend gently re-suspending the cell suspension approximately every 2–3 min when seeding multiple dishes.
After counting (see section B), seed 1.5 × 104 cells per 35 mm polymer bottom dishes and add growth medium (pre-warmed at 37 °C, see Recipe 1) up to a final volume of 1 mL per dish.
Note: Prepare sufficient Petri dishes for control samples (see section C). We used a low cell number for seeding because C2C12 is a fast-growing cell line (doubling time: ~20 h) and this cell number guarantees that single cells are still present when the assay is performed the next day.
Keep the cells in a tissue culture incubator (37 °C, 5% CO2, 95% humidity) overnight.
Cell counting (supplemental; if information on cell counting is not required, proceed to section C)
The purpose of this step is to quantify the cell concentration to resuspend the cells at the appropriate concentration for the assay. We routinely use the trypan blue hemocytometer assay. Alternative cell counting methods such as automatic cell counters may be used.
Prepare the hemocytometer by cleaning the chambers and coverslip with ethanol. Dry the hemocytometer by using lint-free tissue. Place the glass coverslip over the counting chambers.
Note: The correct placement is indicated by the appearance of the Newton rings.
Add 100 μL of 0.4% trypan blue solution to 100 μL of cell suspension (step A7) to obtain a 1:1 dilution using a P200 Eppendorf pipette.
Load the hemocytometer with 10 μL of cell suspension per counting chamber with a P20 Eppendorf pipette and examine immediately under an inverted phase contrast microscope at low magnification (e.g., 5–10× magnification).
Critical: To ensure accurate results, it is important to avoid over- or underfilling the cell suspension chamber.
Count the number of viable (seen as bright cells) and non-viable cells (stained blue) in the large outer quadrants.
Calculate the percentage of viable cells: % viable cells = [1.00 - (number of blue cells ÷ number of total cells)] × 100. Cell viability should be at least 95%.
Calculate the cell concentration based on the premise that each square accounts for a volume of 10-4 mL of cell suspension.
To obtain the total number of viable cells per milliliter of aliquot, multiply the total number of viable cells by 2 (the dilution factor for trypan blue) and the correction factor of 104 (volume of each square).
Annexin V staining of adherent cells
We suggest that at least four samples are prepared (Figure 1): i) a negative control without staining to determine background fluorescence, ii) a sample stained with DAPI only, iii) a sample to be stained with DAPI and annexin V in the presence of calcium, and iv) a sample to be stained with DAPI and annexin V in the absence of calcium.
Figure 1. Schematic illustration of the annexin V binding assay. Preparation of a negative control without DAPI and annexin V staining, a control with DAPI-stained cells, and cells stained with DAPI and annexin V conjugated to Alexa Fluor 568 for 10 min on ice with TBSS with or without calcium, respectively. The negative control without staining is used to assess the level of background fluorescence in the sample. This is important because even in the absence of a fluorescent stain, there may still be some level of background fluorescence present due to autofluorescence or other sources. DAPI staining (at a final concentration of 10 μg/mL) is used for visualization of nuclei and cell counting. Non-specific binding of Ca2+-dependent annexin V is tested in the same assay by using TBSS without Ca2+.
Prior to the start of the assay, prepare TBSS with and without calcium (± Ca2+; see Recipes 2 and 3) and a DAPI stock solution (see Recipe 4).
Note: Annexin V requires the presence of Ca2+ to bind to PS. TBSS without Ca2+ is used as a negative control.
To prepare the cells for labeling, carefully aspirate and discard the growth medium using a P1000 Eppendorf pipette.
Wash cells twice with 1 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette.
After washing, add 0.5 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette.
Add 5 μL of the annexin V conjugated to Alexa Fluor 568 using a P20 Eppendorf pipette and incubate for 10 min on ice in the dark.
Note: Skip steps C5–C8 for the negative control (i); skip steps C5–C7 for the sample only stained with DAPI (ii).
Remove staining solution and wash the cells twice with 1 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette.
After washing, add 0.5 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette.
Add DAPI stock solution to a final concentration of 10 μg/mL (stock 1 mg/mL: add 5 μL using a P20 Eppendorf pipette). Incubate the cells at room temperature for 10 min in the dark.
Note: At the concentration used here, DAPI stains the nucleus of both live and dead cells and is used for the visualization of nuclei and cell counting. When using DAPI for dead cell staining, a final concentration of 0.1 μg/mL is recommended. Alternatively, other dead cell staining reagents such as SYTOX Blue can also be used.
Remove staining solution and wash cells twice with 1 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette.
Add 1 mL of TBSS (± Ca2+; cooled on ice, see Recipes 2 and 3) using a P1000 Eppendorf pipette and image on the microscope.
Preparation of GUVs
Prepare the stock and working solutions of the lipids
Clean the Hamilton syringes by flushing them ten times with chloroform:methanol (1:1, v:v).
Caution: Chloroform is a hazardous solvent. Conduct all work in a fume hood, while wearing proper protective clothing.
To have a 5 mg/mL lipid film, transfer 500 μL of a 10 mg/mL DOPC stock solution (see Recipe 5) into a round bottom glass tube on ice using Hamilton syringes. For 5 mg/mL DOPC:DOPE or DOPE:DOPS lipid mix, add 9 mol DOPC and 1 mol DOPE or DOPS (lipid stocks, see Recipe 5). In this study, the lipid mixtures DOPC:DOPE and DOPC:DOPS will be named only DOPE and DOPS, respectively.
Caution: Avoid any use of plastic ware when handling organic solvents. For more complex or different lipid mixtures, the volume of used lipids needs to be adjusted.
Evaporate the organic solvent at room temperature under reduced pressure in a rotary evaporator at 250 mbar for 2–4 h followed by evaporation at ~10 mbar for 15 min (see Figure 2A and 2B).
Store the lipid film at -20 °C until use.
Dissolve the lipid film in 1 mL of chloroform:methanol (1:1; v:v).
Transfer into a glass vial with screw caps closed with parafilm and store it at -20 °C.
Cleaning of the glass slides
Clean the glass slides with detergent, deionized water, and 70% ethanol.
Dry the slides with wipes.
Place glass slides in the UV/ozone cleaner.
Run the UV/ozone cleaner for 30 min.
Caution: The UV/ozone cleaner must be placed under a fume hood.
Turn off the UV/ozone cleaner and wait at least 15 min before opening the chamber.
Preparation of GUVs
Glue the O-ring using vacuum grease onto the cleaned side of one of the glass slides to have a tight, closed chamber. Apply 35–40 drops of 1 μL of lipid mix inside the O-ring under the fume hood (see Figure 2C).
Evaporate the solvent under vacuum for 30–60 min at 250 mbar in a desiccator.
Fill the O-ring with swelling buffer (see Recipe 6) and place the other glass slide on top with the cleaned side facing downwards.
Place the chamber in a dark room for 2–4 h at room temperature (see Figure 2D).
Carefully tap on each side of the chamber.
Transfer the GUVs to a 2 mL tube by removing the glass slide and O-ring.
Store the GUVs covered in aluminum foil at room temperature.
Figure 2. Generation of giant unilamellar vesicles (GUVs) from lipid mixtures. (A, B) Preparation of the lipid mixture. The desired volume and type of lipid is mixed in chloroform:methanol (1:1; v:v) in a glass tube and (I) the solvent is evaporated in rotary evaporator under the reduced pressure of 250 mbar for 2–4 h; (II) the resulting thin lipid film on the glass tube is dissolved in chloroform:methanol (1:1; v:v) and (III) stored in a glass vial with screw caps sealed with parafilm. (B) The glass tube containing the lipids in chloroform is connected to the rotary evaporator. (C) Equipment for a GUV formation of the home-made chamber. (D) Schematic workflow of GUV formation by the swelling method. The lipid mixture is applied to the cleaned glass slides and (I) dehydrated in a desiccator at 250 mbar for 30–60 min. The dried lipid film is then (II) rehydrated in a swelling buffer for 2–4 h, resulting in the formation of giant vesicles.
Annexin V staining of GUVs
Place 25 μL of GUVs (with a cut tip) in 25 μL of binding buffer (see Recipe 7) on a cover glass slide.
Note: As additional control, a GUV sample in buffer without Ca2+ can be prepared.
Place the cover glass slide under the microscope.
Let the GUVs settle for approximately 5 min before imaging in a white light channel and an annexin V channel.
Data analysis
Data acquisition
Images are acquired at a Leica TCS SP8 confocal laser scanning microscope (Leitz, Wetzlar, Germany) equipped with a 63×/1.20 water objective. For detailed imaging settings, see Table 1. All images are acquired at the same resolution, magnification, and orientation. This allows direct comparison of images and saves time when arranging figures.
Table 1. Confocal microscope settings
Channel Laser Detector Scan speed Excitation, nm Intensity, % Emission, nm Gain
Annexin V White light laser, intensity at 85%
Hybrid Detector
(HyD)
400 Hz 577 5 587–757 100
Bright light
White light laser, intensity at 85%
Photomultiplier
tube (PMT)
400 Hz trans channel - - 300
For analysis of the adherent cells
Export the raw image data in a format compatible with ImageJ (e.g., .tif) from the imaging system.
Open the ImageJ software and import the images of the blue fluorescence (DAPI) and the red fluorescence (Annexin V conjugated to Alexa Fluor 568).
Click on Image in the upper operation row of the interface and select in the color menu Merge Channels.
For C1 (red), the image showing the red fluorescence needs to be selected; for C3 (blue), the DAPI picture needs to be chosen. Press Ok to merge the two images.
Click on Image in the upper operation row of the interface and change the image type to RGB Color.
Save the images as a .tif or .jpg file by selecting the File menu in the upper operation row of the interface and press Save As. Representative images are shown in Figure 3.
Figure 3. Representative confocal images of proliferating C2C12 wild-type and CDC50A knockout cells. Cells are stained with Alexa Fluor 568 conjugated annexin V (red) for 10 min in ice-cold TBSS with and without calcium. DAPI staining is used for visualization of nuclei. In contrast to C2C12 wild-type cells, CDC50A-deficient cells stained positive for annexin V, indicating increased surface exposure of aminophospholipids. Images are representative of three independent experiments. Scale bar: 30 μm.
For analysis of GUVs
ImageJ is used to analyze the membrane fluorescence intensity of individual GUVs before and after annexin V treatment. Only unilamellar giant vesicles are used for analysis.
Open the images with ImageJ by importing the LIF-file.
Split the channels to have bright light and annexin channel separately and save as Tiff.
Tiff images can be assembled (see Figure 4).
Figure 4. Imaging the binding of fluorescent annexin V on giant unilamellar vesicles (GUVs). Giant unilamellar vesicles are prepared from different lipids and incubated without (-) and with (+) annexin V in the presence of Ca2+. Vesicles are observed in bright light and annexin channel. DOPC, PC (18:1/18:1) only; DOPE, PC (18:1/18:1)/PE (18:1/18:1), (9/1, mol/mol); DOPS, PC (18:1/18:1)/PS (18:1/18:1), (9/1, mol/mol). Data shown are from one experiment representative of two independent vesicle preparations. Scale bar: 10 μm.
Extract signal intensities with ImageJ software
Continue with the annexin channel.
A region of interest (ROI) is placed around the GUV (ROIouter) of interest to measure the annexin V fluorescence intensity of the membrane, by measuring the integrated density value per pixel.
The second ROI is placed within the GUV lumen (ROIinner) (see Figure 5A).
The inner ROI is subtracted from the outer ROI to have the membrane density value per pixel of the annexin V fluorescent intensity (ΔI):
ΔI = I(ROIouter) - I(ROIinner)
Repeat the procedure for each GUV and save the data as an Excel (.xls) file.
The annexin V fluorescence intensities are plotted in form of a bar diagram using the OriginPro (see Figure 5B and Table 2).
Figure 5. Fluorescence intensity analysis on giant unilamellar vesicles (GUVs). (A) A first region of interest (ROI) is placed around the GUV (ROIouter), and a second ROI is placed within the GUV lumen (ROIinner) of interest to measure the annexin V fluorescence intensity of the membrane, by measuring the integrated density value per pixel using the software ImageJ. (B) The mean annexin V fluorescence intensities for GUVs with the indicated lipid compositions are presented as averages, based on individual measurements of n ≤ 10 GUVs. Error bars indicate standard deviations. Data are from one experiment representative of two independent vesicle preparations.
Table 2. The mean annexin V fluorescence intensities (ΔI) for giant unilamellar vesicles (GUVs) with the indicated lipid compositions. Data are from one experiment representative of two independent vesicle preparations. Data are plotted in Figure 5.
Lipids of GUVs -Annexin V +Annexin V
ΔI, average (n = 10) s.d. ΔI, average (n = 13–15) s.d.
DOPC 0.001 0.001 1.430 1.814
DOPE 0.086 0.114 1.107 1.229
DOPS 0.001 0.002 16.732 8.934
Recipes
Buffers were prepared using double-distilled water (ddH2O), which was obtained using an in-house water distillation system. Alternatively, all buffers are prepared using ultrapure water with purification sensitivity of 18 MΩ·cm-1 at 25 °C.
Growth medium
Open a 500 mL flask of high-glucose DMEM medium
Add 100 mL of FBS (heat-inactivated)
Optional: add 5 mL of 100× penicillin-streptomycin solution
Prepare in sterile cabinet; store at 4 °C
TBSS buffer + Ca2+ (0.5 L)
136 mM NaCl (3.97 g)
2.6 mM KCl (96.9 mg)
1.8 mM CaCl2 (132.3 mg)
1 mM MgCl2·6H2O (101.6 mg)
0.36 mM NaH2PO4·2H2O (24.8 mg)
5.56 mM D-glucose (500.8 g)
5 mM HEPES (600 mg)
Adjust pH to 7.4 with 1 M NaOH. Complete volume to 0.5 L. Sterilize by filtering using a 0.22 μm filter. Store at 4 °C up to several months.
TBSS buffer - Ca2+ (0.5 L)
136 mM NaCl (3.97 g)
2.6 mM KCl (96.9 mg)
1 mM MgCl2·6H2O (101.6 mg)
0.36 mM NaH2PO4·2H2O (24.8 mg)
5.56 mM D-glucose (500.8 g)
5 mM HEPES (600 mg)
100 μM EGTA (19 mg)
Adjust pH to 7.4 with 1 M NaOH. Complete volume to 0.5 L. Sterilize by filtering using a 0.22 μm filter. Store at 4 °C up to several months.
DAPI stock solution (1 mg/mL)
Dissolve DAPI in ultrapure water to 1 mg/mL. Stock solution is stable for several months and repeated use, if stored protected from light at -20 °C.
Lipid stocks in chloroform
Lipids are ordered in chloroform at a concentration of 25 mg/mL and stored at -20 °C until further use. For longer storage, aliquot 10 mg of lipids in glass vials with screw caps, evaporate the chloroform, and store the dried lipid at -20 °C. Before using it, dissolve the 10 mg of lipid in 1 mL of chloroform:methanol (1:1; v:v).
Critical: Some lipids may have limited or very poor solubility in chloroform:methanol (1:1; v:v) and require a mixture of chloroform:methanol:water.
Swelling buffer (320 mM)
Dissolve 5.48 g of sucrose to a final volume of 50 mL in deionized water. The buffer is filter-sterilized over a 0.2 μm Acrodisc® syringe filter and stored at 5 °C.
Binding buffer (100 mL)
10 mM HEPES-KOH pH 7.4 (238.31 mg)
150 mM NaCl (876.6 mg)
5 mM KCl (37.275 mg)
1 mM MgCl2·6H2O (20.33 mg)
1 mM CaCl2 (11.098 mg)
Adjust pH to 7.4 with 1 M NaOH. Complete volume to 100 mL. Store at 4 °C up to several months.
Acknowledgments
We gratefully acknowledge Michelle Werner for technical assistance. This protocol was adapted from our previous work (Weingärtner et al., 2012; Grifell-Junyent et al., 2022; Herrera et al., 2022). The work was supported by the Lundbeckfonden (R221-2016-1005 to T.G.P.) and an instrument grant from the Deutsche Forschungsgemeinschaft (INST 213/886-1 FUGG to T.G.P.). HDU is a scholar of the Friedrich Ebert Foundation.
Competing interests
The authors declare that no competing interests exist.
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Effective Hand Rearing of Neonatal Mice for Developmental Studies
ML Marcus N. Leiwe
SF Satoshi Fujimoto
Takeshi Imai
Published: Vol 13, Iss 11, Jun 5, 2023
DOI: 10.21769/BioProtoc.4755 Views: 1040
Reviewed by: Pengpeng LiMatthew Grubb Anonymous reviewer(s)
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Abstract
Chronic manipulation in neonatal mice is a technical challenge, but it can achieve greater insights into how mice develop immediately after birth. However, these manipulations can often result in maternal rejection and consequently serious malnourishment and occasional death. Here, we describe a method to effectively hand rear mice to develop normally during the first post-natal week. In our experiments, we were able to negate the feeding deficiencies of anosmic mutant mice when compared to littermate controls. As a result, the delayed neuronal remodeling seen in maternally reared mutant mice was not seen in the hand-reared mutant mice. This methodology is user intensive but can be useful for a broad range of studies either requiring many interventions or one intervention that can result in maternal rejection or being outcompeted by healthy littermates.
Keywords: Neonatal Hand rearing Mice Development Feeding
Background
The common use of neonatal mice in developmental studies can result in maternal rejection (or even, in some circumstances, cannibalization) after experimental or genetic manipulations. In other situations, neonatal mice may be unable to suckle on their own; for example, anosmic mutant mice face severe difficulties in their suckling behavior (Brunet et al., 1996; Fujimoto, et al., 2023). These difficulties make it hard to determine whether altered development is due to the manipulation or simply due to a lack of maternal feeding.
In these cases, it is often necessary to hand rear the pups to prevent any difficulties. Successful hand rearing is dependent on more than simply delivering sufficient nutrients. For example, nesting conditions can have an influence on thermal stress and postnatal development (Gaskill et al., 2012; Mason et al., 2018). However, the majority of previous attempts have focused on nutrient delivery. To this end, a variety of techniques have been employed, such as force feeding with a syringe (Pleasants, 1959; Hoshiba, 2004) or directly adding nutrients to the stomach via a gavage (Messer et al., 1969; Hall, 1975). However, these methods were either only partially effective or technically difficult, which resulted in unreliable results or a low success rate. Additionally, varying the type of milk only produced worse results (Smart et al., 1984; Tonkiss et al., 1987).
Could a combination of improving the environmental conditions, feeding techniques, and neonatal care result in more effective and reliable growth of pups? There is a wide array of non-academic advice on how to rear mice (Creek Valley Critters, https://www.youtube.com/user/CreekValleyCritters), and we sought to harness this knowledge and apply it to biological research (Fujimoto et al., 2023). We discovered that the key features for successful growth were warm temperatures, regular feeding every two hours, and belly massages to avoid bloating. While this technique is manually very intensive, it produces pups that are healthy, and it does not require technically difficult procedures.
Materials and reagents
All the materials and reagents for hand rearing are shown in Figure 1.
Pipettes (Gilson, catalog numbers: P200 and P1000)
1.5 mL Eppendorf tubes (Bio-bik, catalog number: CF-0150)
Needles for injection (29 G) (BD, catalog number: 326666)
Paintbrush (Tokyu Hands, round No. 1)
Cotton buds
Tissue paper for nesting material
Cardboard box as a cage (W 16 cm × D 16 cm × H 6 cm)
Warm water (for cleaning pups)
Saline (Otsuka, catalog number: 3311401A7028)
70% ethanol
Powdered dog milk (PetAG, Ebsilac) (see Recipes)
Figure 1. Materials required for hand rearing pups
Equipment
All equipment items for hand rearing are shown in Figure 2.
Warm hood (W 60 cm × D 50 cm × H 52 cm) (As One, Pasaurina Glove Box with outlet, model: AS-600SC, https://www.axel-gl.com/en/asone/d/3-4041-03/)
Note: Most chambers should be adequate, provided they are able to maintain a warm and stable environment that can also be easily cleaned.
Heat mat (Natsume, model: KN-475-3-40)
Temperature monitor (As One, model: A-230-W)
Pet heater (Marukan, model: HD-40c)
Thermostat (Marukan, model: HD-1)
Dry bath incubator (Major science, model: MD-02N) or water bath (Sansyo, model: SWS-181D)
Electronic scale (A&D company, model: HT-120)
Pipettes (Gilson, catalog numbers: P200 and P1000)
Ultraviolet lamp
Figure 2. Schematic and photo of the hand rearing chamber
Procedure
Sterilize the hood 24 h prior to the experiment.
Note: Use the built-in UV light and 70% ethanol in order to prevent infections during the hand rearing period.
Prepare powdered dog milk in 1.5 mL tubes (see Recipes).
Prewarm the chamber to ~30 °C by turning on the heat pad and pet heater.
Note: The temperature probe should be placed in the cardboard box where the mice will be located.
Turn on the dry bath incubator or water bath and set it to 37 °C.
Note: Place the milk required for the first feeding session as well as saline (for intraperitoneal injections if the pups become dehydrated) and water (used as a saliva substitute; see step 8) into the dry bath incubator or water bath ~10 min before use.
Provide nesting material.
Note: Two to three sheets of tissue paper torn into strips should suffice. Autoclaving the nesting materials is not necessary for successful rearing, but you should follow the regulations of the specific mouse facility and/or laboratory.
Select up to three pups to hand rear.
Notes:
One person can take care of up to three pups at the same period. It is recommended to avoid using the runt of the litter as they tend to not feed so well.
Leave some pups behind with the dam to compare growth rates with those that are hand reared.
Prior to touching the hand-reared pups at any time, make sure your hands are clean. This is to prevent infection; usually we clean our hands with 70% ethanol. Skin-to-skin contact seems to be beneficial for the pups, as it helps to reduce heat loss without gloves; however, follow the regulations for mouse handling of the mouse facility and/or the laboratory.
It is advisable to hand rear more than one pup, in order to allow them to nest together and use their body warmth to warm each other.
Our experiments began from P1; however, it is possible to rear pups immediately from birth.
Weigh pups prior to starting the experiment.
Notes:
Use a scale that is at least sensitive to 0.01 g.
In our experiments, we weighed mice every 12 h to check if the weight gain was sufficient.
Clean the pup prior to feeding.
Steps 8–12 should be performed in the warm hood. Pups should not leave the hood except when performing weight checks.
Clean hands with 70% ethanol.
Notes:
i. Warm your hands with the pet heater to as close to the ambient temperature as possible.
ii. Neonatal mice are small and can lose body heat very quickly. Having warm hands prevents this.
Dip a cotton bud in water at 37 °C (heated in a 1.5 mL tube in the water bath).
Gently clean the genital region with the cotton bud.
Remove any fecal material.
Notes:
i. The fecal material should be a yellow-orange color; if the color of the feces changes to black, it indicates that there may be internal bleeding.
ii. Do not be surprised if the pup begins to urinate.
Use a new cotton bud to clean the face, whiskers, and rest of the body.
Note: Dried milk can occasionally get stuck between the fingers and toes of the pup and fuse them together. To prevent infections, it is best to remove this.
Using the paintbrush, feed the pup with the required volume of milk (100 μL/g body weight).
Prior to feeding, aliquot the volume required (100 μL/g body weight) into a new 1.5 mL tube and preheat in the dry bath incubator or the water bath so it reaches 37 °C before feeding.
Dip the paintbrush into the milk and carefully place the tip of the brush just inside the mouth of the pup (see Video 1).
Notes:
i. The first several sessions may take some time, as the pup begins to learn how to suckle from the paintbrush.
ii. If milk begins to flow out from the nostrils, stop feeding and clean the pup. It can either indicate that the pup is full or is drinking too quickly.
1) If this is the case, turn the pup upside down to aid the flow out through the nostril and remove as much milk as possible with a cotton bud.
2) If the pup still suckles, continue feeding slowly.
Continue feeding until the milk in the 1.5 mL tube from step 9a is finished or if 30 min of feeding time have elapsed. The stomach should now be white and clearly visible.
Video 1. Feeding procedure. See step 9 for details.
Using a new cotton bud soaked in warm water, clean the pup until milk and dry skin are removed, mimicking the licking behavior that dams exhibit for their pups.
Massage the stomach of the pups to prevent bloating (see Video 2).
Notes:
Repeated gentle downward movements can often help in this process.
Bloating can be seen by either the presence of a swollen abdomen or several dark spots (air pockets) within the stomach (normally white when filled with milk).
i. In case of bloating, attempt to remove the air pockets from the stomach by increased massages.
ii. If the abdomen appears to be bloated and firm, and massaging with the cotton bud does not aid the process, a stronger form of massage may be necessary. Place the pup on the heating pad with the ventral side in contact with the pad (see Video 3). Gently hold and push down on the abdomen and begin to massage the abdomen in a circular manner. In extreme cases, approximately 10 min of massaging may be necessary to remove the blockages in the digestive system. A successful massage should result in urination and excretion from the pup in question.
Video 2. Massage and excretion. See steps 9, 10, and 11 for details.
Video 3. Robust massage and excretion. See step 11b.ii.
Return the pup to the nest in the cardboard box.
Notes:
Once finished, return the pup to the nesting material.
The other pups can be maintained in the cardboard cage until they are ready to be fed. Allow the pups to nest together to warm each other.
Repeat steps 8–12 for the other pups. Steps 8–12 typically take 30 min per pup.
Clean the paintbrush by washing out the milk with water and then spraying with 70% ethanol and leave to dry.
Prepare the milk for the next feeding session.
Notes:
If the pups are easily able to finish the allocated milk, it may be necessary to increase the quantity of milk provided.
Conversely, if the pup is bloated it may be best to either reduce the quantity of milk or skip one feeding session to focus on the massages to remove the bloating.
Begin the feeding and cleaning process again two hours after starting (see Figure 3).
Continue the schedule every two hours until P6 or your desired time point. We only have experience with up to P6. Note that circadian rhythms are not apparent until ~P10. As manual feeding will be very laborious (1.5 h feeding for three pups and 0.5 h break, 12 cycles a day), it is almost impossible to do with only one person. Feeding should be performed in shifts with at least two people.
Figure 3. Schedule for hand rearing pups from P1 to P6
Data analysis
Measure the weight using an electronic scale every 12 h. Figure 4 shows one of our examples. Data analysis has been described in the original publication (Fujimoto et al., 2023).
Figure 4. Rescuing anosmic mutant mice from growth defects (Fujimoto et al., 2023). Olfactory sensory neuron-specific tetanus toxin light chain mice (OSN-TeNT, OMP-Cre;R26-CAG-LoxP-TeNT) were used. OSN-TeNT mice are anosmic, because neurotransmission from OSNs is blocked. A. Differences in development can clearly be seen when both pups are maternally reared. B. After using our hand-rearing protocol, no differences were visible between the two groups. C. Timeline depicting the weight changes during hand rearing. See Fujimoto et al. (2023) for more details. Scale bars: 10 mm.
Notes
Handling: initially, pups will be nervous as they are unused to being handled. During the first 24 h, provide as much exposure as possible. Additionally, the temperature of the experimenters’ hands is important. We suggest warming them up on the pet heater prior to handling but after cleaning.
Feeding: pups will take a while to learn how to suckle from the paintbrush; be patient. Additionally, different pups will prefer different feeding positions.
Bloating: feeding the pups vertically appears to increase the likelihood of bloating. It is recommended that you feed the pups while they are on their back or on their abdomen.
Dehydration: dehydration can be detected by the transparency of the skin. The greater the transparency, the greater the state of dehydration. We recommend delivering 10 μL of saline subcutaneously every two hours until the pup improves.
Weight gain: when comparing with maternally reared littermate controls, it is worth noting that the weight of the hand-reared pups is still lower (50%–90% at P6 in our examples). This may indicate that hand rearing cannot fully rescue the mice from growth defects. It should also be considered that the mother from whom the pups have been taken has now fewer pups to care for, and thus their growth may be higher than in a truly separate control case. Thus, the maternally reared littermates may not be appropriate controls. Hand-reared wild-type mice should be used as a control for hand-reared mutant mice.
Recipes
Powdered dog milk
Use 9 g of powdered milk per 30 mL of warm tap water. Do not use deionized water, as the ions are needed for correct development
Note: To help dissolve the powder, it is recommended to use a 50 mL centrifuge tube and then vortex the content until the powder is fully dissolved. We did not filter sterilize the milk, but you can if it is a requirement from the regulation of the mouse facility/laboratory.
Aliquot 1 mL of liquid into a 1.5 mL tube.
Note: Repeat until the centrifuge tube is empty.
Store aliquots at 4 °C for up to one week.
Note: If you need to store the milk for longer, store at -20 °C for up to 1–2 months. However, it is recommended to prepare fresh solutions when needed.
Acknowledgments
This work was supported by grants from the PRESTO program of the Japan Science and Technology Agency (JST) (T.I.), the JSPS KAKENHI (23680038, 15H05572, 15K14336, 16K14568, 16H06456, and 17H06261 to T.I., 15K14327 and 17K14944 to S.F., 17K14946 to M.N.L.), Sumitomo Foundation (T.I.), Nakajima Foundation (T.I.), and RIKEN CDB intramural grant (T.I.).
The procedures described here were based on the advice from the YouTube channel Creek Valley Critters (https://www.youtube.com/user/CreekValleyCritters) with some modifications by our hands. This YouTube channel contains additional useful information regarding hand rearing.
This protocol has been used in our research work (Fujimoto et al., 2023).
Competing interests
There are no conflicts of interest or competing interests.
Ethics
All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of the RIKEN Kobe Branch and Kyushu University. Both males and females were used for our experiments.
References
Brunet, L. J., Gold, G. H., and Ngai, J. (1996). General anosmia caused by a targeted disruption of the mouse olfactory cyclic nucleotide-gated cation channel. Neuron 17(4): 681–693.
Fujimoto, S., Leiwe, M. N., Aihara, S., Sakaguchi, R., Muroyama, Y., Kobayakawa, R., Kobayakawa, K., Saito, T. and Imai, T. (2023). Activity-dependent local protection and lateral inhibition control synaptic competition in developing mitral cells in mice. Dev Cell doi: https://doi.org/10.1016/j.devcel.2023.05.004
Gaskill, B. N., Gordon, C. J., Pajor, E. A., Lucas, J. R., Davis, J. K. and Garner, J. P. (2012). Heat or insulation: behavioral titration of mouse preference for warmth or access to a nest. PLoS One 7(3): e32799.
Hall, W. G. (1975). Weaning and growth of artificially reared rats. Science 190(4221): 1313–1315.
Hoshiba, J. (2004). Method for hand-feeding mouse pups with nursing bottles. Contemp. Top Lab Anim. Sci. 43(3): 50–53.
Mason, B., Rollins, L. G., Asumadu, E., Cange, C., Walton, N. and Donaldson, S. T. (2018). Nesting environment provides sex-specific neuroprotection in a rat model of neonatal hypoxic-ischemic injury. Front. Behav. Neurosci. 12: 221.
Messer, M., Thoman, E. B., Galofre, A., Dallman, T. and Dallman, P. R. (1969). Artificial feeding of infant rats by continuous gastric infusion.J. Nutr. 98(4): 404–410.
Pleasants, J. R. (1959). Rearing germfree cesarean-born rats, mice, and rabbits through weaning. Ann. N. Y. Acad. Sci. 78: 116–126.
Smart, J. L., Stephens, D. N., Tonkiss, J., Auestad, N. S. and Edmond, J. (1984). Growth and development of rats artificially reared on different milk-substitutes. Br. J. Nutr. 52(2): 227–237.
Tonkiss, J., Smart, J. L. and Massey, R. F. (1987). Growth and development of rats artificially reared on rats' milk or rats' milk/milk-substitute combinations. Br. J. Nutr. 57(1): 3–11.
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A Simple Sonication Method to Isolate the Chloroplast Lumen in Arabidopsis thaliana
JH Jingfang Hao
AM Alizée Malnoë
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4756 Views: 599
Reviewed by: Aswad KhadilkarSam-Geun Kong Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Plants Jul 2022
Abstract
The chloroplast lumen contains at least 80 proteins whose function and regulation are not yet fully understood. Isolating the chloroplast lumen enables the characterization of the lumenal proteins. The lumen can be isolated in several ways through thylakoid disruption using a Yeda press or sonication, or through thylakoid solubilization using a detergent. Here, we present a simple procedure to isolate thylakoid lumen by sonication using leaves of the plant Arabidopsis thaliana. The step-by-step procedure is as follows: thylakoids are isolated from chloroplasts, loosely associated thylakoid surface proteins from the stroma are removed, and the lumen fraction is collected in the supernatant following sonication and centrifugation. Compared to other procedures, this method is easy to implement and saves time, plant material, and cost. Lumenal proteins are obtained in high quantity and purity; however, some stromal membrane–associated proteins are released to the lumen fraction, so this method could be further adapted if needed by decreasing sonication power and/or time.
Keywords: Thylakoid lumen Thylakoid membrane Sonication Protein Arabidopsis thaliana
Background
The chloroplast is the organelle that conducts photosynthesis in plants and algae. A major compartment of the chloroplast is the thylakoid lumen, which is enclosed by the thylakoid membrane. The Arabidopsis lumen proteome consists of at least 80 proteins based on mass spectrometry analyses (Peltier et al., 2002; Schubert et al., 2002) and up to 127 proteins according to lumenal targeting peptide prediction (Almagro Armenteros et al., 2019). All lumenal proteins characterized so far are nuclear-encoded and post-translationally transported into chloroplasts [see for reviews on lumenal proteins targeting (Albiniak et al., 2012) and function (Kieselbach and Schröder, 2003; Järvi et al., 2013)]. Lumenal proteins support and modulate photosynthetic activity directly or indirectly (examples of proteins are given in parenthesis), with known function in electron transport (plastocyanin PC, photosystem I subunit PsaN, and photosystem II subunit PsbO), in protein processing (C-terminal processing protease ctpA), assembly (immunophilins), and degradation (Deg proteases), in redox regulation (membrane protein with lumen thioredoxin domain HCF164) and in photoprotection (violaxanthin de-epoxidase and lipocalin in the plastid LCNP). However, the function of several lumenal proteins remains to be elucidated (some of which are Psb-like and pentapeptide repeat-containing proteins) and their regulation is not fully understood. The activity, stability, and distribution in the lumen compartment as soluble or membrane-associated proteins, or protein–protein interactions can be regulated by post-translational modifications such as phosphorylation or N-terminal acetylation (Gollan et al., 2021), or through redox modification or disulfide bond modulation, e.g., by lumen thiol oxidoreductase 1 of the lumenal domain of the kinase STN7 (Wu et al., 2021; for reviews, see Buchanan and Luan, 2005; Kang and Wang, 2016).
The major challenge when performing studies on the lumen proteome is the balance between purity and the quantity of protein needed for further analysis. Indeed, lumenal proteins are lowly abundant compared to thylakoid membrane protein light harvesting complex LHCII and stromal Rubisco [which represent more than 50% of total chloroplast protein content (Hall et al., 2011)]. Fractionation of the chloroplast and isolation of lumenal proteins thus enable their detection and accurate quantification by working within the dynamic range of detection, for example in immunoblot assays (the dynamic range is the lowest to highest concentration of a given protein that can be reliably detected). Then, the accumulation of a protein of interest can be investigated in different growth conditions or stress treatments and in mutants; the lumen proteome has been analyzed from 6–8-week-old plants grown under standard growth conditions (i.e., 120 μmol photons m-2·s-1, 21 °C, 8:16 h light/dark) (Peltier et al., 2002; Schubert et al., 2002) and also comparing the end of the dark vs. light period (Granlund et al., 2009) or after cold acclimation (Goulas et al., 2006). In addition, localization and distribution of the proteins, as well as assessment of protein complexes and interaction, can be inferred. Recently, Gollan et al. (2021) refined further localization studies to distinguish free lumenal proteins from membrane-associated ones, using Yeda press to isolate soluble lumenal proteins and subsequent washes with urea and salt to release inner membrane-associated proteins.
To isolate lumen proteins from plants, first the chloroplasts are obtained by homogenization of leaves followed by centrifugation; then, chloroplasts are lysed by osmotic shock, and thylakoid membranes are collected by centrifugation and washed to remove stromal proteins and peripheral membrane proteins. Finally, thylakoid membranes are ruptured to release lumenal proteins with a Yeda press (Kieselbach et al., 1998) or a sonicator (Peltier et al., 2000; Levesque-Tremblay et al., 2009); alternatively, thylakoid membranes are solubilized to release lumenal proteins using a detergent such as Triton X-114 followed by phase partitioning at 30–37 °C (Bricker et al., 2001), 0.04% Triton X-100 (McKinnon et al., 2020), or 0.05% n-Dodecyl β-D-maltoside (Chang et al., 2021).
Here, we report a simple procedure by which a lumenal fraction can be isolated in pure form from the thylakoids in Arabidopsis by sonication, which we adapted from previously described methods (Peltier et al., 2000; Levesque-Tremblay et al., 2009). Lumen isolation by sonication is of interest for several reasons: 1) the Yeda press is no longer commercially available (Hall et al., 2011), so unless already existing in the laboratory, access to one is limiting; 2) operating a sonicator is easier and faster than using the Yeda press for more than two samples (due to faster washing time of instruments parts); and 3) use of detergent can be costly and can disrupt native interactions. Also, the yield with sonication is comparable to using the Yeda press (15–30 μg of lumenal proteins per gram of leaf) and a decreased isolation time (from six to three hours for two samples) is valuable to limit proteolysis and preserve more native states of proteins and complexes during the lumen isolation. All methods present the disadvantage of stromal membrane–associated protein contaminants in the lumenal fraction that are not removed during the washes of thylakoid membranes (Bricker et al., 2001) (for example, see Figure 2, ATPb); the presence of these contaminants could be decreased by using a shorter sonication time and/or decreased power (Peltier et al., 2000). Overall, the sonication method is easy to implement, saves time, plant material, and cost, and is suitable for most studies—unless a large quantity of proteins is required, in which case the Yeda press method should be favored.
Materials and reagents
1.5 mL microcentrifuge tube (Sigma-Aldrich, catalog number: HS4323)
15 mL centrifuge tube (Thermo Fisher Scientific, catalog number: 339650)
Amicon ultra-0.5 centrifugal filter unit (EMD Millipore, catalog number: UFC500324)
10.4 mL polycarbonate bottle with cap assembly (Beckman Coulter, catalog number: 355603)
50 mL open-top thick-wall polycarbonate tube (Beckman Coulter, catalog number: 363647)
10 μL pipette tip (Thermo Fisher Scientific, catalog number: 3521-HR)
200 μL pipette tip (Thermo Fisher Scientific, catalog number: 3551-HR)
1,000 μL pipette tip (Thermo Fisher Scientific, catalog number: 3101-HR)
Disposable glass Pasteur pipettes 230 mm (VWR, catalog number: 612-1702)
1,000 mL plastic bag (e.g., Tingstad, catalog number: 398301-1)
Glass funnel (e.g., Sagitta, catalog number: 87807)
250 mL flask (e.g., Sagitta, catalog number: 86425)
25 mL beaker (e.g., VWR, catalog number: 213-0192)
Cuvette for chlorophyll quantification [e.g., Hellma, catalog number: HL104-002-10-40 (quartz, preferred) or Sarstedt, catalog number: 67.742 (plastic; ensure to measure right away so acetone does not degrade plastic and affect spectrophotometer reading)]
Paintbrush 6 mm (Ahlsell, catalog number: 384364)
Miracloth 22–25 μm pore size (Calbiochem, catalog number: 475855)
Arabidopsis thaliana plants (wild type and soq1-1 mutant, ecotype: Columbia-0)
Milli-Q water
2-Mercaptoethanol (Thermo Fisher Scientific, catalog number: 21985023)
Glycerol (Sigma-Aldrich, catalog number: G5516)
Bromophenol blue (Sigma-Aldrich, catalog number: B0126)
Ethanol (Sigma-Aldrich, catalog number: EX0290)
Acetic acid (Sigma-Aldrich, catalog number: 695092)
Coomassie blue R-250 (Sigma-Aldrich, catalog number: 1.12553)
Tris(hydroxymethyl)aminomethane (Tris base) (Sigma-Aldrich, catalog number: 648310-M)
Glycine (Sigma-Aldrich, catalog number: G8898)
Sodium dodecyl sulfate (SDS) (Sigma-Aldrich, catalog number: L3771)
Tween 20 (Sigma-Aldrich, catalog number: P9416)
Non-fat dried milk (e.g., Semper)
Tricine (Sigma-Aldrich, catalog number: T0377)
D-sorbitol (Sigma-Aldrich, catalog number: S1876)
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: E9884)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: 529552)
Sodium L-ascorbate (Sigma-Aldrich, catalog number: A4034)
L-cysteine (Sigma-Aldrich, catalog number: C7352)
Sodium fluoride (Sigma-Aldrich, catalog number: 215309)
MgCl2 (Sigma-Aldrich, catalog number: M2670)
Concentrated HCl (37%) (Sigma-Aldrich, catalog number: 320331)
Disodium hydrogen phosphate (Na2HPO4) (Sigma-Aldrich, catalog number: 567550)
Sodium dihydrogen phosphate (NaH2PO4) (Sigma-Aldrich, catalog number: 1.06370)
Benzamidine (Sigma-Aldrich, catalog number: 12072)
ϵ-Aminocaproic acid (Sigma-Aldrich, catalog number: A2504)
Phenylmethanesulfonyl fluoride solution (PMSF) (Sigma-Aldrich, catalog number: 93482)
cOmpleteTM, EDTA-free protease inhibitor cocktail (Roche, catalog number: 4693132001)
Quick StartTM Bradford Protein Assay kit (Bio-Rad, catalog number: 5000201)
PageRuler prestained protein ladder (Thermo Fisher Scientific, catalog number: 26616)
Immobilon-P PVDF Membrane (Millipore Sigma, catalog number: IPVH00005)
ECL bright kit for immunodetection (Agrisera, catalog number: AS16 ECL-N)
Anti-PsaD, 1:1,000 dilution (Agrisera, catalog number: AS09 461)
Anti-Lhcb4, 1:7,500 dilution (Agrisera, catalog number: AS04 045)
Anti-RbcL, 1:7,500 dilution (Agrisera, catalog number: AS03 037)
Anti-PC, 1:2,000 dilution (Agrisera, catalog number: AS06 141)
Anti-ATPb, 1:5,000 dilution (Agrisera, catalog number: AS05 085)
Horseradish peroxidase (HRP)-conjugated anti-rabbit secondary antibody, 1:10,000 dilution (Sigma-Aldrich, catalog number: A6154)
1 M Tris-HCl stock solution (6.8 and 7.5) (see Recipes)
100 mM Sodium phosphate stock solutions (pH 7.8) (see Recipes)
Extraction buffer (see Recipes)
Resuspension buffer (see Recipes)
Lysis buffer (see Recipes)
Washing buffer (see Recipes)
4× sample loading buffer for SDS-PAGE (see Recipes)
Running buffer for SDS-PAGE (see Recipes)
Transfer buffer for immunodetection (see Recipes)
TBST buffer (see Recipes)
Blocking buffer (see Recipes)
Coomassie blue stain (see Recipes)
Coomassie blue destain (see Recipes)
Equipment
10 μL pipette (Thermo Fisher Scientific, catalog number: 4641040N)
20 μL pipette (Thermo Fisher Scientific, catalog number: 4641060N)
200 μL pipette (Thermo Fisher Scientific, catalog number: 4641080N)
1,000 μL pipette (Thermo Fisher Scientific, catalog number: 4641100N)
Plant growth cabinet (CLF plant Climatics, model: E-41L2)
Custom-designed LED panel, built by JBeamBio with cool white LEDs BXRA-56C1100-B-00 (Farnell)
Laboratory balance (e.g., Fisher Scientific, catalog number: 14-557-421)
Blender (e.g., Coline, 300 mL mixer cup with 2-bladed knife)
High-speed centrifuge (Beckman Coulter, model: Avanti J-20XP) with JA-25.50 fixed angle aluminum rotor (Beckman Coulter, catalog number: 363055)
Ultracentrifuge (Beckman Coulter, model: LE-70) with type 70.1 Ti fixed-angle titanium rotor (Beckman Coulter, catalog number: 342184)
Benchtop centrifuge (Beckman Coulter, catalog number: B06322)
UV-visible spectrophotometer (Hitachi, catalog number: U-5100)
Sonicator (Sonics, model: VCX130) with 2 mm microtip (Sonics, catalog number: 630-0417)
Dry block heater (MRC, catalog number: DBSC-001)
Immunodetection imaging system (Azure, model: c600)
Procedure
Plant material and growth conditions
Wild-type Arabidopsis thaliana and soq1-1 mutant were grown in a growth chamber with 120 μmol photons m-2·s-1 light intensity and 60% humidity at 20 °C during the day for 8 h and 18 °C during the night for 16 h. For cold and high light conditions (Cold HL), plants are illuminated in the cold room (4 °C) for 6 h at 1,600 μmol photons m-2·s-1 light intensity.
Thylakoid lumen isolation by sonication
Prepare extraction buffer [see Recipe 1 (Fristedt et al., 2009)], resuspension buffer (see Recipe 2), lysis buffer (see Recipe 3), and washing buffer (see Recipe 4). Keep at 4 °C.
Harvest the Arabidopsis rosettes from intended conditions (e.g., growth light and cold and high light conditions) into plastic bags. After removing the air, seal the bags and put them in the dark in the cold room. We usually use four rosettes from plants grown under short-day (8:16 h day/night) conditions for seven weeks (~10 g of fresh weight).
Blend the plant samples in 90 mL of extraction buffer three times for 5 s using a blender.
Filtrate the homogenate through a glass funnel with four layers of Miracloth to a 250 mL flask on ice.
Transfer the filtrated homogenate to 50 mL open-top thick-wall polycarbonate tubes (two tubes) and centrifuge at 1,000× g for 5 min.
Resuspend the chloroplast pellet in 35–40 mL of resuspension buffer using a soft paintbrush and centrifuge at 1,000× g for 5 min.
Note: We initially use a small volume (~3 mL) to resuspend the chloroplast using a paintbrush, flash freeze one or two aliquots (100 μL) in liquid nitrogen, and store them at -70 °C until immunoblot analysis. Then, we add resuspension buffer to 35–40 mL.
Resuspend the chloroplast in lysis buffer at a final chlorophyll concentration of 0.2 mg/mL and centrifuge at 6,000× g for 5 min.
Note: We usually keep the chloroplast in lysis buffer at 4 °C for 10 min to break the chloroplast envelopes by osmotic shock before centrifugation.
Concentrate approximately 2 mL of supernatant using Amicon ultra-0.5 centrifugal filters. Flash freeze the stromal aliquots (50 μL) in liquid nitrogen and store them at -70 °C until immunoblot analysis.
Wash the thylakoid pellet with 35–40 mL of washing buffer and centrifuge at 6,000× g for 5 min.
Note: We usually use a small volume (~3 mL) to resuspend the pellet using a paintbrush and then add washing buffer to 35–40 mL.
Resuspend the thylakoids in washing buffer, usually ~4 mL, so that the chlorophyll concentration is approximately 0.5 mg/mL. The method to measure the chlorophyll concentration is from Porra et al. (1989). Flash freeze the thylakoid aliquots (100 μL) at -70 °C until immunoblot analysis.
Transfer the thylakoid mixture to a 25 mL pre-cooled beaker and sonicate 10 times for 30 s ON (power: 130 watt) and 10 s OFF at 4 °C.
Note: We usually put half of the beaker into ice to keep the samples cold during the sonication operation.
Transfer the samples from the beaker to a 10.4 mL polycarbonate bottle, balance the samples with the washing buffer carefully, and ultracentrifuge at 200,000× g for 2 h at 4 °C to separate the thylakoid membranes in the pellet and the soluble lumenal proteins in the supernatant.
Transfer the supernatant (~3 mL) carefully to a new 15 mL centrifuge tube using a disposable glass Pasteur pipette and concentrate the soluble lumen sample (final volume is ~150 μL) at 14,000× g for 10 min at 4 °C using Amicon ultra-0.5 centrifugal filters. Flash freeze the lumen aliquots (25 μL) and store them at -70 °C until immunoblot analysis.
Resuspend the thylakoid membrane after isolation of lumen proteins in washing buffer (~4 mL) at a chlorophyll concentration of 0.5 mg/mL. Flash freeze the thylakoid membranes aliquots (100 μL) and store them at -70 °C until immunoblot analysis.
Sample preparation for SDS-PAGE
Take out samples (thylakoids, thylakoid membrane after isolation of lumen proteins, and lumen fraction) from -70 °C on ice and thaw rapidly at 37 °C on a heat block (less than a minute).
Wash the thylakoid and thylakoid membrane samples with 1 mL of 120 mM Tris-HCl (pH 6.8) and centrifuge at 20,000× g for 5 min at 4 °C.
Resuspend the pellets gently by adding 200 μL of 120 mM Tris-HCl (pH 6.8).
Measure the samples concentration using the Bradford assay (Kruger, 1994)
SDS-PAGE and immunoblot
Prepare the polyacrylamide gels for SDS-PAGE.
Note: The optimal concentration of separating gel depends on the protein of interest. We usually use a 15% separating gel for lumenal plastocyanin protein (PC; PC1 runs closer to 14 kDa while PC2 runs closer to 19 kDa) and 12% separating gel for thylakoid membrane light-harvesting complex b4 protein (Lhcb4, 29 kDa), core subunit A of photosystem I (PsaA, 55–60 kDa), stromal ribulose bisphosphate carboxylase oxygenase large subunit (RbcL, 53 kDa), beta subunit of ATP synthase (ATPb, 54 kDa), and suppressor of quenching 1 (SOQ1, 108 kDa).
Load the samples (5 μg of protein and 5 µL PageRuler prestained protein ladder) and run the polyacrylamide gels.
Note: We run the gels at a lower voltage (80 V) for 20 min and then increase to a higher voltage (120 V) until the dye front has reached the bottom of the gel.
After running the gel, transfer the proteins to a PVDF membrane in transfer buffer (see Recipe 7) by wet transfer at 200 mA for 90 min at 4 °C.
After transferring, incubate the PVDF membrane with blocking buffer (see Recipe 9) for 1 h at room temperature (20–25 °C).
Incubate the PVDF membrane with the primary antibody diluted with blocking buffer for 1 h at room temperature (20–25 °C).
Note: Here, we used anti-PsaD, anti-Lhcb4, anti-RbcL, anti-PC, rabbit-specific antibodies against a C-terminal peptide of SOQ1 (TVTPRAPDAGGLQLQGTR) (1:200 dilution, produced and purified by peptide affinity by ThermoFisher), and anti-ATPb.
Wash the PVDF membrane with TBST buffer (see Recipe 8) (10 min × three times)
Note: We used the horseradish peroxidase (HRP)-conjugated anti-rabbit secondary antibody.
Incubate the PVDF membrane with the secondary antibody diluted with blocking buffer for 1 h at room temperature (20–25 °C).
Wash the PVDF membrane with TBST buffer (10 min × three times).
Incubate the PVDF membrane with the ECL bright kit for immunodetection.
Data analysis
We used sonication to break the thylakoids and collected thylakoid lumen by ultracentrifugation. The purity of the different fractions was assessed by immunoblot analysis using various antibodies against stromal (RbcL), thylakoid membrane (PsaA, Lhcb4), and thylakoid lumen (PC) proteins (Figure 1). The results in Figure 1 confirmed the purity of thylakoids, thylakoid membrane after isolation of lumen proteins (named membrane), and lumen. There are some lumen contaminants in the stroma fractions, possibly due to some thylakoids being broken when we lysed chloroplasts by osmotic shock. Of note, the lumen fraction is not completely pure. We observed a small amount of the stromal subunit of ATP synthase (ATPb) in the lumen, suggesting that there are some thylakoid membrane contaminants in the isolated lumen fraction (Figure 2).
Figure 1. Immunoblot analysis of marker proteins in chloroplast sub-fractions. The protein samples were prepared from 7-week-old Arabidopsis plants (Col-0) under growth light (GL, 120 μmol photons m-2·s-1) and 6 h cold and high light (Cold HL, 4 °C, 1,600 μmol photons m-2·s-1) conditions. Samples were loaded at the same quantity of total protein (5 μg). Four chloroplast proteins (RbcL, PsaA, Lhcb4, and PC) are shown. From Yu et al. (2022).
By applying this method, we can investigate the localization and accumulation of chloroplast proteins under different conditions (e.g., Cold HL) (Yu et al., 2022). Using either the Yeda press or sonication methods, we found that SOQ1, a thylakoid membrane-anchored protein involved in negative regulation of photoprotection qH (Brooks et al., 2013; Malnoë et al., 2018), accumulated as full length and also as three distinct truncated lumenal forms [comprising three domains thioredoxin-like (T) NHL (N) C-terminal domain (C) named TNC, two domains NC, or the C-terminal domain only (CTD)] in growth light and Cold HL conditions (Figure 2). Because of the large volumes of buffers used during the preparation with the Yeda press, no protease inhibitors were added except EDTA for metalloproteinases inhibition; together with the long isolation time, a concern was that the truncated SOQ1 forms observed in the lumen fraction were an artifact of the preparation. In comparison, the small volumes of buffers used in the sonication method allow to add inhibitors of different proteases (e.g., Roche cOmpleteTM protease inhibitor cocktail tablet, one tablet for 50 mL buffer). In the presence of protease inhibitors and a shorter isolation time, we confirmed the existence of SOQ1 truncated forms in the lumen fraction. The full-length SOQ1 present in the lumen fraction is likely due to contamination from the thylakoid membrane. These results were published in Yu et al. (2022).
Figure 2. The accumulation of SOQ1 in chloroplast sub-fractions. The protein samples were prepared from 7-week-old wild type (Col-0) and soq1 mutant under growth light (GL, 120 μmol photons m-2·s-1) and 6 h cold and high light (Cold HL, 4 °C, 1,600 μmol photons m-2·s-1) conditions. Samples were loaded at the same quantity of total protein (5 μg). Four chloroplast proteins (SOQ1, Lhcb4, PC, and ATPb) are shown. Three distinct truncated forms (TNC, NC, and CTD) of SOQ1 are accumulated in the lumen in GL and Cold HL conditions. TNC: thioredoxin-like, NHL and C-terminal domains (calculated molecular weight is 73 kDa); NC: NHL and C-terminal domains (calculated molecular weight is 53 kDa); CTD: C-terminal domain (calculated molecular weight is 17 kDa). From Yu et al. (2022).
Notes
All steps for thylakoid lumen isolation are performed in the cold room (4 °C). The centrifuge equipment with the rotor is pre-cooled to 4 °C. The paintbrushes should be washed after each isolation step. All buffers for thylakoid lumen isolation are prepared one day before and stored in the cold room. BSA, sodium ascorbate, and L-cysteine should be added in the extraction buffer right before use [see also notes from Hall et al. (2011)]. The protease inhibitors can be prepared with the stock concentration of 100 mM benzamidine (stored at -20 °C), 500 mM ϵ-aminocaproic acid, and 100 mM PMSF stored at 4 °C for several months [see also notes from Bouchnak et al. (2018)]. The protease inhibitors with final concentration of 1 mM benzamidine, 5 mM ϵ-aminocaproic acid, and 0.2 mM PMSF and/or 1× Roche cOmpleteTM protease inhibitor cocktail tablet must be added in all buffers right before use.
Recipes
(*) Added right before use
Extraction buffer
50 mM tricine-NaOH (pH 7.8)
330 mM sorbitol
1 mM EDTA
10 mM KCl
*0.15% (w/v) bovine serum albumin (traps fatty acids)
*4 mM sodium ascorbate (limits protein oxidation)
*7 mM L-cysteine (limits protein oxidation)
*Protease inhibitors
Resuspension buffer
50 mM sodium phosphate (pH 7.8)
330 mM sorbitol
10 mM sodium fluoride (NaF) (inhibits phosphatase)
*Protease inhibitors
Lysis buffer
10 mM sodium phosphate (pH 7.8)
5 mM MgCl2
10 mM NaF
*Protease inhibitors
Washing buffer
50 mM sodium phosphate (pH 7.8)
100 mM sorbitol
10 mM NaF
5 mM MgCl2
*Protease inhibitors
4× sample loading buffer for SDS-PAGE
8% (w/v) SDS
20% (w/v) 2-mercaptoethanol
40% (v/v) glycerol
0.008% (w/v) bromophenol blue
25 mM Tris-HCl (pH 6.8)
Running buffer for SDS-PAGE
25 mM Tris base
192 mM glycine
0.1% (w/v) SDS
Transfer buffer for immunodetection
25 mM Tris base
192 mM glycine
20% (v/v) ethanol
0.0375% (w/v) SDS
Tris-buffered saline with 0.1% Tween 20 (TBST) buffer
20 mM Tris-HCl (pH 7.5)
150 mM NaCl
0.1% (v/v) Tween 20
Blocking buffer
5% (w/v) non-fat dried milk in TBST
Coomassie blue stain
0.1% (w/v) Coomassie blue R-250
40% (v/v) ethanol
10% (v/v) acetic acid
50% (v/v) Milli Q water
Coomassie blue destain
30% (v/v) ethanol
10% (v/v) acetic acid
60% (v/v) Milli Q water
1 M Tris-HCl stock solution (6.8 and 7.5, 500 mL)
Mix 60.57 g of Tris base with 400 mL of Milli Q water
Adjust the pH to 6.8 or 7.5 by adding concentrated HCl
Add Milli Q water until final volume is 500 mL
Note: The stock solution is used to prepare 25 mM and 120 mM Tris-HCl (pH 6.8) or 20 mM Tris-HCl (pH 7.5)
100 mM Sodium phosphate stock solution (pH 7.8)
10.4 mM NaH2PO4
89.6 mM Na2HPO4
Note: The stock solution is used to prepare 10 mM and 50 mM sodium phosphate (pH 7.8).
Acknowledgments
This protocol was adapted from Peltier et al. (2000) and Levesque-Tremblay et al. (2009). We thank W.P. Schroder for help with lumen preparation using the Yeda press method and critical reading of this protocol. This work was supported by a starting grant to A.M. from the Swedish Research Council Vetenskapsrådet (2018-04150), European Commission Marie Skłodowska-Curie Actions Individual Fellowship Reintegration Panel (845687) to A.M., by a consortium grant from the Swedish Foundation for Strategic Research (ARC19-0051) and by grants to UPSC from the Knut and Alice Wallenberg Foundation (2016.0341 and 2016.0352), and the Swedish Governmental Agency for Innovation Systems (2016-00504).
Competing interests
The authors declare that there is no conflict of interest.
Ethics considerations
The material used in this protocol is Arabidopsis thaliana.
References
Albiniak, A. M., Baglieri, J. and Robinson, C. (2012). Targeting of lumenal proteins across the thylakoid membrane. J. Exp. Bot 63(4): 1689–1698.
Almagro Armenteros, J. J., Salvatore, M., Emanuelsson, O., Winther, O., von Heijne, G., Elofsson, A. and Nielsen, H. (2019). Detecting sequence signals in targeting peptides using deep learning. Life Sci. Alliance 2(5): e201900429.
Bouchnak, I., Moyet, L., Salvi, D., Kuntz, M. and Rolland, N. (2018). Preparation of Chloroplast Sub-compartments from Arabidopsis for the Analysis of Protein Localization by Immunoblotting or Proteomics. J. Vis. Exp. 140: e58581
Bricker, T. M., Prevost, M., Vu, V., Laborde, S., Womack, J. and Frankel, L. K. (2001). Isolation of lumenal proteins from spinach thylakoid membranes by Triton X-114 phase partitioning. Biochim. Biophys. Acta-Bioenerg. 1503(3): 350–356.
Brooks, M. D., Sylak-Glassman, E. J., Fleming, G. R. and Niyogi, K. K. (2013). A thioredoxin-like/β-propeller protein maintains the efficiency of light harvesting in Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 110: 2733–2740.
Buchanan, B. B. and Luan, S. (2005). Redox regulation in the chloroplast thylakoid lumen: a new frontier in photosynthesis research. J. Exp. Bot. 56: 1439–1447.
Chang, W., Li, C., Cui, Z., Li, W., Song, H., Chang, H., Fu, W., Wang, C., Huang, T., Luo, Y., et al. (2021). Diverged Early From CtpB and CtpC, CtpA Has Evolved to Process D1 Precursor in Oxygenic Photosynthetic Organisms. Front. Plant Sci. 12: e676036.
Fristedt, R., Willig, A., Granath, P., Crèvecoeur, M., Rochaix, J. D. and Vener, A. V. (2009). Phosphorylation of Photosystem II Controls Functional Macroscopic Folding of Photosynthetic Membranes in Arabidopsis. Plant Cell 21(12): 3950–3964.
Gollan, P. J., Trotta, A., Bajwa, A. A., Mancini, I. and Aro, E. M. (2021). Characterization of the Free and Membrane-Associated Fractions of the Thylakoid Lumen Proteome in Arabidopsis thaliana. Int. J. Mol. Sci 22(15): 8126.
Goulas, E., Schubert, M., Kieselbach, T., Kleczkowski, L. A., Gardeström, P., Schröder, W. and Hurry, V. (2006). The chloroplast lumen and stromal proteomes of Arabidopsis thaliana show differential sensitivity to short- and long-term exposure to low temperature. Plant J. 47(5): 720–734.
Granlund, I., Hall, M., Kieselbach, T. and Schröder, W. P. (2009). Light Induced Changes in Protein Expression and Uniform Regulation of Transcription in the Thylakoid Lumen of Arabidopsis thaliana. PLoS One 4(5): e5649.
Hall, M., Mishra, Y. and Schröder, W. P. (2011). Preparation of stroma, thylakoid membrane, and lumen fractions from Arabidopsis thaliana chloroplasts for proteomic analysis. Methods Mol. Biol. 775: 207–222.
Järvi, S., Gollan, P. J. and Aro, E. M. (2013). Understanding the roles of the thylakoid lumen in photosynthesis regulation. Front. Plant Sci. 4: e00434.
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Malnoë, A., Schultink, A., Shahrasbi, S., Rumeau, D., Havaux, M. and Niyogi, K. K. (2018). The Plastid Lipocalin LCNP Is Required for Sustained Photoprotective Energy Dissipation in Arabidopsis. Plant Cell 30(1): 196–208.
McKinnon, L. J., Fukushima, J., Endow, J. K., Inoue, K. and Theg, S. M. (2020). Membrane Chaperoning of a Thylakoid Protease Whose Structural Stability Is Modified by the Protonmotive Force. Plant Cell 32(5): 1589–1609.
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Plant Science > Plant physiology > Abiotic stress
Biochemistry > Protein > Isolation and purification
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4,757 | https://bio-protocol.org/en/bpdetail?id=4757&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Preparation of Human Kidney Progenitor Cultures and Their Differentiation into Podocytes
MM Maria Elena Melica
MA Maria Lucia Angelotti
GA Giulia Antonelli
AP Anna J. Peired
CC Carolina Conte
LC Letizia De Chiara
BM Benedetta Mazzinghi
EL Elena Lazzeri
LL Laura Lasagni
PR Paola Romagnani
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4757 Views: 890
Reviewed by: Rajesh RanjanFereshteh Azedi Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Translational Medicine Aug 2022
Abstract
Kidney diseases are a global health concern. Modeling of kidney disease for translational research is often challenging because of species specificities or the postmitotic status of kidney epithelial cells that make primary cultures, for example podocytes. Here, we report a protocol for preparing primary cultures of podocytes based on the isolation and in vitro propagation of immature kidney progenitor cells subsequently differentiated into mature podocytes. This protocol can be useful for studying physiology and pathophysiology of human kidney progenitors and to obtain differentiated podocytes for modeling podocytopathies and other kidney disorders involving podocytes.
Graphical overview
Keywords: Kidney progenitor cells Kidney Podocytes Tubular cells Organoids Chronic kidney disease
Background
Kidney diseases, a global health issue, are the consequence of injury to the functional components of the kidney, the nephrons (Romagnani et al., 2017). Nephrons are constituted by a blood filtering unit, the glomerulus, and the respective tubule where the filtrate is modified by solute reabsorption and metabolite secretion up to when the final urine is excreted via the urinary tract (Romagnani et al., 2017). The nephrons respond to injury in two ways: a) differentiated epithelial cells undergo polyploidization and hypertrophy to rapidly support residual kidney function and b) immature epithelial cells, referred to as kidney progenitors (Lazzeri et al., 2019), proliferate to recover at least a part of the lost cells, i.e., kidney regeneration (Lazzeri et al., 2019). Kidney progenitors are localized along the inside of the Bowman capsule of the glomerulus and are scattered among tubular epithelial cells along the tubule, being identified by the expression of the surface markers CD133 and CD24, in humans (Sagrinati et al., 2006; Lazzeri et al., 2019). Kidney progenitors can be obtained from kidney tissue or urine and cultured long term (Angelotti et al., 2012) because they retain the capacity for self-renewal. Kidney progenitors have the capacity to differentiate into multiple types of kidney epithelial cells in vitro and in vivo (Sagrinati et al., 2006; Lazzeri et al., 2019). Hence, kidney progenitors can be expanded and differentiated into different types of tubular epithelial cells (Angelotti et al., 2012) and even cultured in 3D, generating tubuloids, a selective property among other kidney tubular cells (Xu et al., 2022). This property makes them ideal for modeling of genetic tubular disorders, e.g., upon isolation from the urine of patients with genetic tubular disorders or upon introduction of pathogenic genetic variants (e.g., using CRISPR-Cas system) (Xu et al., 2022). In addition, kidney progenitors can be differentiated in culture into podocytes, the main constituent of the glomerular filtration barrier. Podocytes are highly differentiated postmitotic cells unable to proliferate (Kopp et al., 2020). For this reason, they are impossible to expand in primary cultures, unless using artificial systems of immortalization (Shankland et al., 2007).
Here, we report detailed protocols on how to prepare human kidney progenitor cultures from human kidney tissue, maintain them, and differentiate them into podocytes. Differentiation of kidney progenitors using specific factors and compounds (Lasagni et al., 2015), as recently reported for the histone deacetylase inhibitor panobinostat, induces a change in their phenotype, promoting transcription of podocyte genes such as nephrin, podocin, and synaptopodin (Melica et al., 2022). We also report methods to assess their phenotype by qRT-PCR, FACS, and confocal and stimulated emission depletion (STED) microscopy. Applying the same culturing method described here to the isolation procedure reported by Lazzeri et al. (2015) permits the preparation of kidney progenitor cultures also from the urine of patients with kidney disorders, making them particularly suitable for studying genetic podocytopathies for diagnostic purposes. Given the importance of kidney progenitors and podocytes in the pathogenesis of chronic kidney disease, the possibility to prepare and maintain these cultures has wide implications and possible uses.
Materials and reagents
Biological materials
Normal-appearing kidney fragments are obtained from the pole opposite to the tumor from patients that underwent nephrectomy for localized renal tumors.
CAUTION: All the experiments involving human specimens should be performed in accordance with the recommendations of the Institutional Ethical Committee for human experimentation. All procedures in this protocol were conducted under protocols approved by the Ethical Committee on human experimentation of the Careggi University Hospital.
Reagents
Physiological saline solution, NaCl 0.9% (B. Braun Melsungen AG, A.I.C. n. 030902391)
HyClone defined fetal bovine serum (FBS), US origin (Cytiva, catalog number: SH30070.03)
Endothelial cell growth basal medium (EBM), 500 mL (LONZA, catalog number: CC-3121)
Microvascular endothelial growth medium (EGM-MV) SingleQuots kit (LONZA, catalog number: CC-4123) [contains growth factors, cytokines, and supplements: bovine brain extract w/o heparin (BBE); hydrocortisone (hEGF); gentamicin, amphotericin B (GA-1000); and FBS]
DMSO (Merck KGaA, catalog number: D8418)
Dulbecco’s modified Eagle’s medium/Ham’s nutrient mixture F-12 (DMEM-F12-1L) (Merck KGaA, catalog number: D2906)
Collagenase type IV (Sigma, catalog number: C-5138)
Panobinostat, LBH589 (MedChem Express, catalog number: HY-10224)
Trypsin/EDTA solution 0.025% (LONZA, catalog number: CC-5012)
Sodium hydrogen carbonate (NaHCO3) (Merck KGaA, catalog number: S5761)
Bi-distilled water (ddH2O)
D-PBS, no calcium, no magnesium (Gibco, catalog number: 14190-094)
Ammonium chloride (NH4Cl) (Carlo Erba, catalog number: 419416, CAS number: 12125-02-9)
FcR blocking reagent human (Miltenyi Biotec, catalog number: 130-059-901)
Bovine serum albumin (BSA) (Merck KGaA, catalog number: A9747)
Sodium azide (NaN3) (Merck KGaA, catalog number: S2002, CAS number: 26628-22-8)
RNeasy Micro kit (Qiagen, catalog number: 74004)
TaqMan reverse transcription reagents (Invitrogen, catalog number: N8080234)
TaqMan Fast Universal PCR Master Mix (2×), no AmpEraseTM UNG (Applied Biosystems, catalog number: 4352042)
Human GAPD (GAPDH) endogenous control (VICTM/TAMRATM probe, primer limited) (Applied Biosystems, catalog number: 4310884E)
TaqMan Gene Expression Assay Mix (20×) (Table 1)
Table 1. TaqMan assay list
Gene symbol Gene name TaqMan assay ID
NPHS1 NPHS1, nephrin Hs00190446_m1
NPHS2 NPHS2, podocin Hs00387817_m1
SYNPO Synaptopodin Hs00200768_m1
KLF15 Kruppel like factor 15 Hs00362736_m1
Paraformaldehyde solution 4% in PBS (PFA) (Santa Cruz, catalog number: sc-281692)
Triton X-100 (Merck KGaA, catalog number: X100RS-SG)
4’,6-Diamidino-2-phenylindole (DAPI) (Merck KGaA, catalog number: D9542)
Goat serum (Vector Laboratories, catalog number: S-1000)
Donkey serum (Merck KGaA, catalog number: D9663)
Antibodies used for FACS assay and immunofluorescence (Table 2)
Table 2.Antibodies list
Antibody Vendor Catalog number Final concentration
Primary antibody
CD133/2 Miltenyi Biotec 130-090-851 10 μL/test
CD24 (clone SN3) Santa Cruz SC-19585 10 μg/mL
mouse IgG1 Miltenyi Biotec 130-106-545 10 μg/mL
mouse IgG2b Miltenyi Biotec 130-106-547 10 μg/mL
α-Tubulin Merck KGaA T6074 2 μg/mL
Sir-Actin Spirochrome SC001 1 μM
Nephrin (NPHS1) R&D system AF4269 4 μg/mL
Podocin (NPHS2) Abcam ab50339 30 μg/mL
Secondary antibody
Goat anti-mouse IgG1-Alexa Fluor 488 Molecular Probes A-21121 6 μg/mL
Goat anti-mouse IgG2b-Alexa Fluor 647 Molecular Probes A-21242 6 μg/mL
Goat anti-mouse IgG1-Alexa Fluor 594 Molecular Probes A-21125 20 μg/mL
Donkey anti-sheep IgG(H+L)-Alexa Fluor 488 Molecular Probes A-11015 2 μg/mL
Goat anti-rabbit IgG(H+L)-Alexa Fluor 546 Molecular Probes A-11035 2 μg/mL
Kidney progenitor cell growth medium (see Recipes)
Kidney progenitor cell washing medium (see Recipes)
Red blood cell lysis buffer (NH4Cl 0.08%) (see Recipes)
Freezing medium (see Recipes)
DMEM-F12 + 10% HyClone defined FBS (see Recipes)
FACS buffer (PBS 1× 0.5% BSA - 0.02% NaN3 buffer) (see Recipes)
Equipment
Pipettes
Vacuum filtration system with 0.22 μm cellulose acetate (CA) membrane, 500 mL filters (Corning, catalog number: 430769).
Sharp forceps, straight (2-biol, catalog number: 91156-11)
Sterile plates 100 mm × 20 mm (Corning, catalog number: 430167)
Cell dissociation sieves (Merck KGaA, catalog number: S1145)
Screen for cell dissociation, size 80 mesh screens (Merck KGaA, catalog number: S3770)
Screen for cell dissociation, size 60 mesh screens (Merck KGaA, catalog number: S1020)
Glass pestle
Ice bucket
6-well clear TC-treated multiple well plates (Corning, catalog number: 3516)
Cryogenic vial (Corning, catalog number: CC430659)
Freezing container (Thermo Scientific, catalog number: 5100-0001)
75 cm2 flask (Corning, catalog number: CC430641)
Polypropylene urine container, 120 mL (Biosigma, catalog number: BSC258)
Serological pipettes 2 mL (Corning, catalog number: CLS4486)
Serological pipettes 5 mL (Corning, catalog number: CLS4487)
Serological pipettes 10 mL (Corning, catalog number: CLS4488)
15 mL tube (Corning, catalog number: 430791)
50 mL tube (Corning, catalog number: 430829)
1.5 mL microcentrifuge tubes (Axygen, catalog number: MCT-150-C-S)
0.2 mL RNase-free PCR tubes (Invitrogen, catalog number: AM12225)
MicroAmp fast optical 96-well reaction plate with barcode (Applied Biosystems, catalog number: 4346906)
MicroAmp optical adhesive film (Applied Biosystems, catalog number: 4360954)
GeneExplorer thermal cycler 96 × 0.2 mL (Bioer, catalog number: GE-96G)
7900HT Fast Real-Time PCR system (Applied Biosystems, catalog number: 4351405)
Centrifuge with plate holders
Inverted phase contrast microscope (Zeiss, Z-AXIO40C)
Heracell 150i CO2 incubator
Bürker counting chamber (Merck KGaA, catalog number: BR719505-1EA)
Flow cytometer (Miltenyi Biotec, MacsQuant Analyzer)
2-well chamber slide coverslip (Nunc Lab-Tek II, catalog number: 155379PK)
Confocal microscope (Leica Microsystems, LEICA SP8 STED 3X confocal microscope)
Biological safety cabinet (Angelantoni Life Science Srl, catalog number: CTH48C2)
4 °C fridge, -20 °C freezer, and -80 °C freezer
Software
Flow Cytometry Analysis software (Inivai, Flowlogic software)
Confocal microscope acquisition software Las X (Leica Microsystems)
Huygens Professional software version 18.04 (Scientific Volume Imaging B.V.)
Procedure
Human kidney progenitor cells (KPC): isolation, maintenance, and cryopreservation
In this section, we describe how to isolate primary kidney progenitor cells from human tissue. The method we describe exploits the ~50-fold higher proliferative capacity of KPC cells in comparison to other renal cell types in a specific growth medium (Peired et al., 2020). Based on our experience, this method allows to obtain a pure population of viable kidney progenitors more easily than the multi-step process based on separation using magnetic beads.
Isolation of KPC from human kidney tissue
Collect a fragment of kidney cortex (from 1 to 3 cm3) from the pole opposite to the tumor. Store the tissue in sterile physiological saline solution during transport to the laboratory. We recommend performing the kidney cell isolation within 1 h after surgical tissue collection.
Remove the kidney capsule and transfer the sample to a 100 mm sterile dish. Mince the cortex in pieces as small as possible using a scalpel. Add to the dish 5 mL of 750 U/mL collagenase type IV prepared in EBM medium. Incubate for 45 min at 37 °C in the incubator. Neutralize the enzymatic reaction by adding 10 mL of EBM containing 10 % FBS. CAUTION: During mincing, maintain the tissue fragments humidified by adding a drop of sterile physiological saline solution.
Transfer the suspension to graded mesh screens (60 and 80 mesh). Mechanically break down the tissue suspension using a glass pestle and pass it through the 60 and 80 mesh screens. Wash thoroughly with 20 mL of kidney progenitor cell washing medium and recover the flowthrough in a polypropylene urine container. Transfer this suspension to a 50 mL polypropylene tube and centrifuge at 400× g for 5 min at 4 °C.
Aspirate the supernatant and wash the pellet with 5 mL of PBS. Centrifuge at 400× g for 5 min at 4 °C. Discard the supernatant and resuspend the pellet in 5 mL of red blood cell lysis buffer.
Incubate for 4 min at 37 °C and then stop the reaction by adding 10 mL of kidney progenitor cell washing medium.
Centrifuge at 400× g for 5 min at 4 °C.
Remove supernatant and resuspend the pellet in 10 mL of kidney progenitor cell growth medium.
Count cell suspension using a Bürker counting chamber.
Transfer the cells in 75 cm2 flask (500,000 cells/flask) in 8 mL/flask kidney progenitor cell growth medium. Label as passage 0.
Place the cells in a 5% CO2, 37 °C incubator.
After three days, replace the medium with fresh kidney progenitor cell growth medium to remove unattached cells and continue with twice-a-week changes until cells reach 80% confluency. It usually takes 7–10 days.
Expansion and sub-culturing of KPC
Sub-culture when the cells are approximately 80% confluent.
Aspirate the medium and wash with 6 mL of PBS.
Aspirate PBS and add 2 mL of a 0.25 mg/mL trypsin/EDTA solution. Incubate in a 37 °C incubator for approximately 5 min. Check under the microscope if cells are detached.
Neutralize the enzymatic reaction by adding 4 mL of kidney progenitor cell washing medium and collect the cells in a 15 mL polystyrene tube. Centrifuge the cells at 400× g for 5 min at 4 °C.
Aspire supernatant without disturbing the cell pellet and resuspend the cells in 5 mL of kidney progenitor cell growth medium.
Count the number of cells using a Bürker counting chamber.
Replate the cells in 75 cm2 flasks with a ratio of 1:3 in kidney progenitor cell growth medium.
Change medium twice a week during maintenance of cultures in a 5% CO2, 37 °C incubator.
Cryopreservation of KPC
Kidney progenitors are cryo-stored in 1 mL of freezing medium at a density from 5 × 105 up to 1 × 106 cells/cryogenic vial.
Aspirate the medium from the 75 cm2 flask and wash with 6 mL of PBS.
Aspirate the PBS and add 2 mL of a 0.25 mg/mL trypsin/EDTA solution. Incubate in a 37 °C incubator for approximately 5 min. Check under the microscope if cells are detached.
Neutralize the enzymatic reaction by adding 4 mL of kidney progenitor cell washing medium and collect the cells in a 15 mL polystyrene tube. Centrifuge the cells at 400× g for 5 min at 4 °C.
Aspire supernatant without disturbing the cell pellet and resuspend the cells in 5 mL of kidney progenitor cell growth medium.
Count the number of cells using a Bürker counting chamber.
Centrifuge the cells at 400× g for 5 min at 4 °C.
Resuspend the cell pellet in freezing medium at a density of 1 × 106 cells/mL. Mix well.
Aliquot in 1 mL per cryovial.
Transfer the cryovials to a freezing container and put the freezing container into a -80 °C freezer.
The next day, transfer the cryovials to a liquid nitrogen tank for long-time storage. CAUTION: Minimize as much as possible the time cells remain in freezing medium at room temperature. Transfer immediately in the freezing container to -80 °C.
Thawing of kidney progenitor cells
Thaw a vial of cells. To achieve rapid warming, place the frozen vial into a 37 °C water bath.
Transfer immediately the content of each vial to a 15 mL tube containing 4 mL of kidney progenitor cell washing medium.
Centrifuge at 400× g for 5 min at 4 °C.
Remove the supernatant and resuspend the cells into 2 mL of kidney progenitor cell growth medium.
Transfer the cell suspension to 75 cm2 flasks at a density of approximately 500,000 cells/flask in 8 mL/flask of kidney progenitor cell growth medium. Place the cells in a 5% CO2, 37 °C incubator.
Differentiation of KPC into podocytes
Detach the cells at 60%–80% confluency with trypsin as described above.
Count cells and plate in a 6-well plate at a density of 80,000 cell/well in 1.5 mL/well of kidney progenitor cell growth medium.
Place the cells in a 5% CO2, 37 °C incubator.
After 5–6 h (or when cells are attached to the plate), gently remove the medium and replace with 1.5 mL/well of EBM without any supplement and without serum. Place the cells in a 5% CO2, 37 °C incubator.
After 16 h, remove the EBM medium and stimulate the cells for 48 h with 1.5 mL/well of differentiation medium containing 0.2 μM panobinostat in DMEM-F12 + 10% HyClone Defined FBS.
At the end of differentiation, characterize the cells using qRT-PCR and immunofluorescence. CAUTION: To obtain better differentiation results, use cells at early passages (P1–P2).
Data analysis
Flow cytometry analysis for purity check
It is important to characterize each primary kidney progenitor cell culture, evaluating in the various passages (from passage P0 to at least passage P3) the expression of surface markers CD133 and CD24. To evaluate CD133 and CD24 expression, perform flow cytometry analysis as reported:
Detach the cells with trypsin as described above.
Count cells.
Prepare two 1.5 mL tubes: label one Isotype control and the other Antibody (CD133 and CD24).
Transfer 100,000 cells in each tube.
Centrifuge at 400× g for 5 min at 4 °C.
Aspirate supernatant without disturbing the cell pellet. Add 5 μL of FcR blocking reagent human on cell pellet.
Prepare the staining solutions as follows:
Isotype control mix I: Add 1 μL of mouse IgG2b (to obtain a final concentration of 10 μg/mL) and 1 μL of mouse IgG1 (to obtain a final concentration of 10 μg/mL) in 98 μL of FACS buffer.
Antibody mix I: Add 10 μL of CD133/2 (10 μL/test) antibody and 2 μL of CD24 (to obtain a final concentration of 10 μg/mL) antibody in 28 μL of FACS buffer.
Add 30 μL of the staining solutions to the corresponding tube containing cells and FcR blocking reagent human. Resuspend the pellet.
Incubate on ice for 15–30 min covered with a tin foil.
Add 500 μL of FACS buffer.
Centrifuge at 400× g for 5 min at 4 °C and aspirate the supernatant without disturbing the cell pellet.
Resuspend each pellet in 30 μL of the staining solution II, prepared as follows:
Staining Solution II: add 1 μL of goat anti-mouse IgG2b-647 (to obtain a final concentration of 5 μg/mL) and 1 μL of goat anti-mouse IgG1-488 (to obtain a final concentration of 5 μg/mL) to 300 μL of FACS buffer.
Incubate on ice for 15–30 min covered with a tin foil.
Add 500 μL of FACS buffer.
Run FACS assay by using MacsQuant Analyzer.
Analyze the FACS data using Flowlogic software. A representative FACS assay for kidney progenitor cells is shown in Figure 1. CAUTION: Use primary cultures consisting of at least 95% of CD133 and CD24 double-positive cells. The percentage of double-positive cells tend to increase during the first two passages because the kidney progenitor cell growth medium allows the selective growth of undifferentiated kidney progenitors.
Figure 1. Evaluation of CD133/2 and CD24 expression in kidney progenitor cells by flow cytometry. Representative flow cytometry dot plot graphs showing the percentage of CD133 and CD24 positive cells in primary kidney progenitor cells at passage P1 (B). Staining of the same cells with isotype control antibodies is shown in (A).
Evaluation of differentiation
After 48 h of stimuli with differentiation medium, it is possible to evaluate differentiation status of the cells by using qRT-PCR and immunofluorescence (Figure 2).
Figure 2. Evaluation of podocytes derived from kidney progenitor cells. Representative phase contrast images of (A) undifferentiated kidney progenitor cells and (B) podocytes derived from kidney progenitor cells after 48 h of differentiation. Scale bars, 100 μm. (C) qRT-PCR assay of the podocyte markers NPHS1, NPHS2, KLF15, and SYNPO in undifferentiated kidney progenitor cells and in podocytes derived from kidney progenitor cells. mRNA expression of the podocytes markers was determined by qRT-PCR and reported as mean ± SEM of fold increase over undifferentiated cells. (D, E) Representative confocal microscopy images showing expression of the podocyte markers NPHS1 and NPHS2 (red) in undifferentiated kidney progenitor cells and in podocytes derived from kidney progenitor cells. DAPI (white) was used to counterstain nuclei. Scale bars, 25 μm. KPC, kidney progenitor cell.
qRT-PCR assay
One of the methods to check the differentiation status of the cells is to use real-time PCR assay to evaluate mRNA expression level of podocyte markers, such as NPHS1, NPHS2, KLF15, and SYNPO. We perform qRT-PCR for each marker on the RNA extracted from an equal number of undifferentiated and differentiated cells independently from the RNA concentration obtained. The qRT-PCR protocol is detailed below:
Collect pellets from 100,000 undifferentiated and differentiated cells and extract RNA using the RNeasy Micro kit, followed by the DNase digestion protocol in a final volume of 15 μL per sample.
Proceed to the synthesis of the cDNA using the TaqMan reverse transcription reagents:
Prepare the following cDNA synthesis mix on ice in a 0.2 mL PCR tube. Mix thoroughly and centrifuge briefly.
Component Volume for reaction
10× buffer 5 μL
25 mM MgCl2 3.5 μL
dNTPs 10 μL
RNase inhibitor 2.5 μL
Random hexamers 2.5 μL
Multiscribe (MULV) 2.5 μL
RNA 15 μL
RNase-free water To 50 μL
Proceed with the following incubation protocol in a thermal cycler.
Step Temperature Run time
1 25 °C 10 min
2 48 °C 30 min
3 95 °C 5 min
4 4 °C Hold
Collect cDNA synthesis product for qPCR or store in a -20 °C freezer.
Prepare the qPCR mix for each podocyte gene (NPHS1, NPHS2, KLF15, or SYNPO) on ice by adding the components below. Mix extra 10% for more reactions. Mix thoroughly and centrifuge briefly.
Component Volume for one reaction
TaqMan Fast Universal PCR Master Mix (2×) 10 μL
TaqMan Assay (NPHS1, NPHS2, KLF15, or SYNPO) 1 μL
ddH2O 4 μL
Final volume 15 μL
Prepare the qPCR mix for GAPDH housekeeping gene on ice by adding the components below. Mix extra 10% for more reactions. Mix thoroughly and centrifuge briefly.
Component Volume for one reaction
TaqMan Fast Universal PCR Master Mix (2×) 10 μL
GAPDH endogenous control 1 μL
ddH2O 8 μL
Final volume 19 μL
Dispense 15 μL of the mix for NPHS1, NPHS2, KLF15, or SYNPO into each well of a 96-well PCR plate. For GAPDH, dispense 19 μL of the mix.
Add 5 μL of cDNA samples to the well containing the mix for NPHS1, NPHS2, KLF15, or SYNPO and add 1 μL of cDNA samples to the well containing the mix for GAPDH.
Seal the plate and centrifuge briefly. Proceed to the incubation below by selecting fast mode:
Step Temperature Run time
1 95 °C 20 s
2 95 °C 1 s
3 60 °C 20 s + data collection
4 Repeat steps C2–C3 40 times
Perform data analysis: gene expression of each marker is normalized to that of the GAPDH. For each marker, results are reported as fold change of expression of the differentiated cells over undifferentiated cells (Figure 2).
Immunofluorescence
A differentiative program induces morphology changes and leads to cell-specific protein expression that can be evaluated by immunofluorescence assay. The differentiation of kidney progenitor cells into podocytes is confirmed on the basis of the NPHS1 and NPHS2 podocyte marker expression (Figure 2), while the tubulin and actin expression assessed by super-resolution microscopy shows the drastic cytoskeleton changes associated with differentiation (Figure 3). The immunofluorescence procedure is detailed below:
Detach the cells with trypsin as described above.
Count cells.
Plate cells onto a 2-well chamber slide at a density of 20,000 cell/well in 1 mL/well of kidney progenitor cell growth medium.
Place the cells in a 5% CO2, 37 °C incubator.
After 5–6 h (or when cells are attached to the plate), gently remove the medium and replace with 1.5 mL/well of EBM without any supplement and without serum. Place the cells in the incubator with 5% CO2 and 37 °C.
After 16 h, remove the EBM medium and stimulate cells for 48 h with 1 mL/well of 0.2 μM panobinostat in DMEM-F12 + 10% HyClone defined FBS.
At the end of the differentiation, remove chamber slides from the incubator.
Aspirate the medium and wash the cells with 500 μL/chamber of PBS.
Add 1 mL/chamber of 4% PFA and incubate for 20 min at room temperature.
Gently wash the slides three times with 1 mL of PBS.
Add permeabilizing solution (composed of 0.5% Triton X-100 in PBS) if required from the antibody user manual (antibody information is reported in Table 2 and Table 3) for 5 min at room temperature.
Table 3. Immunofluorescence details
Primary antibody Secondary antibody
Antibody Final concentration Antibody Final concentration Permeabilization Blocking serum required
α-Tubulin 2 μg/mL Goat anti-mouse IgG1-Alexa Fluor 594 20 μg/mL Required Goat
Sir-Actin 1 μM / / Required /
Nephrin (NPHS1) 4 μg/mL Donkey anti-sheep IgG(H+L)-Alexa Fluor 488 2 μg/mL Not required Donkey
Podocin (NPHS2) 30 μg/mL Goat anti-rabbit IgG(H+L)-Alexa Fluor 546 2 μg/mL Required Goat
Wash for 5 min with PBS.
Incubate with blocking solution containing 3% BSA and 0.3% serum (goat or donkey, as reported in Table 3) in PBS.
Remove blocking solution without washing.
Incubate with primary antibody (Table 3) for 15 min at 37 °C and subsequently for 1 h at 4 °C covered with a tin foil.
Wash for 5 min with PBS.
Incubate with the secondary antibodies listed in Table 3 and with 1 μg/mL DAPI in PBS 1× for 30 min at room temperature covered with a tin foil to block the light.
Acquire images using a LEICA SP8 STED 3X confocal microscope.
For STED analysis, frame sequential acquisition can be applied to avoid fluorescence overlap. A 775 nm pulsed-depletion laser was used and a gating between 0.3 and 6 ns was applied to avoid collection of reflection and autofluorescence. Images were acquired with Leica HC PL APO CS2 100×/1.40 oil STED white objective. De-convolve with Huygens Professional software (Romoli et al., 2018).
Figure 3. Stimulated emission depletion (STED) super-resolution microscopy shows cytoskeleton changes associated with differentiation. Representative STED images showing cytoskeleton changes associated with differentiation into podocytes based on α-Tubulin (green) and Actin (red) expression in primary human kidney progenitor cells before (A) and after 48 h differentiation (B–G). Compared to confocal microscopy (D, F), the use of STED microscopy and deconvolution software allows to identify the cytoskeleton organization with nanoscopic spatial resolution (C, E, and G).
Recipes
Kidney progenitor cell growth medium (EGM-MV medium + 20% HyClone defined FBS)
The renal progenitor cells are grown in EGM-MV medium supplemented with 20% HyClone defined FBS, prepared as follows:
Supplement 400 mL of EBM medium with BBE and hEGF provided in the microvascular endothelial growth medium (EGM-MV) SingleQuots kit and with 100 mL of HyClone defined FBS. Filter using a vacuum filtration system with 0.22 μm CA membrane and store at 4 °C.
Kidney progenitor cell washing medium (EBM medium + 10% FBS)
Supplement EBM medium with 10% FBS by using the serum provided with the SingleQuots kit (and not used for EGM-MV + 20% HyClone formulation). Other commercial FBS can be used. Filter the solution using a vacuum filtration system with 0.22 μm CA membrane.
Red blood cell lysis buffer (NH4Cl 0.08%)
Dissolve 0.4 g of NH4Cl in 500 mL of bi-distilled water. Filter using a vacuum filtration system with 0.22 μm CA membrane and store at 4 °C.
Kidney progenitor cell freezing medium
Immediately before freezing the cells, prepare a solution containing HyClone defined FBS supplemented with 10% DMSO.
DMEM-F12 + 10% HyClone defined FBS
Dissolve 15.6 g of powder DMEM-F12 (one vial of DMEM-F12) in 1 L of MilliQ water and supplement with 1.2 g/L NaHCO3. Filter using 0.22 μm filters and store at 4 °C. Add FBS HyClone at a final concentration of 10% (w/v) only for the volume of medium necessary for the experiment.
FACS BUFFER (PBS 1× 0.5% BSA - 0.02% NaN3 buffer)
Dissolve 2 g of NaN3 in 10 mL of PBS to obtain a 20% (w/v) NaN3 stock solution, which can be stored at room temperature for at least two years.
Dissolve 1.25 g of BSA in 250 mL of D-PBS, no calcium, no magnesium, and then add 250 μL of 20% NaN3 stock solution. Store at 4 °C.
Acknowledgments
This protocol was derived from the original work of Melica et al. (2022).
This study was funded by the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (grant agreement no. 101019891). M.E.M. was supported by a FIRC-AIRC fellowship for Italy.
Competing interests
The authors have no competing financial interests.
Ethics considerations
Normal-appearing kidney fragments were obtained from the pole opposite to the tumor of patients who underwent nephrectomy for renal tumors, in agreement with the Ethical Committee on human experimentation of the Careggi University Hospital.
References
Angelotti, M. L., Ronconi, E., Ballerini, L., Peired, A., Mazzinghi, B., Sagrinati, C., Parente, E., Gacci, M., Carini, M., Rotondi, M., et al. (2012). Characterization of renal progenitors committed toward tubular lineage and their regenerative potential in renal tubular injury. Stem Cells 30(8): 1714-1725.
Kopp, J. B., Anders, H. J., Susztak, K., Podesta, M. A., Remuzzi, G., Hildebrandt, F. and Romagnani, P. (2020). Podocytopathies. Nat Rev Dis Primers 6(1): 68.
Lasagni, L., Angelotti, M., Ronconi, E., Lombardi, D., Nardi, S., Peired, A., Becherucci, F., Mazzinghi, B., Sisti, A., Romoli, S., et al. (2015). Podocyte Regeneration Driven by Renal Progenitors Determines Glomerular Disease Remission and Can Be Pharmacologically Enhanced. Stem Cell Reports 5(2): 248-263.
Lazzeri, E., Angelotti, M. L., Conte, C., Anders, H. J. and Romagnani, P. (2019). Surviving Acute Organ Failure: Cell Polyploidization and Progenitor Proliferation. Trends Mol Med 25(5): 366-381.
Lazzeri, E., Ronconi, E., Angelotti, M. L., Peired, A., Mazzinghi, B., Becherucci, F., Conti, S., Sansavini, G., Sisti, A., Ravaglia, F., et al. (2015). Human Urine-Derived Renal Progenitors for Personalized Modeling of Genetic Kidney Disorders. J Am Soc Nephrol 26(8): 1961-1974.
Melica, M. E., Antonelli, G., Semeraro, R., Angelotti, M. L., Lugli, G., Landini, S., Ravaglia, F., Regina, G., Conte, C., De Chiara, L., et al. (2022). Differentiation of crescent-forming kidney progenitor cells into podocytes attenuates severe glomerulonephritis in mice. Sci Transl Med 14(657): eabg3277.
Peired, A. J., Antonelli, G., Angelotti, M., Allinovi, M., Guzzi, F., Sisti, A., Semeraro, R., Conte, C., Mazzinghi, B., Nardi, S., et al. (2020). Acute kidney injury promotes development of papillary renal cell adenoma and carcinoma from renal progenitor cells. Sci Transl Med 12(536): eaaw6003.
Romagnani, P., Remuzzi, G. Glassock, R. Levin, A., Jager, K. J., Tonelli, M., Massy, Z., Wanner, C. and Anders, H. J. (2017). Chronic kidney disease. Nat Rev Dis Primers 3: 17088.
Romoli, S., Angelotti, M. L., Antonelli, G., Kumar Vr, S., Mulay, S. R., Desai, J., Anguiano Gomez, L., Thomasova, D., Eulberg, D., Klussmann, S., et al. (2018). CXCL12 blockade preferentially regenerates lost podocytes in cortical nephrons by targeting an intrinsic podocyte-progenitor feedback mechanism. Kidney Int 94(6): 1111-1126.
Sagrinati, C., Netti, G. S., Mazzinghi, B., Lazzeri, E., Liotta, F., Frosali, F., Ronconi, E., Meini, C., Gacci, M., Squecco, R., et al. (2006). Isolation and characterization of multipotent progenitor cells from the Bowman's capsule of adult human kidneys. J Am Soc Nephrol 17(9): 2443-2456.
Shankland, S. J., Pippin, J. W., Reiser, J. and Mundel, P. (2007). Podocytes in culture: past, present, and future. Kidney Int 72(1): 26-36.
Xu, Y., Kuppe, C., Perales-Patón, J., Hayat, S., Kranz, J., Abdallah, A. T., Nagai, J., Li, Z., Peisker, F., Saritas, T., et al. (2022). Adult human kidney organoids originate from CD24+ cells and represent an advanced model for adult polycystic kidney disease. Nat Genet 54(11): 1690-1701.
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Measuring Myeloperoxidase Activity as a Marker of Inflammation in Gut Tissue Samples of Mice and Rat
NH Nikita Hanning
JM Joris G. De Man
BW Benedicte Y. De Winter
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4758 Views: 800
Reviewed by: Pilar Villacampa AlcubierreRajesh ThippeshappaMarieta Ruseva
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Abstract
Myeloperoxidase (MPO) is an enzyme contained in lysosomal azurophilic granules of neutrophils. MPO activity has been shown to correlate with the number of neutrophils in histological sections of the gastrointestinal tract and is therefore accepted as a biomarker of neutrophil invasion in the gut. This protocol describes an easy, cost-effective kinetic colorimetric assay to quantify myeloperoxidase activity in intestinal tissue samples. It is explained using tissue collected in mice but can also be used for other laboratory animals. In a first step, tissue specimens are homogenized using a phosphate buffer containing 0.5% hexadecyltrimethylammonium bromide (HTAB), which extracts MPO from neutrophils. The obtained supernatant is added to a reagent solution containing o-dianisidine dihydrochloride, which is a peroxidase substrate. Finally, the change in absorption is measured via spectrophotometry and converted to a standardized unit of enzyme activity. The assay is illustrated and compared to a commercially available enzyme-linked immunoassay (ELISA), demonstrating that MPO activity does not necessarily correlate with MPO protein expression in tissue samples.
Key features
• Optimized for use in mice and rats but can also be used for samples of other species.
• Measures enzymatic activity instead of mRNA or protein expression.
• Requires a spectrophotometer.
• Can be performed in duplo using 10 mg of (dry-blotted) gut tissue or more.
Graphical overview
Keywords: Colitis Enteritis IBD Inflammation Intestines Myeloperoxidase activity Neutrophils Peroxidase
Background
The presence of inflammatory infiltrates in the gastrointestinal tract is one of the hallmark characteristics of diseases such as ulcerative colitis and Crohn’s disease (Xavier and Podolsky, 2007). The pathogenesis of these diseases is frequently studied in rodents exposed to colitis-inducing chemicals such as acetic acid, dextran sodium sulfate (DSS), and 2,4,6-trinitrobenzene sulfonic acid (TNBS), or in genetically engineered mice that are prone to develop gastrointestinal inflammation spontaneously (Mizoguchi, 2012). To assess which factors contribute to the development, maintenance, and resolution of inflammation in the gut, the availability of sensitive assays to quantify this inflammation is essential.
Several methods are used to quantify the presence of inflammatory infiltrates in the gut. First, the degree of inflammation can be scored by quantifying the shortening and thickening of the intestinal wall via the weight:length ratio of the relevant region within the gastrointestinal tract (Solomon et al., 2010). Second, scoring systems can be used to assess the presence of ulcerations at an endoscopic, macroscopic, or microscopic level (Solomon et al., 2010; Heylen et al., 2013; Erben et al., 2014). Third, quantitative polymerase chain reactions (qPCR), bead-based immunoassays, or conventional enzyme-linked immunoassays (ELISAs) can be used to quantify the expression of pro-inflammatory cytokines such as tumor necrosis factor α (TNF-α) or interleukin 1β (IL-1β) (Khan et al., 2004; Yan et al., 2009; Kim et al., 2012). Finally, myeloperoxidase (MPO) activity can be measured as a biomarker of inflammation. MPO is a peroxidase contained within lysosomal azurophilic granules in neutrophils and released upon activation of the immune system (Schultz and Kaminker, 1962). It has been shown to correlate well with the number of neutrophils in histological sections and is therefore a simple, objective biochemical alternative to the labor-intensive histological analysis of inflammatory infiltrates (Krawisz et al., 1984).
The presence of MPO in tissue samples can be assessed in different ways. First, expression at the mRNA or protein level can be measured by means of qPCR or commercially available sandwich ELISAs, respectively. Second, kinetic assays can be used to evaluate enzymatic activity. In this protocol, we describe a simple, cost-effective kinetic colorimetric assay to determine MPO activity in intestinal tissue supernatant of mice. In addition, we compare our assay to an ELISA, demonstrating that the MPO concentration of the tissue does not necessarily reflect its enzymatic activity. The protocol consists of several steps. First, obtained tissue samples are homogenized in a potassium phosphate buffer containing 0.5% hexadecyltrimethylammonium bromide (HTAB), which solubilizes the enzyme. Next, the supernatant is added to a reagent solution containing o-dianisidine dihydrochloride. This peroxidase substrate turns brownish upon oxidation and can be detected via spectrophotometry (λ = 460 nm).
This protocol was developed and used to measure MPO activity in intestinal tissues of mice and rat (Moreels et al., 2001; Ruyssers et al., 2009; Vermeulen et al., 2011). It could, however, be easily adapted to measure MPO activity in human gastrointestinal biopsies, or in samples derived from other organs. However, spectrophotometric interference by tissue myoglobin or vascular hemoglobin can occur, requiring adaptations of the protocol for organs such as the heart or brain (Kuebler et al., 1996; Xia and Zweier, 1997).
Materials and reagents
Reagents
di-Potassium hydrogen phosphate (K2HPO4) (VWR, catalog number: 71003, CAS number: 7758-11-4), store at room temperature
Dulbecco’s phosphate buffered saline (DPBS), including calcium and magnesium (Gibco, catalog number: 14040133), store at room temperature
Hexadecyltrimethylammonium bromide (Sigma-Aldrich, catalog number: H5882, CAS number: 57-09-0), store at room temperature
Hydrogen peroxide 30% (H2O2) (Merck, catalog number: 107209, CAS number: 7722-84-1), store at 4 °C
o-Dianisidine dihydrochloride (Sigma-Aldrich, catalog number: D3252, CAS number: 20325-40-0), store at 4 °C
Potassium dihydrogen phosphate (KH2PO4) (Merck, catalog number: 104873, CAS number: 7778-77-0), store at room temperature
Sodium chloride 0.9% for injection (NaCl) (B. Braun, catalog number: 76363, CAS number: 7647-14-5), store at room temperature
Deionized water (H2O), prepared using the Elix® water purification system (see Equipment), store at room temperature
Solutions
Potassium phosphate buffer (pH = 6.0) (see Recipes)
Homogenization buffer (see Recipes)
Assay reagent (see Recipes)
Recipes
Potassium phosphate buffer (pH = 6.0)
Prepare a 50 mM KH2PO4 solution by dissolving 6.805 g of KH2PO4 in 1 L of deionized H2O.
Prepare a 50 mM K2HPO4 solution by dissolving 5.706 g of K2HPO4 in 0.5 L of deionized H2O.
Adjust the pH of the KH2PO4 solution by gradually adding the 50 mM K2HPO4 solution, until a pH of 6.0 is reached.
Homogenization buffer
Dissolve 2.5 g of hexadecyltrimethylammonium bromide in 500 mL of potassium phosphate buffer.
Assay reagent (light sensitive; use amber bottles wrapped in silver foil to store this solution)
Prepare a 1% H2O2 solution by adding 100 μL of a 30% H2O2 solution to 2,900 μL of the potassium phosphate buffer.
Prepare a 0.05% H2O2 solution by adding 200 μL of a 1% H2O2 solution to 3,800 μL of the potassium phosphate buffer.
Dissolve 66.8 mg of o-dianisidine dihydrochloride in 4 mL of 0.9% NaCl solution.
Prepare the assay reagents by mixing 4 mL of 0.05% H2O2 solution, 4 mL of the o-dianisidine dihydrochloride solution, and 392 mL of the potassium phosphate buffer.
Laboratory supplies
Cryotubes Cryo.STM, 2 mL (Greiner Bio-One, catalog number: 126263)
Precellys® tubes, 2 mL (Bertin Technologies, catalog number: 9000538)
Cuvettes, 4.2 mL (Sarstedt, catalog number: 67.741)
Reusable oral gavage needle, 10 G × 25 mm (PetSurgical, catalog number: AFN2425S)
Syringe with luer lock tip, 5 mL (Terumo, catalog number: SS*05SE1)
Dissection material (scissors, fine tweezers)
Equipment
Elix® water purification system (Millipore, catalog number: ZLXS50020)
Progard® TL1 Cl2 pretreatment tap water filter (Milli-Q, catalog number: PR0GTLCS1)
Handheld pH meter (Oakton Instruments, model: pH 6+, catalog number: 35613-22)
Precellys® 24 tissue homogenizer (Bertin Technologies, catalog number: P000669-PR240-A)
Centrifuge (Eppendorf, catalog number: 5425R)
UV/Vis Spectrometer (ATI Unicam, model: UV2-100)
Software
Excel® (Microsoft) or any other software suitable for basic data manipulation
Procedure
Tissue collection
Euthanize the mouse by cervical dislocation. Alternatively, a different humane method of euthanasia such as carbon dioxide inhalation can be used.
Carefully open the abdominal cavity without damaging the intestines by making a midline incision.
Remove ±3 cm of the relevant part of the gastrointestinal tract of the abdominal cavity, place it in a Petri dish containing ice-cold DPBS solution, and flush with a gavage needle attached to a 5 mL syringe containing DPBS to remove the luminal content. Alternatively, fecal material can be removed by gently squeezing the intestines with a pair of tweezers.
Clean the gut tissue by removing any remaining mesenteric fat and then cut open the intestines along the mesenteric border.
Collect a representative tissue sample (0.5 cm × 0.5 cm; ± 20–30 mg), dry it on a paper towel, weigh it, place it in a cryovial, and subsequently freeze the specimen in liquid nitrogen. To remain unbiased, it is good practice to always collect samples in the same region of the gastrointestinal tract (i.e., always take the most distal or proximal part of the colon). Examples of tissue samples of correct size are shown in Figure 1.
After collection of all mouse samples, store them at -80 °C until further experiments are performed.
Figure 1. Gut tissue samples of an appropriate size for the myeloperoxidase (MPO) assay. The exact size depends on the animal species and inflammatory state of the tissue. From left to right, representative samples of distal colon (25 mg), proximal colon (27 mg), ileum (22 mg), and glandular stomach (22 mg) of a healthy C57BL/6J mouse are shown.
Tissue homogenization
Take the tissue samples from the freezer, transfer them to Precellys® tubes, and place these on ice. When labeling the Precellys® tubes, place the identifier on the side of the tube since the homogenization procedure can make identifiers written on the cap illegible.
Use a pipette to add 0.02 mL/mg tissue of homogenization buffer (Recipe 2) to the samples.
Place the tubes in the homogenizer and dissociate the tissue samples (6,000 rpm; 20 s shaking > 20 s break > 20 s shaking > 60 s break).
Place the tubes on ice and wait for 30 min to allow for the foam, which is formed during the homogenization, to disappear.
Centrifuge the samples (18,000× g for 15 min at 4 °C).
Use a pipette to collect the supernatant in 1.5 mL Eppendorf tubes and place these on ice. The pellet can be discarded.
Myeloperoxidase activity assay
Start the UV/Vis spectrometer and specify the necessary settings:
Measurements are kinetic.
Absorbance is measured at a wavelength of 460 nm (bandwidth 2.0 nm).
The change in absorbance is measured during the first 60 s after the start of the chemical reaction.
Open the lid of the spectrophotometer and place a 3 mL cuvette in the cuvette holder of the device. Pay attention to the correct orientation of the cuvette, i.e., with the clear windows in line with the light source and detector.
Use a pipette to add 2,900 μL of assay reagent (Recipe 3) to the cuvette.
Use a pipette to add 100 μL of sample supernatant to the cuvette, mix the sample by repetitive pipetting (5–10 times using a pipette set at 2 mL), close the lid of the spectrophotometer, and then promptly start the spectrophotometric measurement.
After the 60-second measurement, note the absolute absorption values at 0 and 60 s in Excel.
Replace the cuvette by a clean one and repeat the procedure for the other samples. Ideally, the assay is performed in duplo or in triplo.
Data analysis
Calculation of MPO activity in tissue samples
Calculate the change in absorbance (ΔAbs) by subtracting the measured absorbance at 0 s from the absorbance at 60 s.
Derive the MPO activity of the sample by implementing Beer’s law:
where ΔC is the change in concentration of the oxidized form of o-dianisidine in μM/min; ΔAbs is the change in absorbance of the sample after 60 s; ϵ is the extinction coefficient of oxidized o-dianisidine, which has been determined to be 1.13 × 104 μM/cm for a wavelength of 460 nm at room temperature (Worthington and Worthington, 2011); and b is the optical path length, which is 1 cm for the cuvettes recommended in this protocol.
Express the MPO activity in units per gram of tissue. A unit of MPO is defined as the enzymatic activity that converts 1 μmol of hydrogen peroxidase to water in 1 min at room temperature and can be calculated using:
where Act is the enzymatic activity in unit per gram of tissue; ΔC is the change in concentration of the oxidized form of o-dianisidine in μM/min; Vreac is the volume of the supernatant that is used in the cuvette, which is 0.1 mL in the described protocol; and Csn is the concentration of the generated supernatant, which is 0.05 g of tissue per milliliter of homogenization buffer.
Note that for samples with very high activity levels, the supernatant could be further diluted to obtain accurate measures of enzyme activity.
Data example
To illustrate the assay, 12-week-old female C57BL/6J mice were exposed to 3% DSS for seven days via autoclaved drinking water to induce acute colitis (Kim et al., 2012; Breugelmans et al., 2020). Control mice received autoclaved drinking water without the addition of DSS. After a one-week treatment, DSS-treated mice showed signs of colitis, i.e., weight loss, diarrhea, and rectal bleeding. To confirm the inflammatory state of the animals, MPO activity was determined in tissue supernatant derived from the distal colon. To illustrate the importance of determining MPO activity instead of MPO content, a commercially available ELISA (Invitrogen, catalog number: EMMPO) was performed to quantify MPO protein expression levels on the same tissue supernatant, according to the instructions of the manufacturer.
MPO activity in the distal colon of DSS-treated mice was significantly higher than in control animals [DSS: 5.46 (4.05–8.14) U/g tissue vs. control: 0.30 (0.22–0.47) U/g tissue, p < 0.01 for n = 6 animals/group, Figure 2A]. Despite the increased peroxidase activity in the distal colon of DSS-exposed mice, the MPO protein expression did not differ between the groups [DSS: 42.67 (34.53–58.75) ng/g tissue vs. control: 45.05 (39.60–49.92) ng/g tissue, p = 0.78, Figure 2B]. Notably, MPO protein content and enzymatic activity were not correlated (rS = 0.30, p = 0.34, Figure 2C). Potentially, the capacity of gut tissue to inhibit peroxidase activity is lost upon the exposure to DSS (Ormrod et al., 1987).
Figure 2. MPO activity and protein expression levels in a mouse model for acute DSS-induced colitis. A) MPO activity, measured according to our protocol, is increased in DSS-exposed mice. B) MPO protein expression, measured using an ELISA kit, is not altered by exposure to DSS. C) MPO protein expression and enzymatic activity are not correlated. All quantifications were performed in duplo. Analysis by means of a Mann-Whitney U test (A, B) or Spearman’s rank correlation test (C) for n = 6 animals per group. DSS, dextran sulfate sodium; MPO, myeloperoxidase activity.
Validation of protocol
This MPO assay was previously used by our group to quantify inflammation in acute and chronic colitis models in both mice and rat (Moreels et al., 2004; Ruyssers et al., 2009; Vermeulen et al., 2011). We demonstrated that MPO activity correlated well with other markers for inflammation in TNBS-induced colitis in the rat (Vermeulen et al., 2011) and in the adoptive transfer colitis model in mice (Heylen et al., 2013).
General notes and troubleshooting
For the tissue homogenization to be successful, it is best to avoid the use of large tissue samples. We have observed that the best results are obtained when the gastrointestinal tract is cut open longitudinally. The Precellys® tubes allow for a maximum volume of ± 1.5 mL, corresponding to a maximum tissue weight of 75 mg.
Our protocol expresses the obtained MPO activity in units per gram of tissue. Alternatively, one can express the MPO activity in units per gram of tissue protein content. To this end, total protein content needs to be assessed on the tissue supernatant obtained in section B. Commercially available kits, such as the Pierce BCA protein assay (Thermo Fisher, catalog number: 23227) can be used for this measurement.
Sections B (tissue homogenization) and C (myeloperoxidase assay) can be performed on separate days. To do so, collect the tissue supernatant in cryovials and store these at -80 °C until the MPO activity is to be determined.
In our hands, intra-assay variability for an assay performed in duplo was 8.89% for DSS-exposed animals and 75.51% for controls. Note that the high coefficient of variability in control animals could be attributed to the very low read-out values in this group.
Acknowledgments
This work was supported by a BOF-TOP grant of the University of Antwerp (grant number 35018). The quantification of MPO activity in gastrointestinal tissue supernatant was adapted from the work of Krawisz and colleagues (Krawisz et al., 1984).
The authors would like to thank the lab technicians of the Laboratory of Experimental Medicine and Paediatrics for their help with experiments and for critically reviewing the protocol. The graphical abstract was constructed using BioRender.
Competing interests
The authors declare that there are no conflicts of interest.
Ethical considerations
Animal experiments were performed after obtaining approval of the Ethical Committee for Animal Testing of the University of Antwerp (ECD-file 2021-88).
References
Breugelmans, T., Van Spaendonk, H., De Man, J. G., De Schepper, H. U., Jauregui-Amezaga, A., Macken, E., Lindén, S. K., Pintelon, I., Timmermans, J. P., De Winter, B. Y., et al. (2020). In-depth study of transmembrane mucins in association with intestinal barrier dysfunction during the course of T cell transfer and DSS-induced colitis. J Crohns Colitis 14(7): 974-994.
Erben, U., Loddenkemper, C., Doerfel, K., Spieckermann, S., Haller, D., Heimesaat, M. M., Zeitz, M., Siegmund, B. and Kühl, A. A. (2014). A guide to histomorphological evaluation of intestinal inflammation in mouse models. Int J Clin Exp Pathol 7(8): 4557-4576.
Heylen, M., Deleye, S., De Man, J. G., Ruyssers, N. E., Vermeulen, W., Stroobants, S., Pelckmans, P. A., Moreels, T. G. and De Winter, B. Y. (2013). Colonoscopy and μPET/CT are valid techniques to monitor inflammation in the adoptive transfer colitis model in mice. Inflamm Bowel Dis 19(5): 967-976.
Khan, S. S., Smith, M. S., Reda, D., Suffredini, A. F. and McCoy, J. P. Jr. (2004). Multiplex bead array assays for detection of solubile cytokines: comparisons of sensitivity and quantitative values among kits from multiple manufacturers. Cytometry B Clin Cytom 61(1): 35-39.
Kim, J. J., Shajib, S., Manocha, M. M. and Khan, W. I. (2012). Investigating intestinal inflammation in DSS-induced model of IBD. J Vis Exp (60): 3678.
Kuebler, W. M., Abels, C., Schuerer, L. and Goetz, A. E. (1996). Measurement of neutrophil content in brain and lung tissue by a modified myeloperoxidase assay. Int J Microcirc Clin Exp 16(2): 89-97.
Krawisz, J. E., Sharon P. and Stenson W. F. (1984). Quantitative assay for acute intestinal inflammation based on myeloperoxidase activity: Assessment of inflammation in rat and hamster models. Gastroenterology 87(6): 1344-1350.
Mizoguchi, A. (2012). Animal models of inflammatory bowel disease. Prog Mol Biol Transl Sci 105(1): 263-320.
Moreels, T. G., De Man, J. G., De Winter, B. Y., Herman, A. G. and Pelckmans, P. A. (2001). How to express pharmacological contractions of the inflamed rat intestine. Naunyn Schmiedebergs Arch Pharmacol 364(6): 524-533.
Moreels, T. G., Nieuwendijk R. J., De Man J. G., De Winter B. Y., Herman A. G., Van Marck E. A. and Pelckmans P. A. (2004). Concurrent infection with Schistosoma mansoni attenuates inflammation induced changes in colonic morphology, cytokine levels, and smooth muscle contractility of trinitrobenzene sulphonic acid induced colitis in rats. Gut 53(1): 99-107.
Ormrod, D. J., Harrison, G. L. and Miller, T. E. (1987). Inhibition of neutrophil myeloperoxidase activity by selected tissues. J Pharmacol Methods 18(2): 137-142.
Ruyssers, N. E., De Winter, B. Y., De Man, J. G., Loukas, A., Pearson, M. S., Weinstock, J. V., Van den Bossche, R. M., Martinet, W., Pelckmans, P. A. and Moreels, T. G. (2009). Therapeutic potential of helminth soluble proteins in TNBS-induced colitis in mice. Inflamm Bowel Dis 15(4): 491-500.
Solomon, L., Mansor, S., Mallon, P., Donnelly, E., Hoper, M., Loughrey, M., Kirk, S. and Gardiner, K. (2010). The dextran sulphate sodium (DSS) model of colitis: an overview. Comp Clin Pathol 19(1): 235-239.
Schultz, J. and Kaminker, K. (1962). Myeloperoxidase of the leucocyte of normal human blood. I. Content and localization. Arch Biochem Biophys 96: 465-467.
Vermeulen, W., De Man, J. G., Nullens, S., Pelckmans, P. A., De Winter, B. Y. and Moreels, T. G. (2011). The use of colonoscopy to follow inflammatory time course of TNBS colitis in rats. Acta Gastroenterol Belg 74(2): 304-311.
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4,759 | https://bio-protocol.org/en/bpdetail?id=4759&type=0 | # Bio-Protocol Content
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In vivo Electroporation of Skeletal Muscle Fibers in Mice
SF Steven J. Foltz
HH H. Criss Hartzell
HC Hyojung J. Choo
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4759 Views: 499
Reviewed by: Salma Merchant Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Jan 2021
Abstract
In vitro models are essential for investigating the molecular, biochemical, and cell-biological aspects of skeletal muscle. Still, models that utilize cell lines or embryonic cells do not fully recapitulate mature muscle fibers in vivo. Protein function is best studied in mature differentiated tissue, where biological context is maintained, but this is often difficult when reliable detection reagents, such as antibodies, are not commercially available. Exogenous expression of tagged proteins in vivo solves some of these problems, but this approach can be technically challenging because either a mouse must be engineered for each protein of interest or viral vectors are required for adequate levels of expression. While viral vectors can infect target cells following local administration, they carry the risk of genome integration that may interfere with downstream analyses. Plasmids are another accessible expression system, but they require ancillary means of cell penetration; electroporation is a simple physical method for this purpose that requires minimal training or specialized equipment. Here, we describe a method for in vivo plasmid expression in a foot muscle following electroporation.
Graphical overview
Keywords: Skeletal muscle In vivo transfection Muscular dystrophy Muscle biology Imaging
Background
Skeletal muscle is the largest human organ by mass and is essential for autonomous locomotion (Janssen et al., 2000). It is also the target of several degenerative genetic diseases collectively termed muscular dystrophies and myopathies. Recent improvements in genome sequencing have rapidly expanded the list of genes and gene variants causing muscle diseases, many of which have unknown or unconfirmed functions (Lek and MacArthur, 2014; Biancalana and Laporte, 2015; Angelini et al., 2018; Fichna et al., 2018). Besides the obvious desire to understand these novel disease-causing genes for therapeutic development, an exploration of their expression patterns and function, including cellular trafficking and localization, binding partners, and responses to various environmental stressors, is required to expand the current understanding of muscle cell biology. In vitro systems are convenient for the study of new gene products, as they consist of a single or defined mixture of cell types, are usually amenable to genetic manipulation (both knockout and overexpression), and deliver results quickly relative to in vivo systems. However, mature skeletal muscle fibers differ substantially from myotubes cultured in vitro in terms of size, structure, and three-dimensional organization (Dessauge et al., 2021). For example, nuclei regularly cluster in cultured myotubes but are separated into myonuclear domains in vivo. Furthermore, muscle in vivo consists primarily of contractile units (sarcomeres), which are interwoven with highly organized mitochondrial, t-tubule, and sarcoplasmic reticular systems, but typical muscle cell culture systems do not develop well-organized sarcomeres (Denes et al., 2019). These factors combine to influence expression, localization, and behavior of proteins in skeletal muscle fibers, which can differ substantively from what is observed in vitro (Deshmukh et al., 2015).
Historically, animal models (knockout or transgenic) have been the gold standard for probing protein function in vivo but are associated with substantial production costs and long characterization times. Another disadvantage of engineered mice is that a separate mouse is required for each protein of interest, and studying the behavior of several interacting proteins becomes daunting. One would like to use common in vitro tools, like fluorescent and biochemical tags, in vivo. Labeled proteins can be expressed via vectors in skeletal muscle, provided that the vector is able to enter the cell. Adeno-associated viral vectors can freely transduce muscle following intramuscular injection but are labor intensive to produce (Gregorevic et al., 2004; Qiao et al., 2011). Other viral vectors (e.g., lentivirus or retrovirus) can transduce stem cells in vitro, which can then be transplanted into muscle, but this is technically impractical in most cases (Li et al., 2005; Ousterout et al., 2015). A simpler approach is to transfect cells in vivo with plasmid DNA. This is efficiently accomplished through electroporation, a physical method for reversibly permeabilizing the cell membrane with strong electric field. Here, we describe a method for expressing tagged proteins in skeletal muscle fibers of the flexor digitorum brevis (FDB) muscle of the foot through direct intramuscular injection of plasmid DNA followed by electroporation. While this method is theoretically applicable to most skeletal muscles, the FDB possesses several advantages: it is small, accessible, and easily dissociable to generate a suspension of single muscle fibers for downstream analyses. This protocol provides a simple system for evaluating protein functions in skeletal muscle in vivo (Demonbreun and McNally, 2015; Foltz et al., 2021) with supplies that are attainable for most research labs.
Materials and reagents
Standard laboratory bulk materials
Microfuge tubes
Pipette tips
1 mL syringe
26 G needles
0.2 μm filters
Dissecting board and pins
Dissection scissors and forceps
Additional materials and reagents
Hyaluronidase (Sigma-Aldrich, catalog number: H4272); make a 0.5 mg/mL stock in sterile H2O, aliquot, and store at -20 °C
Isoflurane: must be stored in a locked drawer and accessible only by authorized users
Hank’s balanced salt solution (HBSS) (Thermo Fisher, Gibco, catalog number: 14025092)
Collagenase A (Roche, catalog number: 10103586001); prepare a 1% (w/v) stock solution in sterile DMEM, aliquot, and store for < 6 months at -20 °C
1 M HEPES (VWR, catalog number: 97064-360); store at room temperature
DMEM (Thermo Fisher, catalog number: 11995); store at 4 °C
Bovine serum albumin (BSA) (fraction V) (Roche, catalog number: 10775835001); store at 4 °C
CaCl2·2H2O (Sigma-Aldrich, catalog number: C5080); store at room temperature
MgSO4·7H2O (Sigma-Aldrich, catalog number: M5921); store at room temperature
KCl (Sigma-Aldrich, catalog number: P8041); store at room temperature
NaHCO3 (Fisher Scientific, catalog number: S233-500); store at room temperature
NaCl (Fisher Scientific, catalog number: BP358); store at room temperature
NaH2PO4·H2O (Sigma, catalog number: 9763); store at room temperature
100 mM sodium pyruvate (Thermo Fisher, catalog number: 11360070); store at 4 °C
D-Glucose (Sigma-Aldrich, catalog number: G7528); store at room temperature
MEM amino acids (Thermo Fisher, catalog number: 11130051); store at 4 °C
L-serine (Sigma, catalog number: S4311); store at room temperature
Glycine (Sigma, catalog number: G4392); store at room temperature
L-glutamine (Thermo Fisher, catalog number: 25030081); aliquot and store at -20 °C
Penicillin/streptomycin (Thermo Fisher, catalog number: 15140122); aliquot and store at -20 °C
Digestion buffer (see Recipes)
Imaging medium (optional) (see Recipes)
Equipment
Electroporator (BTX, model: ECM 830), Tweezertrodes platinum plated 7 mm (BTX, catalog number: 45-0488), or needle electrodes (2-needle array electrode 5 mm; BTX, catalog number: 45-0206)
Isoflurane flow regulator (Pro 5 oxygen concentrator 625), induction chamber, and nose cone
Cell culture incubator or bead bath at 37 °C (e.g., Fisher Scientific, model: Isotemp 210)
Procedure
Inject plasmids and electroporate FDB
Prepare a solution of 0.5 mg/mL hyaluronidase in sterile deionized H2O (< 18 MOhm). Aliquot and store at ≤ -20 °C.
Note: It is best to prepare ~100 μL per mouse, to allow 20 μL injection into each footpad and dead volume in the syringe needle.
Anesthetize the mouse in an induction chamber with 3% isoflurane and 0.8 L/min of O2. Once the mouse has stopped moving and breathing has slowed, depth of anesthesia should be confirmed through toe-pinch reflex.
Note: We have performed this procedure on mice ranging from three to six months of age, but generally 16–20 weeks; even so, the procedure is expected to be appropriate for mice outside this range.
During the induction period, load a syringe (1 mL or less) with hyaluronidase solution from step A1. Inject 20 μL into each footpad.
Transfer the mouse from the induction chamber to a nose cone maintaining flow of 1.5% isoflurane. The mouse should lay belly down with legs extended away from its head. Clean the skin of the foot with isopropanol.
The FDB is a long, thin muscle spanning the length of the mouse foot. It is the most superficial of the foot muscles in the mouse. Hyaluronidase should be injected beneath the skin but above this muscle. There is an excess of skin at the heel of the mouse foot that can guide injection into the footbed. The syringe needle can be pressed upward against the excess skin as the needle is inserted down the length of the foot, to ensure that this is achieved. Insert the needle to a point just above the branching of the foot into toes. Eject 20 μL of hyaluronidase solution while slowly pulling the needle out. This helps to deliver the solution evenly throughout the footbed.
Remove the mouse from isoflurane and observe its recovery. Mice should be allowed to recover for several minutes on a heated pad. Once the mouse is awake and alert, return it to its home cage. Allow 2 h for hyaluronic acid to digest extracellular matrix before injecting plasmids.
Prepare plasmids for injection. Injection of 20 μg per foot is standard, although more or less may be desirable (this must be determined empirically). Injection volumes should range between 10 and 20 μL. Therefore, dilute plasmid to a concentration between 1 and 2 μg/mL (if necessary) in sterile water.
Note: Overexpression of some proteins can be toxic, which manifests experimentally as relatively few transfected fibers.
Repeat steps A1–A4 to anesthetize the mouse and prepare for plasmid injection to the FDB. Additionally, while anesthesia is induced, prepare the electroporator, connect electrodes, adjust settings, and sterilize electrode tips.
Inject plasmid into one foot, as in step A5. Immediately position the needle tips of the electrode at opposite ends of the foot (one near the heel and one near the toes). Electroporate the muscle with 20 pulses of 150 V and 20 ms duration separated by 980 ms. The foot should twitch with each pulse. If the voltage is too high, the mouse may convulse throughout its body; if the voltage is too low, the foot will not twitch, and transfection will be poor.
Note: These settings are guidelines that have worked well in our hands but may require adjusting. Transduction efficiency will depend on accuracy of injections and electrode placement as well as the strength of electroporation used.
Allow the mouse to recover as in step A6. Robust expression of plasmids is typically seen within 7–10 days after the procedure.
Dissect FDB and isolate individual muscle fibers
Prepare stocks of 1% collagenase in sterile DMEM (high glucose, with L-glutamine and sodium pyruvate).
Prepare digestion buffer (Recipe 1) and warm to 37 °C until FDB muscles are dissected. Prepare 1 mL of digestion buffer per mouse. Both FDB muscles can be digested in a single microfuge tube with 1 mL of digestion buffer, or individual muscles can be digested in 500 μL each.
Euthanize the mouse with an inhalation overdose of isoflurane and perform a secondary means of euthanasia (e.g., cervical dislocation). Pin the foot to a dissecting board just above the branching of the digits using a dissection pin.
Find the excess skin at the heel of the foot and grasp with forceps. Using dissecting scissors, clip the excess skin. This creates a small hole in the skin at the heel of the foot. Using forceps, gently pull up on the skin at this incision. The skin should be a thin layer, distinct from the muscle underneath. Holding the skin away from the muscle with forceps will help to avoid cutting into and damaging the muscle in the next step.
Using dissecting scissors, cut through the skin on either side of the foot from heel to toe. Leave the skin attached at the toes. The skin should now form a triangle, point detached at the heel, and base attached at the toes. From the tip, peel the skin back to expose the underlying foot muscles.
The FDB tendon is found at the base of the foot near the heel. Grasp the tendon with forceps and cut such that the forceps are still holding the tendon after it is released from the bone. The FDB is the long, superficial muscle directly beneath the skin. It can be released from underlying muscles by pulling up on the tendon with forceps and clipping at the sides along the length of the muscle. It terminates where the toes branch. Cut the tendon to fully release the muscle from the foot. OPTIONAL STOPPING POINT: If individual muscle fibers are not required for downstream analysis, the following steps are not necessary. Transfection efficiency is generally low, so detection of the target protein may require a biochemical tag with strong detection reagents. If the target protein is fluorescently tagged, the following steps are recommended.
Note: See Video 1 for a demonstration of the dissection.
Video 1. Dissection of mouse flexor digitorum brevis muscle
Transfer the whole muscle to a Petri dish containing ~20 mL of sterile HBSS. With forceps, briefly rinse the muscle by moving it through the solution to remove any blood, hair, or other contaminants. Move rinsed FDB muscles to digestion buffer. Incubate at 37 °C for 3 h.
Prepare 10% BSA in DMEM (digestion stop solution, 1 mL per FDB). Warm to 37 °C.
Note: Fetal bovine serum (FBS) can replace BSA, depending on the application.
Transfer FDB muscle from digestion buffer to 1 mL of digestion stop solution. Using a wide-bore P200 pipette tip (cut the narrow tip of a normal P200 tip to make a tip diameter of approximately 2 mm), dissociate the fibers with smooth pipetting, 10 times. Repeat this process twice more, using consecutively narrower pipette tip bores. Finally, using a standard P200 pipette tip, pipette the solution 40 times to fully release individual muscle fibers.
Allow the contents of the tube to settle for 2 min and then remove the supernatant containing individual muscle fibers.
Allow the supernatant to settle for an additional 15 min. The muscle fibers will settle, while small debris, dead cells, and single cells from the tissue will remain floating. Remove and discard the supernatant. Resuspend the individual muscle fibers in a buffer suitable for the desired analysis. For any experiment involving visualization of a fluorescent protein, we recommend modified DMEM imaging buffer (Recipe 2) or commercially available phenol-free DMEM, because phenol red and several vitamins contained in standard cell culture media fluoresce when excited with ~488 nm light.
Data analysis
This method can be used to set up a variety of downstream analyses, including molecular biology and biochemistry, which should be adapted to the preferred protocols for individual labs. We find that the procedure described above has particular utility in live-cell imaging applications. Single muscle fibers from step B11 can be resuspended in a small volume (< 100 μL) of imaging buffer and seeded directly onto a Matrigel-coated glass-bottom culture dish. The low plating volume facilitates adherence within approximately 30 min, at which point additional imaging buffer should be applied to the dish. We provide some examples of experiments that can be done on individual muscle fibers, but these are only a small sampling of what is possible. Detailed instructions for analysis can be found in the methods and Figure S5 of Foltz et al. (2021).
Muscle fibers can be electroporated with plasmids to modulate gene expression (e.g., short-hairpin or micro-RNAs) and a reporter gene (in this case, green fluorescent protein), and then analyzed for phenotype seven days later. In this example, a small molecule sensor for phosphatidylserine (fusion protein of lactadherin C2-domain and mCherry fluorescent protein) was included in imaging medium, the muscle fiber membrane was damaged by 405 nm light, and accumulation of the sensor was observed at the damage site with timelapse confocal microscopy. Images are shown from approximately 7 min post-laser injury. In this example, the shRNA did not influence the localization of the phosphatidylserine probe after injury (Figure 1). Note that the ability of a given plasmid to modulate gene expression should be validated in vitro before in vivo use.
Figure 1. Exposure of intracellular membrane lipids after membrane injury
Amphyphysin-2 (BIN1) is a membrane-remodeling protein with a putative role in transverse-tubule biogenesis. Expression of BIN1-mCherry shows expression apparently in t-tubules (Figure 2A), which is in agreement with subcellular localization reported (Prokic et al., 2020). Seven minutes after damage to the sarcolemma, BIN1 becomes enriched at the plasma membrane near the site of injury (white arrowheads, Figure 2B).
Figure 2. Protein localization in vivo
Notes
We have injected up to two plasmids per FDB in mice, for total expression of four plasmids per animal. When two plasmids were injected into a single FDB, the plasmids were pre-mixed and injected simultaneously. We have observed variability in terms of the efficiency of expression. The likeliest causes are imprecise injections of the plasmid or placement of the electroporation electrodes. However, there is a strong possibility that it is also dependent on the gene expressed. Overexpression of some genes may be toxic. In rare cases, few to no fibers may be expressing the plasmid. Users should keep note of transfection efficiency with all plasmids to track emergent patterns. Changes in the expression vector, for example the promoter, can reduce the level of gene expression and mitigate potential toxicity.
Successful electroporation requires the use of a strong electrical field. It poses a high risk of pain to the mouse and should only be performed on animals maintained under deep anesthesia. Every care should be taken to minimize distress to the animals.
Recipes
Digestion buffer
Reagent Final concentration Amount
Collagenase A 0.2% 200 μL
HEPES (1 M) 25 mM 25 μL
DMEM n/a 775 μL
Total n/a 1 mL
Imaging medium (optional)
Reagent Final concentration Amount
CaCl2·2H2O 1.8 mM 26.4 mg
MgSO4·7H2O 0.8 mM 197.2 mg
KCl 5.3 mM 39.5 mg
NaHCO3 44 mM 369.6 mg
NaCl 110 mM 642.8 mg
NaH2PO4·H2O 0.9 mM 127.7 mg
Sodium pyruvate 1 mM 1 mL
D-Glucose 5.3 mM 100 mg
MEM amino acids 2× 4 mL
L-serine 0.4 mM 42 mg
Glycine 0.4 mM 30 mg
L-glutamine 4 mM 2 mL
Penicillin/streptomycin 100 U/L 1 mL
H2O 92 mL
Total 100 mL
Acknowledgments
This protocol is derived from the original research paper (Foltz et al., 2021; DOI: 10.1083/jcb.202007059). This work was supported by grants from the National Institutes of Health: the National Institute of Arthritis and Musculoskeletal and Skin Diseases awards R01AR067786 (to H. Criss Hartzell), R01AR071397 (to H.J. Choo), and F32AR074249 (to S.J. Foltz); National Eye Institute award R01EY114852 (to H. Criss Hartzell); and National Institute of General Medical Sciences grant R01GM132598 (to H. Criss Hartzell). Figure prepared in BioRender.com.
Competing interests
The authors declare no competing interests.
Ethics
All procedures involving animals were approved by the Emory Institutional Care and Use Committee under protocols 201800130 (Hartzell) and 201700233 (Choo).
References
Angelini, C., Giaretta, L. and Marozzo, R. (2018). An update on diagnostic options and considerations in limb-girdle dystrophies. Expert Rev Neurother 18(9): 693-703.
Biancalana, V. and Laporte, J. (2015). Diagnostic use of Massively Parallel Sequencing in Neuromuscular Diseases: Towards an Integrated Diagnosis. J Neuromuscul Dis 2(3): 193-203.
Demonbreun, A. R. and McNally, E. M.(2015). DNAElectroporation, Isolation and Imaging of Myofibers. J Vis Exp (106): e53551.
Denes, L. T., Riley, L. A., Mijares,J. R., Arboleda, J. D., McKee, K., Esser, K. A. and Wang, E. T. (2019). Culturing C2C12 myotubes on micromolded gelatin hydrogelsaccelerates myotube maturation. Skelet Muscle 9(1): 17.
Deshmukh, A. S., Murgia, M., Nagaraj,N., Treebak, J. T., Cox, J. and Mann, M. (2015). Deep proteomics of mouse skeletal muscle enables quantitation of protein isoforms,metabolic pathways, and transcription factors. Mol Cell Proteomics 14(4): 841-853.
Dessauge, F., Schleder, C., Perruchot,M. H. and Rouger, K. (2021). 3Din vitro models of skeletal muscle: myopshere, myobundle and bioprinted muscle construct. Vet Res 52(1): 72.
Fichna, J. P., Macias, A., Piechota, M., Korostynski, M., Potulska-Chromik, A., Redowicz, M. J. and Zekanowski, C. (2018). Whole-exome sequencing identifies novel pathogenic mutations and putative phenotype-influencing variants in Polish limb-girdle muscular dystrophy patients. Hum Genomics 12(1): 34.
Foltz, S. J., Cui, Y. Y., Choo, H. J. and Hartzell, H. C. (2021). ANO5 ensures trafficking of annexins in wounded myofibers. J Cell Biol 220(3): e202007059.
Gregorevic, P., Blankinship, M. J., Allen, J. M., Crawford, R. W., Meuse, L., Miller, D. G., Russell, D. W. andChamberlain, J. S. (2004). Systemic delivery of genes to striated muscles using adeno-associated viral vectors. Nat Med 10(8): 828-834.
Janssen, I., Heymsfield, S. B., Wang, Z. M. and Ross, R. (2000). Skeletal muscle mass and distribution in 468 men and women aged 18-88 yr. J Appl Physiol (1985) 89(1): 81-88.
Lek, M. and MacArthur, D. (2014). The Challenge of Next Generation Sequencing in the Context of Neuromuscular Diseases. J Neuromuscul Dis 1(2): 135-149.
Li, S., Kimura, E., Fall, B. M., Reyes, M., Angello, J. C., Welikson, R., Hauschka, S. D. and Chamberlain, J. S.
(2005). Stable transduction of myogenic cells with lentiviral vectors expressing a minidystrophin. Gene Ther 12(14): 1099-1108.
Ousterout, D. G., Kabadi, A. M., Thakore, P. I., Majoros, W. H., Reddy, T. E. and Gersbach, C. A. (2015). Multiplex CRISPR/Cas9-based genome editing for correction of dystrophin mutations that cause Duchenne muscular dystrophy. Nat Commun 6: 6244.
Prokic, I., Cowling, B. S., Kutchukian, C., Kretz, C., Tasfaout, H., Gache, V., Hergueux, J., Wendling, O., Ferry, A., Toussaint, A., et al. (2020). Differential physiological roles for BIN1 isoforms in skeletal muscle development, function and regeneration. Dis Model Mech 13(11): dmm044354.
Qiao, C., Koo, T., Li, J., Xiao, X. and Dickson, J. G. (2011). Gene therapy in skeletal muscle mediated by adeno-associated virus vectors. Methods Mol Biol 807: 119-140.
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4,760 | https://bio-protocol.org/en/bpdetail?id=4760&type=0 | # Bio-Protocol Content
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Peer-reviewed
Inoculation of Maize with Sugarcane Mosaic Virus Constructs and Application for RNA Interference in Fall Armyworms
IG Iram Gull
GJ Georg Jander
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4760 Views: 570
Reviewed by: Zhibing LaiDemosthenis ChronisMalgorzata LichockaJuliane K Ishida
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Original Research Article:
The authors used this protocol in Plant Biotechnology Journal Sep 2021
Abstract
Virus-mediated transient gene overexpression and gene expression silencing can be used to screen gene functions in plants. Sugarcane mosaic virus (SCMV) is a positive strand RNA virus in the Potyviridae family that has been modified to be used as vector to infect monocots, including maize (Zea mays), for transient gene overexpression and gene expression silencing. Relative to stable transformation, SCMV-mediated transient expression in maize has the advantages of being faster and less expensive. Here, we describe a protocol for cloning constructs into the plasmid vector pSCMV-CS3. After maize seedlings are transformed with pSCMV-CS3 constructs by particle bombardment, the virus replicates and spreads systemically in the plants. Subsequent infections of maize seedlings can be accomplished by rub inoculation with sap from SCMV-infested plants. As an example of a practical application of the method, we also describe virus-induced gene silencing (VIGS) of fall armyworm (Spodoptera frugiperda) gene expression. Transgenic viruses are created by cloning a segment of the fall armyworm target gene into pSCMV-CS3 prior to maize transformation. Caterpillars are fed on the virus-infected maize plants, which make dsRNA to silence the expression of the fall armyworm target gene after ingestion. This use of SCMV for plant-mediated VIGS in insects allows rapid screening of gene functions when caterpillars are feeding on their host plants.
Graphical overview
Keywords: Sugarcane mosaic virus Biolistic transformation Zea mays Spodoptera frugiperda VIGS
Background
Viruses can be engineered as vectors for overexpression of heterologous coding or non-coding RNA sequences in plants. Additionally, viruses carrying a fragment of a target gene can cause virus-induced gene silencing (VIGS) by activating the internal RNA interference (RNAi) machinery of their host plants (Pasin et al., 2019). The rapid replication of plant viruses makes VIGS an efficient approach for in vivo investigation of gene function in plants.
Several virus vectors have been used for VIGS in monocot plant species, including Chinese wheat mosaic virus (Yang et al., 2018), tobacco rattle virus (TRV) (Zhong et al., 2014), barley stripe mosaic virus (Buhrow et al., 2016), brome mosaic virus (Wang et al., 2021), bamboo mosaic virus, cymbidium mosaic virus (Hsieh et al., 2013), rice tungro bacilliform virus (Purkayastha et al., 2010), cucumber mosaic virus (Tzean et al., 2019), foxtail mosaic virus (Liu et al., 2016), maize rayado fino virus (Mlotshwa et al., 2020), and sugarcane mosaic virus (SCMV) (Mei et al., 2019; Chung et al., 2022). Several of these viruses, including SCMV, have been used for VIGS in maize (Mei et al., 2016; Wang et al., 2016; J. Zhang et al., 2017; Ding et al., 2018; Jarugula et al., 2018; Mlotshwa et al., 2020; Chung et al., 2022)
SCMV is a positive strand RNA virus of the Potyviridae family that infects many monocots including sugarcane, maize, wheat, and sorghum (Xiao et al., 1993). Previously, SCMV has been used as a vector for transient overexpression of non-maize genes (Mei et al., 2019), overexpression of maize endogenous genes (Chung et al., 2021), VIGS of maize endogenous genes (Chung et al., 2022), and plant-mediated VIGS of gene expression in corn leaf aphids (Rhopalosiphum maidis) feeding on maize (Chung and Jander, 2022). The plasmid vector pSCMV-CS3 (Mei et al., 2019; Chung et al., 2022) allows for both efficient gene overexpression and gene expression silencing by RNA interference. The entire SCMV virus is encoded between a cauliflower mosaic virus 35S promoter and a nopaline synthase terminator in the Escherichia coli plasmid pSCMV-CS3. A multiple cloning site, which allows cutting by the enzymes PspOMI, ApaI, PmeI, PstII, and SbfI, is inserted between the P1 and HC-Pro protein-coding regions of the virus. Since the virus is translated as a single polyprotein, prior to being cleaved by virus-encoded proteases, it is essential that there is no in-frame stop codon in the sequence that is cloned into pSCMV-CS3. A NIa protease cleavage site immediately downstream of the multiple cloning site catalyzes the release of the cloned protein from the endogenous viral proteins. Biolistic transformation of pSCMV-CS3 into plant cells causes the virus to be transcribed from the 35S promotor, thereby initiating a systemic viral infection.
The fall armyworm (Spodoptera frugiperda), a lepidopteran insect in the Noctuidae family, is one of the most important pests of maize, causing considerable damage to this important crop worldwide. It is a native North American species that has expanded its range to Africa and Asia in recent years (Rwomushana, 2019). The larval stage preferably consumes the tender shoots and leaves of maize plants (He et al., 2020). Currently available strategies such as use of chemical insecticides (Sisay et al., 2019) and transgenic maize producing Bt (Bacillus thuringiensis) toxin (Niu et al., 2016) are becoming less effective due to resistance that is developing in the pests (Banerjee et al., 2017; Zhang et al., 2020) However, RNAi has emerged as a promising approach for agricultural pest control (Zotti et al., 2018). RNAi of insect genes can be achieved by feeding them on host plants with either stable or transient expression of double-stranded RNA (dsRNA) targeting specific inset genes.
Plant-mediated RNAi has been used to silence expression of lepidopteran genes, including silencing of tobacco hornworm (Manduca sexta) genes by stable (Poreddy et al., 2017) or transient expression of dsRNA in tobacco using TRV (Kumar et al., 2012), and stable expression of dsRNA of cotton bollworm (Helicoverpa armigera) genes in Arabidopsis thaliana (Chen et al., 2019), Nicotiana benthamiana (Bally et al., 2020), cotton (Gossypium hirsutum) (Tian et al., 2015; Mao et al., 2007), and tomato (Lycopersicum esculentum) (Mamta et al., 2016). Most of these studies used expression of RNA in stable transgenic plants, an approach that is a laborious, relatively expensive, and low-throughput strategy for studying gene function in plant pests. By contrast, using viruses as vectors for transient expression of dsRNA is a robust method for generating a rapid and systemic RNAi signal in plants.
Here, we describe a general protocol for infecting maize with pSCMV-CS3 gene constructs, as well as the specific use of SCMV-mediated VIGS for silencing gene expression in fall armyworm caterpillars that are feeding on infected maize plants. When the maize RNAi pathway is activated by SCMV infection, viral dsRNA is cleaved into short interfering RNA (siRNA) of 21–24 nucleotides in length (Ding and Voinnet, 2007). Upon feeding on SCMV-infected maize, fall armyworm caterpillars ingest dsRNA and/or siRNA, which silences expression of the target gene.
Materials and reagents
Plants, insects, vectors, and cell lines
Zea mays inbred line P39 seeds (Maize COOP, http://maizecoop.cropsci.uiuc.edu)
Sweet corn variety Golden Bantam (Burpee Seeds, Warminster Township, PA, or www.amazon.com)
Fall armyworm (Spodoptera frugiperda) eggs (Benzon Research, Carlisle, PA, USA, www.benzonresearch.com)
pSCMV-CS3 (plasmid with a SCMV cloning site #3). This plasmid can be obtained with a material transfer agreement from Iowa State University; contact Dr. Steve Whitham ([email protected])
Escherichia coli DH5α competent cells (Thermo Fisher, catalog number: 18265017)
pSCMV-GFP and pSCMV-GOI (Gene of Interest)
Oligonucleotides (Integrated DNA Technologies, USA)
Flanking multiple cloning site of pSCMV-CS3
SP7788: 5′-GCACAAATGGTTTCCAACG-3′
SP7789: 5′-ATGTTGCATGTCTTGCATG-3′
Reference maize gene primers for reverse-transcriptase PCR (RT-PCR)
Actin-F, 5′-GGTTTCGCTGGTGATGATGC-3′
Actin-R, 5′-CAATGCCATGCTCAATCGGG-3′
EF-1α-F, 5′-TGGGCCTACTGGTCTTACTACTGA-3′
EF-1α-R, 5′-ACATACCCACGCTTCAGATCCT-3′
Reference fall armyworm gene primers for quantitative reverse transcriptase-PCR (qPCR)
qrSF-RPL13-F, 5′-GCCTTAACCCTGCTTTTGCTAG-3′
qrSF-RPL13-R, 5′-GCTTCGCCCTTCAATACCTTC-3′
qrSF-EF1α-F, 5′-TGGGCGTCAACAAAATGGA-3′
qrSF-EF1α-R, 5′-TCTCCGTGCCAGCCAGAAAT-3′
Primers for VIGS target gene cloning and quantitative PCR (will vary based on the gene of interest)
Materials
Wizard® SV Gel and PCR clean-up system (Promega, catalog number: A9282)
Wizard® Plus SV Minipreps DNA purification system (Promega, catalog number: A1460)
Wizard® SV total RNA isolation system (Promega, catalog number: Z3100)
High-Capacity cDNA Reverse Transcription kits (Applied Biosystems, catalog number: 4374966)
GoTaq® Green Master Mix (Promega, catalog number: M7123)
PowerUpTM SYBRTM Green Master Mix (Applied Biosystems, catalog number: A25742)
Restriction enzymes: PspOMI, SbfI (NEB Biolabs, catalog number: R0653S and R3642)
T4 DNA ligase (Promega, catalog number: M1804)
100 bp DNA ladder (Promega, catalog number: G210A)
Nuclease-free water
Absolute ethanol (Sigma-Aldrich, catalog number: E7023)
Liquid nitrogen
Carborundum powder (Fisher, catalog number: C192-500)
Mortar and pestle
Pellet pestle (Fisher Scientific, catalog number: 12-141-363)
Biolistic Optimization kit [1.0 μm Gold Microcarriers, Macrocarrier disks, stopping screens, and 7,600 kPa (1,100 psi) rupture disks (Bio-Rad Laboratories, catalog number: 165-2279)]
0.1 M spermidine (Sigma-Aldrich, catalog number: 05292-1ML-F)
Pipettes and pipette tips (various)
Agarose low-EEO/multi-purpose/molecular biology grade (Fisher BioReagentsTM, catalog number BP160-100)
Microperforated plastic vented bread bags (30 cm × 60 cm) (www.amazon.com)
Plant trays without holes (28 cm wide × 54 cm long × 6 cm deep), plastic pots (9 cm square, 8 cm deep) (www.amazon.com)
qPCR plates (MicroAmpTM optical 384-well reaction plate with barcode (Applied Biosystems, catalog number: 43-098-49)
Cornell maize soil mix: [0.16 m3 Metro-Mix 360 (Scotts, Marysville, OH, USA), 0.45 kg finely ground lime, 0.45 kg Peters Unimix (Griffin Greenhouse Supplies, Auburn, NY, USA), 68 kg Turface MVP (Banfield-Baker Corp., Horseheads, NY, USA), 23 kg coarse quartz sand, and 0.018 m3 pasteurized field soil]
Fall armyworm artificial diet (Southland Products Inc, Lake Village, AR, USA, www.southlandproducts.net)
50 mg/mL kanamycin stock solution (Sigma-Aldrich, catalog number: K0254-20ML)
CaCl2·6H2O (Sigma-Aldrich, catalog number: 21108-500G)
Tryptone (Sigma-Aldrich, catalog number: T7293-1KG)
NaCl (Sigma-Aldrich, catalog number: S9888-1G)
Yeast extract (Sigma-Aldrich, catalog number: 70161-500G)
Agar (Sigma-Aldrich, catalog number: A1296-500G)
15 cm Petri dishes (Fisher Scientific, catalog number FB0875714)
0.22 μm syringe filters (Corning®, catalog number: CLS431219-50EA)
Monobasic potassium phosphate (KH2PO4) (Sigma-Aldrich, catalog number: P0662-500G)
Dibasic potassium phosphate, (K2HPO4) (Sigma-Aldrich, catalog number: P3786-500G)
Tris base (Sigma-Aldrich, catalog number: T1503-500G)
Acetic acid (Sigma-Aldrich, catalog number: A6283-1L)
EDTA (Sigma-Aldrich, catalog number: E4884-500G)
Glycerol (Sigma-Aldrich, catalog number: G7893-IL)
Buffers and solutions
2.5 M CaCl2 (see Recipes)
LB medium (see Recipes)
Inoculation buffer (see Recipes)
1× TAE buffer (see Recipes)
50% (v/v) glycerol (see Recipes)
70% ethanol (see Recipes)
LB agar plates (see Recipes)
Recipes
2.5 M CaCl2
Dissolve 11.0 g of CaCl2·6H2O in deionized water and make the volume up to 20 mL, filter sterilize with a 0.22 μm filter, and store at 4 °C.
LB medium
1% (w/v) tryptone
1% (w/v) NaCl
0.5% (w/v) yeast extract
pH 7.5
Inoculation buffer (50 mM potassium phosphate)
43.4 mL of 1 M KH2PO4
6.6 mL of 1 M K2HPO4
Bring volume to 1 L with deionized water
pH 6.0
1× Tris-Acetate-EDTA (TAE) buffer
242 g of Tris base
57.1 mL of glacial acetic acid
18.6 g of EDTA
Bring final volume to 1 L with deionized water
This is a 50 × TAE stock solution, dilute 1:50 to make 1× TAE buffer
50% (v/v) glycerol
50 mL of 100% glycerol in 50 mL of deionized water
70% ethanol
70 mL of absolute ethanol in 30 mL of deionized water
LB agar plates
LB medium (see Recipe 2)
1.5% w/v agar
Boil to dissolve the agar
Pour approximately 25 mL of heated LB agar into each Petri dish
Equipment
NanoDrop ND-1000 spectrophotometer (Thermo Fisher, USA)
PDS-1000/He biolistic particle delivery system with metal mesh (Bio-Rad laboratories, USA)
1600 MiniG® automated tissue homogenizer and cell lyser (SPEX® Sample Prep, USA)
QuantStudio 6 Flex real-time PCR system (Applied Biosystems, USA)
Ultrasonic water bath with 40 kHz frequency and 284 W heater power (FS140H, Fisher Scientific, USA)
Plate centrifuge (Benchmark Scientific, USA)
Vortex (American Scientific Products, USA)
Benchtop centrifuge (Thermo Scientific, USA)
Incubator with temperature set at 37 °C
Incubator with temperature set at 28 °C
Shaker with temperature set at 37 °C
Plant growth chambers
Procedure
Selection of target sequence and primer design
Select a sequence of size between 200 and 400 bp from the fall armyworm gene of interest, which should be a multiple of three base pairs, avoiding stop codons (in the antisense direction), and avoiding restriction sites of selected restriction enzymes. Check for off targets in the selected sequence against the fall armyworm and maize genomes using blastn at the National Center for Biotechnology Information (NCBI; https://blast.ncbi.nlm.nih.gov). Select a sequence with no off targets.
Design PCR primers to specifically amplify the selected sequence and clone the sequence into the pSCMV-CS3 vector (Figure 1 and Supplementary information) in the antisense direction using the SbfI and PspOMI restriction enzymes.
Figure 1. Plasmid map of pSCMV-CS3, showing the restriction sites used for cloning gene fragments into sugarcane mosaic virus (SCMV)
Following the same parameters, clone a 240 bp fragment of the jellyfish green fluorescent protein (Gfp) gene into the pSCMV-CS3 vector to use as negative control for VIGS experiments.
Synthesis of SCMV-VIGS constructs
Hatch fall armyworm eggs on fall armyworm diet at 28 °C in an incubator. Use 7-day-old fall armyworm caterpillars for RNA isolation. Freeze caterpillars using liquid nitrogen. Grind the frozen larval tissue in a microfuge tube using a pellet pestle and isolate the total RNA using Wizard® SV Total RNA isolation system following the manufacturer’s instructions (Note 1).
Quantify the RNA and use 1 μg for reverse transcription using High-Capacity cDNA Reverse Transcription kits following the manufacturer’s instructions.
Using the cDNA as a template, amplify the fragment of target gene using 400 nM of each target-specific primer with GoTaq® Green Master Mix.
Analyze the specific amplification by 1.5% agarose electrophoresis using 1× TAE buffer, checking the correct size of the fragments by running a 100 bp DNA ladder in a parallel lane on the gel.
Purify the amplicon using Wizard® SV Gel and PCR clean-up system following the manufacturer’s instructions.
Double-digest the purified amplicons and pSCMV-CS3 vector with the selected restriction enzymes and, after analyzing the digestion by 1.5% agarose gel electrophoresis, purify fragments from gel using Wizard® SV Gel and PCR clean-up system following the manufacturer’s instructions.
Quantify the restricted purified products and ligate the target fragments in linearized pSCMV-CS3 using a 1:3 equimolar end ratio and T4 DNA ligase according to the manufacturer’s instructions.
Transform the ligation mixture into E. coli Top 10 competent cells (1:10 dilution) using a heat shock method and select the transformants after overnight growth at 37 °C on LB agar plates containing 50 μg/mL kanamycin.
Verify insertion of the fragment of interest in pSCMV-CS3 by screening a few colonies by PCR using the SP7788/SP7789 primers (flanking the multiple cloning site of the pSCMV-CS3 vector).
Verify the sequence of the inserted fragments by Sanger sequencing with the SP7788 primer, using plasmid DNA isolated using the Wizard® Plus SV Minipreps DNA purification system from positive colonies after overnight growth at 37 °C with shaking at 180 rpm in LB broth containing 50 μg/mL kanamycin.
Biolistic transformation of SCMV-VIGS constructs into maize and in planta detection of RNA expression
Prepare seedlings at the two-leaf stage by sowing five seeds of maize variety Golden Bantam in a pot containing Cornell maize mix. Allow seeds to germinate and grow for a week in a growth chamber with a 16:8 light/dark cycle at 23 °C with 60% humidity (Figure 2) (Notes 2 and 3). Prepare Gold microcarriers (1.0 μm) at the concentration of 100 mg/mL in 50% glycerol. For this, first vortex 50 mg of gold particles in 1 mL of 70% ethanol for 5 min at 2,000 rpm. Then, keep at room temperature for 15 min and collect the gold particles by centrifugation at 900× g for 20 s. Wash the gold particles three times with 1 mL of sterile water by following this sequence: vortex for 1 min at 2,000 rpm, incubation at room temperature for 1 min, and centrifugation for 10 s. Finally, resuspend particles in 500 μL of 50% glycerol (Note 4).
Figure 2. Maize variety Golden Bantam seedlings at the two-leaf stage for biolistic transformation. Five seedlings are planted in each pot.
Place 30 μL of prepared gold microcarriers in a 1.5 mL microfuge tube and vortex for 30 s at 2,000 rpm, followed by sonication for 10 s in an ultrasonic water bath at room temperature with 40 kHz frequency.
Add 5 μg of the pSCMV-VIGS construct during sonication. After 10 s of further sonication, add 25 μL of 2.5 M CaCl2 and 10 μL of 0.1 M spermidine and vortex for 3 min at 2,000 rpm, followed by centrifugation at 2,700× g for 30 s to collect the pellet of gold microcarriers.
Discard supernatant and wash the gold particles with 150 μL of absolute ethanol followed by vortexing at 2,000 rpm for 10 s and centrifugation at 2,700× g for 10 s to collect the gold microcarriers.
Remove the supernatant and resuspend the DNA-coated gold particles in 50 μL of absolute ethanol (see Figure 3 for a flowchart of these steps).
Figure 3. Steps for coating of DNA on gold microcarriers
Place the microfuge tubes having DNA-coated gold microcarriers in an ultrasonic water bath, remove μL of microcarrier suspension, evenly spread it on macrocarrier, and air dry. Prepare four more macrocarriers with the remaining suspension of DNA-coated gold microcarriers.
Arrange the PDS-1000/He biolistic particle delivery system (gene gun) with a microcarrier disk, stopping screen, and 7,600 kPa rupture disk following the manufacturer’s instructions (https://www.bio-rad.com/webroot/web/pdf/lsr/literature/M1652249.pdf). Place one pot of maize seedlings in the instrument chamber and keep the leaves flat against the solid support with a metal mesh that is supplied along with the biolistic particle delivery system (Figure 4). After closing the door, turn on the vacuum and shoot the gold particles when the vacuum reaches 7,600 kPa.
Figure 4. Maize variety Golden Bantam seedlings in the chamber of the PDS-1000/He biolistic particle delivery system
When the helium pressure reaches approximately 7,600 kPa, a clear pop is heard. Release vacuum at this point, remove the pot, mist with water, and transfer the plants to a growth chamber.
Transfer the inoculated plants to individual pots the next day and grow them in a growth chamber under a 16:8 h photoperiod at 23 °C for three weeks, until the appearance of viral symptoms.
After three weeks, collect the tissue from the seventh leaf of each plant and immediately place it in liquid nitrogen.
Grind the leaf tissue using 1600 MiniG® automated tissue homogenizer and cell lyser and extract the total RNA using Wizard® SV Total RNA isolation system following the manufacturer’s instructions.
Quantify the RNA and use 1 μg of RNA for reverse transcription using High-Capacity cDNA Reverse Transcription kit following the manufacturer’s instructions.
Use SP7788 & SP7789 primers for PCR amplification and analyze the expression of target sequence in maize. Maize actin and EF-1α gene are amplified as reference genes using Actin-F, Actin-R & EF-1α-F, and EF-1α-R primer pairs. Analyze the amplicons by 1.5% agarose gel electrophoresis (Note 5).
Preparation of sap and rub inoculation of maize with SCMV-VIGS constructs
Separate the whole seventh leaf from the infected plant in which the expression of dsRNA has been confirmed, grind the tissue finely in a mortar and pestle in the presence of liquid nitrogen, wait for a few minutes until sap comes out of tissue, add inoculation buffer in 1:10 ratio (w/v), and grind again to prepare sap for inoculation (Figure 5A) (Note 6).
Grow maize inbred line P39 plants in individual pots in a growth chamber with a 16:8 h photoperiod at 23 °C for one week until the two-leaf stage (Note 7).
Spray the seedling leaves with deionized water and dust (light sprinkling) the leaves with 600 mesh carborundum.
Take a cotton swab, wet it with prepared maize sap, and use it to rub the top surface of both leaves (Figure 5B) (Note 8).
Figure 5. Sap preparation (A) and rub inoculation (B). Seven-day-old maize seedlings of inbred line P-39 at the two-leaf stage are rub inoculated with sap prepared from plants infected with pSCMV-VIGS constructs.
Transfer the plants to the growth chamber and look for the appearance of viral infection symptoms after three weeks under a 16:8 h photoperiod at 23 °C.
The expression of viral RNA can be confirmed by PCR as described in section C, step 15, as plants without viral symptoms may also have active infections and expression of the gene of interest (Figure 6).
Figure 6. Maize leaves collected from sugarcane mosaic virus (SCMV)-induced gene silencing (VIGS)-infected plants and confirmation of viral infection by PCR. Leaves without (A) and with (B) mosaic symptoms can be positive for SCMV infection. (C) 1.5% agarose gel electrophoresis analysis of PCR for dsRNA from leaves with (lane 1) and without (lane 2) viral symptoms; negative control using water (lane C). (D) 1.5% agarose gel electrophoresis analysis of PCR for maize reference genes actin (lane 1) and EF-1α (lane 2) in the same virus-infected maize leaves.
Caterpillar bioassay for SCMV VIGS in maize and measurement of gene silencing in caterpillars by qRT-PCR
Three weeks after infecting the maize plants with pSCMV-GOI and pSCMV-GFP control constructs by rub inoculation, put five pre-weighed two-day-old caterpillars onto each plant. One caterpillar will be used for measurement of gene silencing after three days of feeding. Growth of the other caterpillars will be monitored after seven days of feeding. Cover the plants with the perforated plastic bags and tie the bags around the stems of the plants with wire twist ties to prevent caterpillar escape (Figure 7).
Figure 7. Bioassays with Spodoptera frugiperda caterpillars. (A) Experimental setup for caterpillar bioassay and gene expression analysis, starting three weeks post rub inoculation. Five two-day-old caterpillars are confined on each plant. The plants are covered with perforated bread bags, which are tied around the plant stem with wire twist ties. (B) Spodoptera exigua (fall armyworm) on a maize leaf.
Transfer the plants back to the growth chamber under the same conditions (16:8 h photoperiod at 23 °C) and monitor the growth of caterpillars after seven days by collecting and weighing the surviving caterpillars.
For measurement of gene silencing, collect one caterpillar from each plant after three days of feeding on pSCMV-VIGS- and pSCMV-GFP-infected plants, place it in a 1.5 mL microfuge tube, and immediately place the tube in liquid nitrogen.
Grind collected caterpillars with a pellet pestle tissue homogenizer and extract the total RNA using Wizard® SV Total RNA isolation system following the manufacturer’s instructions.
Quantify the RNA using a Nanodrop and synthesize the cDNA with 1 μg of RNA using High-Capacity cDNA Reverse Transcription kit following the manufacturer’s instructions.
Perform quantitative PCR using a QuantStudio 6 Flex real-time PCR system, with a five-fold dilution of cDNA samples using qPCR primers and PowerUpTM SYBRTM Green Master Mix. The expression levels are normalized using the fall armyworm RPL13 and EF1α reference genes (with primers qrSF-RPL13-F, qrSF-RPL13-R, qrSF-EF1α-F, and qrSF-EF1α-R) as internal controls.
The gene expression levels are calculated using the 2-ΔΔCt method (Livak and Schmittgen, 2001) (Figure 8). Gene expression levels of caterpillars fed on pSCMV-GOI infected plants is compared to those on control plants (pSCMV-GFP infected plants).
Figure 8. Sample quantitative PCR amplification plots from a QuantStudio 6 Flex real-time PCR system. (A) Amplification of a control housekeeping gene, showing a threshold cycle (CT) value of 19.5. (B) Amplification of a gene of interest from control and experimental (RNA interference) samples, showing CT values of 23.5 and 26, respectively. (C) Formula used to calculate the relative gene expression in the RNAi sample compared to that of the untreated control.
Notes
A similar approach can be used for isolating RNA from other insect species if their gene expression will be targeted by VIGS.
Use pots (9 cm square × 8 cm deep) to grow seedlings. Prepare five pots of seedling per construct transformation. In our hands, Golden Bantam works well for biolistic transformation of maize, but inbred line P39 can also be used.
The efficiency of transformation decreases after the two-leaf stage of seedling. Keep the seedlings in the dark for 24 h before biolistic transformation.
The prepared gold microcarriers can be stored at -20 °C and used for up to three months. Three milligrams of prepared microcarrier is sufficient for five bombardments per construct.
The amplified band is expected at 475 bp + the size of the target gene fragment. You can also confirm the sequence of the target fragment by Sanger sequencing.
Maize leaf sap can be stored indefinitely at -80 °C. Avoid frequent freeze thaw of the sap, as it will reduce the infection efficiency. You can also store the infected leaf tissue (0.25 g of leaf tissue per tube) at -80 °C after snap freezing in liquid nitrogen. This frozen tissue can be used to prepare fresh sap for inoculation of plants. Infection efficiency in both cases will be same.
Use of seedlings after the two-leaf stage results in low infection efficiency.
Change gloves and the cotton swab for sap from different pSCMV-VIGS constructs to avoid cross-contamination.
Acknowledgments
This work was supported by an International Postdoc Scholarship from the Punjab Higher Education Commission to I.G., NSF award 2019516 to G.J., and USDA award 2021-67014- 342237 to G.J.
Competing interests
The authors declare that they have no competing interests.
References
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Supplementary information
The following supporting information can be downloaded here:
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An ex vivo Model of Paired Cultured Hippocampal Neurons for Bi-directionally Studying Synaptic Transmission and Plasticity
RS Ruslan Stanika
GO Gerald J. Obermair
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4761 Views: 720
Reviewed by: Verena BurtscherMohammed Mostafizur RahmanWilly R Carrasquel-Ursulaez
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Original Research Article:
The authors used this protocol in The Journal of Neuroscience Apr 2019
Abstract
Synapses provide the main route of signal transduction within neuronal networks. Many factors regulate critical synaptic functions. These include presynaptic calcium channels, triggering neurotransmitter release, and postsynaptic ionotropic receptors, mediating excitatory and inhibitory postsynaptic potentials. The key features of synaptic transmission and plasticity can be studied in primary cultured hippocampal neurons. Here, we describe a protocol for the preparation and electrophysiological analysis of paired hippocampal neurons. This model system allows the selective genetic manipulation of one neuron in a simple neuronal network formed by only two hippocampal neurons. Bi-directionally analyzing synaptic transmission and short-term synaptic plasticity allows the analysis of both pre- and postsynaptic effects on synaptic transmission. For example, with one single paired network synaptic responses induced by both, a wild-type neuron and a genetically modified neuron can be directly compared. Ultimately, this protocol allows experimental modulation and hence investigation of synaptic mechanisms and thereby improves previously developed methods of studying synaptic transmission and plasticity in ex vivo cultured neurons.
Key features
• Preparation of ex vivo paired cultured hippocampal neurons.
• Bi-directional electrophysiological recordings of synaptic transmission and plasticity.
• Genetic modulation of synaptic network formation (demonstrated by presynaptic viral overexpression of the auxiliary calcium channel α2δ-2 subunit).
Graphical overview
Keywords: Primary hippocampal culture Paired patch clamp recording Synaptic transmission Calcium channel α2δ subunit Viral infection Electrophysiology Short-term plasticity Postsynaptic currents Paired pulse facilitation and depression
Background
Signal transmission between neurons occurs via neurotransmitter release into the synaptic cleft. The temporal and spatial relation of pre- and post-synaptic firing modulates the strength of synaptic connections between neurons (Deperrois and Graupner, 2020). Two types of synaptic activity can be registered at the single cell level. First, miniature excitatory or inhibitory postsynaptic currents (mEPSC or mIPSC, respectively) appear as result of spontaneous local fusion of single synaptic vesicles. Second, excitatory and inhibitory postsynaptic currents (EPSC or IPSC, respectively) can be recorded in response to action potential firing by presynaptic glutamatergic or GABAergic neurons, respectively. Analysis of miniature postsynaptic potentials provides information on the amount and density of synaptic connections (frequency), as well as the postsynaptic receptor abundance (amplitude), and hence helps to study elementary synapse properties. However, higher levels of synaptic function, including the responses of synapses in regard to action potential firing, as well as short- and long-term adaptations of synaptic strength, require the analysis of evoked synaptic transmission. For example, paired-pulse stimulation protocols can serve as a basic model for studying short-term synaptic plasticity ex vivo (Bouteiller et al., 2010). Evoked synaptic transmission and plasticity in specific neuronal pathways can typically be studied in brain slices (Wang and Baudry, 2019). Alternatively, synaptic transmission and plasticity can be studied in cultured neurons, such as by employing optogenetic activation of neuronal cell populations (Barral and Reyes, 2017). In an acute brain slice, which is the standard model for the electrophysiological analysis of synaptic functions, presynaptic stimulation and postsynaptic responses can only be analyzed in one direction. However, as synaptic plasticity involves the possibility of changes in both pre- and post-synaptic components, one-directional measurements limit the study of mechanisms involved in modulating plasticity.
Here, we describe a protocol for culturing simple networks of paired hippocampal neurons for the bi-directional electrophysiological analysis of synaptic functions. This cellular ex vivo model has the following advantages: first, it allows the easy identification of the innervated cells. In classical neuronal cell cultures employing dispersed neurons, this is inherently difficult due to excessive branching of the axons and the possibility of hetero-synaptic innervation of the target neuron. Second, due to the defined simple network, all synapses are formed between the paired neurons. This results in increased amplitudes of postsynaptic responses and hence allows the reliable detection of changes in postsynaptic receptor function. Third, both cells of the cultured paired network can function as presynaptic (stimulated) and postsynaptic (innervated) neurons. Hence, this method allows recording synaptic transmission bi-directionally. This is particularly relevant in the context of genetic manipulations of one of the two paired neurons: as one of the paired neurons can be genetically modified by overexpression or knockdown of specific proteins, bi-directional stimulation protocols allow analyzing pre- and post-synaptic consequences in comparison with wild-type synaptic connections in the same neuronal network. As a proof of principle, we altered the expression of α2δ proteins, which act, on the one hand, as auxiliary subunits of voltage-gated calcium channels (Geisler et al., 2015; Ablinger et al., 2020; Dolphin and Obermair, 2022), and on the other hand as critical synaptic organizers (Eroglu et al., 2009; Geisler et al., 2019; Schöpf et al., 2021; Ablinger et al., 2022). Hence, we employed cultured paired hippocampal neurons to investigate the role of a splice variant of the α2δ-2 isoform in the trans-synaptic regulation of synapse formation and synaptic transmission, including short-term synaptic plasticity.
Materials and reagents
Animals
Timed pregnant wild-type mice (BALB/c, gestational age 16–17 days; Charles River Laboratories, Sulzfeld, Germany).
Biological materials
Lentiviral particles, carrying RNA encoding for the α2δ-2_ΔE23 splice variant and soluble eGFP as fluorescent marker (Geisler et al., 2019). Lentiviral particles were generated as previously described (Nasri et al., 2014; Benskey and Manfredsson, 2016).
Critical: Lentiviruses are classified as a biosafety level 2 (BSL-2) organism.
Materials
Surgical scissors, sharp blunt, straight 14.5 cm (Fine Science Tools, catalog number: 14001-14)
Tissue forceps, slim 1 × 2 teeth 10 cm (Fine Science Tools, catalog number: 11023-10)
Fine scissors, sharp, curved 10.5 cm (Fine Science Tools, catalog number: 14061-10)
Fine scissors, sharp, straight 10.5 cm (Fine Science Tools, catalog number: 14060-10)
Dumont #5 standard forceps (Fine Science Tools, catalog number: 11251-30)
Dumont #5 biology forceps (Fine Science Tools, catalog number: 11252-30)
Vannas-Tübingen spring scissors (Fine Science Tools, catalog number: 15004-08)
18 mm glass coverslips (Marienfeld Superior, catalog number: 0111580)
Rack for coverslips (custom build, Institute of Physiology, Medical University Innsbruck, Austria)
PTFE dish (Carl Roth, catalog number: K837.1)
12.5 cm filter paper (Carl Roth, catalog number: AP86.1)
Hemacytometer (Neubauer, catalog number: Brand 717805)
72 μm nylon mesh (Falcon, catalog number: 352350)
T75 flask (Falcon, catalog number: 353810)
Transfer pipette 3.5 mL (Sarstedt, catalog number: 86.1171.001)
15 mL centrifuge tube (Falcon, catalog number: 352070)
50 mL centrifuge tube (Falcon, catalog number: 352096)
60 mm plastic Petri dish (Falcon, catalog number: 353802)
60 mm Primaria plastic Petri dish (Falcon, catalog number: 353004)
15 cm glass Petri dish (Duran, catalog number: 237555201)
Pasteur pipette (Assistent, catalog number: 40567002)
5 mL serological pipette (Sarstedt, catalog number: 86.1253.001)
10 mL serological pipette (Sarstedt, catalog number: 86.1254.001)
25 mL serological pipette (Sarstedt, catalog number: 86.1685.001)
1.5 mL miniature spray (Rene Lezard)
Borosilicate glass with filament (Sutter Instrument, model: BF150-75-10)
2.5% trypsin (10×) (Gibco, catalog number: 15090-046)
0.5% Trypsin-EDTA (10×) (Gibco, catalog number: 15400-054)
B-27 supplement (50×) (Gibco, catalog number: 17504-044)
GlutaMAX (Gibco, catalog number: 35050-038)
Horse serum (Gibco, catalog number: 16050-122)
PenStrep (Penicillin-Streptomycin) (Gibco, catalog number: 15140-122)
MEM (Gibco, catalog number: 41090-028)
Neurobasal medium (Gibco, catalog number: 21103-049)
HBSS (10×) (Gibco, catalog number: 14180-046)
HEPES 1 M solution (Gibco, catalog number: 15630-056)
Poly-L-lysine (Sigma, catalog number: P2636)
Ara-C (Sigma, catalog number: C6645)
DNase (Sigma, catalog number: DN-25)
Sodium pyruvate (Sigma, catalog number: P2256)
Paraffin (Carl Roth, catalog number: X880.1)
Gelatine (Fluka, catalog number: 48722)
Nitric acid (Carl Roth, catalog number: 4989.2)
Glucose (Carl Roth, catalog number: HN06.3)
Boric acid (Sigma, catalog number: B6768)
Borax (sodium tetraborate decahydrate) (Sigma, catalog number: B9876)
Sodium chloride (NaCl) (Carl Roth, catalog number: 3957.1)
Potassium chloride (KCl) (Carl Roth, catalog number: 6781.3)
Calcium chloride dihydrate (CaCl2·2H2O) (Carl Roth, catalog number: 5239.2)
Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma, catalog number: M0250)
Sodium hydroxide (NaOH) (Carl Roth, catalog number: 6771.3)
Potassium hydroxide (KOH) (Carl Roth, catalog number: 6751.1)
Gluconic acid, potassium salt (K-gluconate) (Carl Roth, catalog number: 4621.1)
HEPES (Carl Roth, catalog number: 6763.1)
EGTA (Sigma, catalog number: E3889)
ATP, magnesium salt (Sigma, catalog number: A9187)
GTP, sodium salt (Sigma, catalog number: G8877)
Solutions
Pyruvate solution, 100 mM, 50 mL (see Recipes)
1% gelatine solution, 50 mL (see Recipes)
HBSS, 500 mL (see Recipes)
Glia medium, 500 mL (see Recipes)
Neuronal maintenance medium, 200 mL (see Recipes)
Neuronal plating medium, 200 mL (see Recipes)
1% DNase solution, 100 mL (see Recipes)
Sodium borate buffer, 500 mL (see Recipes)
Poly-L-lysine solution, 1 mg/mL (see Recipes)
EGTA solution, 0.5 M, 1 mL (see Recipes)
Extracellular solution, 100 mL, adjust pH 7.4 with NaOH (see Recipes)
Intracellular solution, 20 mL, adjust to pH 7.2 with KOH (see Recipes)
Sodium hydroxide solution, 1 M, 5 mL (see Recipes)
Potassium hydroxide solution, 1 M, 5 mL (see Recipes)
20% glucose solution, 50 mL (see Recipes)
Recipes
Pyruvate solution, 100 mM, 50 mL
Reagent Amount
Sodium pyruvate 550 mg
Milli-Q water Add to the total volume of 50 mL
1% gelatine solution, 50 mL
Reagent Amount
Gelatine 500 mg
Milli-Q water Add to the total volume of 50 mL
HBSS, 500 mL
Reagent Amount
HBSS (10×) 50 mL
PenStrep 5 mL
HEPES, 1 M 5 mL
Milli-Q water 440 mL
Glia medium, 500 mL
Reagent Amount
MEM 430 mL
PenStrep 5 mL
Glucose, 20% 15 mL
Horse serum 50 mL
Neuronal maintenance medium, 200 mL
Reagent Amount
Neurobasal medium 194 mL
B-27 supplement, (50×) 4 mL
GlutaMAX 2 mL
Neuronal plating medium, 200 mL
Reagent Amount
MEM 172 mL
Pyruvate solution, 100 mM 2 mL
Glucose, 20% 6 mL
Horse serum 20 mL
1% DNase solution, 100 mL
Reagent Amount
DNase 1 g
HBSS 100 mL
Sodium borate buffer, 500 mL
Reagent Final concentration Amount
Boric acid 50 mM 1.54 g
Borax (sodium tetraborate) 12.5 mM 2.376 g
Milli-Q water Add to the total volume of 500 mL
Poly-L-lysine solution, 1 mg/mL
Reagent Amount
Poly-L-lysine 100 mg
Sodium borate buffer 100 mL
EGTA solution, 0.5 M, 1 mL
Reagent Amount
EGTA 190.2 mg
1 M KOH Add to the total volume of 1 mL
Extracellular solution, 100 mL, adjust pH 7.4 with 1 M NaOH solution
Reagent Final concentration Amount
NaCl 137 mM 800.62 mg
KCl 3 mM 22.36 mg
Glucose 10 mM 180.2 mg
HEPES 10 mM 238.3 mg
CaCl2·2H2O 1.8 mM 26.46 mg
MgCl2·6H2O 2 mM 40.66 mg
Intracellular solution, 20 mL, adjust to pH 7.2 with 1 M KOH solution
Reagent Final concentration Amount
K-gluconate 125 mM 585.62 mg
KCl 10 mM 14.9 mg
HEPES 10 mM 47.66 mg
MgCl2·6H2O 1 mM 4.06 mg
EGTA solution, 0.5M 2 mM 80 μL
ATP (Mg) 4 mM 40.57 mg
GTP (Na) 0.3 mM 3.14 mg
Sodium hydroxide solution, 1 M, 5 mL
Reagent Amount
NaOH 200 mg
Milli-Q water Add to the total volume of 5 mL
Potassium hydroxide solution, 1 M, 5 mL
Reagent Amount
KOH 280.6 mg
Milli-Q water Add to the total volume of 5 mL
20% glucose solution, 50 mL
Reagent Amount
Glucose 10 g
Milli-Q water Add to the total volume of 50 mL
Equipment
Stereo dissection microscope (Olympus, model: SZX2-ILLTQ)
Class II biological safety cabinet (Thermo Scientific, model: HERAsafe KS12)
Suction system (Welch, model: 112037-08)
Safety Bunsen burner (INTEGRA Biosciences, model: Fireboy Plus 144000)
Pipette controller (INTEGRA Biosciences, model: PIPETTEBOY acu 2)
Magnetic stirrer with heating (Phoenix Instrument, model: RMS-10HS)
CO2 incubator (Thermo Scientific, model: HERAcell 240i)
Water baths (VWR, catalog number: 462-0558)
pH meter (inoLab, model: pH 7110)
Micropipette puller (Sutter Instrument, model: P-97)
Microforge (Narishige, model: MF-830)
Ultrapure water purification system (Merck Millipore, model: Milli-Q® EQ 7000)
Universal oven (Memmert, model: UF260)
Experimental setup
CleanBench lab table (TMC, model: TM-63-9012S)
Inverted fluorescent microscope (Olympus, model: IX-83, equipped with LUCPLFLN40XPH/0.6 objective, Lumencor LED lamp, eGFP filter)
Microscope dependent platforms (Sutter Instrument, model: MDM-83-2)
Micromanipulators (Sutter Instrument, model: MPC-325-2)
Two-channel patch clamp amplifier (HEKA Elektronik, model: EPC 10 USB Double)
Quick change chamber RC-41LP (Harvard Apparatus, catalog number: 64-0368)
Software
PatchMaster v2x90.5 (HEKA Elektronik)
FitMaster v2x90.5 (HEKA Elektronik)
Procedure
Critical: Steps A1e–A1k, A1m, A2d–A2f, A2h–A2l, A3b–A3h, and A3j–A3t should be performed under sterile conditions. Steps A2i–A2j should be performed in a biosafety level 2 cabinet.
Culturing paired hippocampal neurons in coculture with an astroglia feeder layer
Cell culture method was adapted from Kaech and Banker (2006) with slight modifications (Obermair et al., 2004; Geisler et al., 2019).
Coverslip preparation
Critical: It is necessary to start preparing the coverslips at least four days prior to the preparation of hippocampal cultures.
Place 18 mm coverslips in custom built racks and rinse in Milli-Q water.
Place racks with coverslips in concentrated nitric acid (63%) for 12 h.
Rinse racks with coverslips in Milli-Q water; perform two changes for 2 h each.
Transfer coverslips into 15 cm glass Petri dishes on top of 12.5 cm filter paper and sterilize with dry heat (200 °C for 10–12 h).
Transfer coverslips into 60 mm plastic Petri dishes (five per dish, see Figure 1).
Note: To prevent coverslips from swimming off, put 10 μL of 1% sterile-filtered gelatine solution into the dish for each coverslip before transferring it into the dish. Coverslips can be easily transferred to the plastic dish by using a glass Pasteur pipette connected to the suction system.
Melt sterile paraffin on the heated magnetic stirrer to 150 °C in a PTFE dish.
Apply three small drops of melted paraffin to each coverslip.
Note: Paraffin can be transferred from PTFE dish to coverslips using fire-polished Pasteur pipettes (Figure 1). Dip the pipette into paraffin and touch the coverslip quickly before paraffin hardens. Repeat procedure for each paraffin dot.
Figure 1. Fire-polished Pasteur pipette and 18 mm cover glasses with attached paraffin dots. Fire-polish the tip of the Pasteur pipette with a Bunsen burner. The tip should melt to form a glass bead with a diameter of approximately 2 mm (while heating the tip of the pipette, rotate it around the longitudinal axis to form a symmetrical bead), and the pipette should be bent by an angle of 50°–80° approximately 3 cm from the end of pipette.
Sterilize coverslips by UV irradiation for 30 min in the laminar flow (class II biological safety cabinet).
Spray poly-L-lysine solution using a 1.5 mL miniature spray bottle onto coverslips and let the poly-L-lysine dry at room temperature overnight.
Critical: The poly-L-lysine solution should not be sprayed directly onto coverslips. Instead, small droplets of the solution, formed at the time of spraying, must settle down on the coverslips by gravity (Video 1). Dish with coverslips should be placed approximately 5 cm below the nozzle of the miniature spray.
Video 1. Small droplets of poly-L-lysine solution are formed during spraying (video at 32× reduced speed). The dish with the coverslips should be placed parallel to and beneath the main spray stream approximately 0.5–1 s after spraying.
Rinse dishes with poly-L-lysine-treated coverslips with sterile Milli-Q water; perform two changes for 2 h each.
Remove final rinse and add 4 mL of neuronal plating medium per dish.
Put dishes in cell culture incubator (37 °C, 5% CO2). The hippocampal neurons will be plated into these dishes.
Before plating neurons on coverslips, replace 4 mL of neuronal plating medium with fresh medium.
Dissection and preparation of paired hippocampal cultures
Kill the pregnant mouse by cervical dislocation, remove uterus, and place it in a sterile 10 cm Petri dish. Remove fetuses from the uterus.
Quickly decapitate fetuses with scissors and place heads in HBSS (4 °C).
Under a dissecting microscope: dissect out brains, separate brain hemispheres, strip away the meninges, and cut out the hippocampi as shown in Video 2.
Video 2. Dissection procedure of hippocampi from 16–17-day-old mouse embryos. This video was made in the MiMo Laboratory at the Karl Landsteiner University, which is a user facility in accordance with §16 TVG 2012, license number 2021-0.412.631, approved by the Austrian Federal Ministry of Science, Research and Economy.
Note: For all steps following the dissection of hippocampi, all media and reagents should be prewarmed to 37 °C before use.
Using the transfer pipette, place all the hippocampi from one litter in a 15 mL Falcon centrifuge tube. Bring the total volume to 4.5 mL with HBSS and add 0.5 mL of 2.5% trypsin (10×). Incubate in a water bath at 37 °C for 15 min.
Remove the trypsin solution, add 5 mL of HBSS (gently tap or swirl the tube to mix), and let stand for 5 min. Repeat this step twice, finally bringing the volume to 3 mL (4 mL if hippocampi from more than five brains are used).
Dissociate the cells by gently pipetting hippocampi up and down, first with a Pasteur pipette with a fire-polished tip to half the normal diameter, and then with a Pasteur pipette with a tip fire-polished to nearly a quarter the normal diameter. Continue pipetting gently until no chunks of tissue remain (approximately 7–8 times).
Determine the density of cells using a hemacytometer.
Add 25,000 cells to each of the dishes containing the poly-L-lysine-treated coverslips in neuronal plating medium (corresponding plating density is 880 cells/cm2).
For viral infection, add medium containing lentiviral particles to the dish with freshly added neurons.
Note: To reach an approximate 50% viral infection efficiency, the volume and concentration of lentivirus added should be defined experimentally. Added medium with lentiviral particles should not exceed 1 mL per Petri dish containing 4 mL of neuronal plating medium.
After 3 h and using forceps, transfer the coverslips with the neurons attached into dishes containing the glial cells in neuronal maintenance medium (see step A3). Turn the coverslips upside down so that the neurons are facing down, towards the glial cells.
To reduce glial proliferation, add Ara-C (5 μM) three days after plating the neurons.
Once a week, remove 2 mL of the neuronal maintenance medium and replace it with fresh medium.
For electrophysiological recordings, use cultured hippocampal neurons at the age 14–17 days in vitro (DIV) (Figure 2).
Figure 2. A pair of synaptically connected hippocampal neurons (left panel, phase contrast micrograph). One neuron is virally transfected with the α2δ-2 protein and soluble eGFP (middle, fluorescence micrograph and right panel, overlayed phase contrast and fluorescent micrographs), the other neuron is an untransfected control neuron (arrow). Scale bar, 20 μm.
Preparation of astroglia feeder layer
Critical: It is necessary to start preparing the glia feeder layer 14 days prior to the preparation of hippocampal cultures.
Prepare brain hemispheres as described in steps A2a–A2c.
Notes:
i. If neuronal cultures are prepared on a regular basis, cells for the glia feeder layer can be prepared from the brains of mice used for the preparation of the neuronal culture.
ii. For all steps following the preparation of brain hemispheres, all media and reagents should be prewarmed to 37 °C before use.
Mince brain hemispheres into small pieces with Vannas-Tübingen spring scissors.
Transfer minced tissue to a 50 mL Falcon centrifuge tube in a final volume of 12 mL of HBSS.
Add 1.5 mL of 1% DNase solution and incubate in water bath for 5 min at 37 °C.
Add 1.5 mL of 2.5% trypsin (10×) and incubate for 15 min at 37 °C. During trypsin treatment, dissociate tissue every 5 min with a 5 mL serological pipette (pipette tissue up and down 7–8 times).
Critical: It is critical to add the DNase solution 5 min before adding the trypsin. Otherwise, DNase will be quickly degraded, resulting in abundant DNA material from minced brain tissues, which will strongly decrease the cell yield.
Add 3 mL of horse serum to inhibit trypsin activity.
Filter the combined supernatants through a 72 μm nylon mesh to remove any undissociated tissue.
Centrifuge supernatants at 200× g for 5 min at 4 °C and resuspend cells in 5 mL of glial medium.
Determine cell density with an hemacytometer.
Transfer 4,000,000 cells into a 75 cm2 T-flask (equivalent to approximately 1–1.5 brain hemispheres per flask). Add glial medium to a total volume of 13 mL. Put the flask in a cell culture incubator (37 °C, 5% CO2).
Note: One 70%–80% confluent T-flask with astroglia cells will be enough to prepare 10 mm × 60 mm dishes of astroglia feeding layer.
Three days after plating, replace glial medium in the flask completely with fresh medium.
Seven days after plating, shake the flask vigorously to dislodge microglia and remove them by washing (replace glia medium completely with fresh medium). Return the flask to the cell culture incubator.
Critical: Slap the flask forcefully 2–3 times on a hard surface so that the medium foams up and the entire content appears messed up. Gently tapping the flask will not dislodge the microglia, which are typically accumulating on top of astrocytes.
When cells in the flask have reached confluence (usually 10 days after plating), remove the glial medium and wash with 10 mL of HBSS.
Remove HBSS, add 10 mL of 0.5% trypsin-EDTA (in HBSS), and put flask in cell culture incubator for 5 min.
Tap the flask gently on the side to detached remaining astrocytes, add 1 mL of horse serum, and place the flask upright to allow the cells to slide to the bottom.
Transfer glial cell suspension into a 50 mL Falcon centrifuge tube and centrifuge at 200× g for 5 min at room temperature.
Resuspend cell pellet in 40 mL of glial medium.
Add 4 mL of glial cell suspension per 60 mm Primaria plastic Petri dish. Put dishes in cell culture incubator (37 °C, 5% CO2).
On the next day, replace glia media in Petri dishes with fresh medium.
On the third day (one day before preparing hippocampal cultures), replace glia medium in dishes with astroglia layer with 6 mL of neuronal maintenance medium. Return dishes to the cell culture incubator.
Electrophysiological recordings of induced postsynaptic responses in paired cultured neurons
Notes:
i. The following steps require the experimenter to have knowledge and proficiency in two-channel patch clamp electrophysiology. All recordings and analyses were performed using PatchMaster software.
ii. To induce synaptic transmission within paired neurons, each neuron will be interchangeably stimulated with a depolarization using an action potential wave form, which needs to be recorded from wild-type (WT) neurons.
Recording of action potentials from WT hippocampal neurons
Pull glass patch pipette with 3–5 mm short taper and fire-polish its tip using a microforge. Resistance of the pipette should be 2–4 MΩ when filled with the intracellular solution.
Place coverslip with WT neurons into the low-profile chamber (Figure 3) and add 300 μL of extracellular solution.
Figure 3. Experimental setup. Low-profile chamber is mounted on the precision controlled XY-stage. Probes 1 and 2 will be used for paired recordings.
Using probe 1, patch any neuron in voltage clamp mode using the whole-cell configuration. Holding potential (Vm) should be set to -70 mV.
Switch from voltage clamp to current clamp mode.
Record spontaneous activity (generation of action potentials) of the neuron using the protocol shown in Figure 4.
Note: If no spontaneous activity is observed in the neuron, depolarize neurons by continuous injection of an electrical current (5 pA, “I-membrane” box in “Amplifier” window). If necessary, increase amplitude of injected current with steps of 5 pA until the neurons start generating action potentials.
Figure 4. Protocol for recording spontaneous activity of neurons using probe 1. Screenshot of the “Pulse generator” window.
In the Replay window, choose the recorded trace and zoom in on one action potential.
Export the recorded action potential as a stimulation template file (Figure 5).
Figure 5. Export of the recorded action potential as stimulation template file
Single stimulation of paired neurons
Prepare two stimulation protocols, as shown in Figure 6.
Note: Duration [ms] of recording time in Segments can be arbitrary but should be long enough to record a postsynaptic response.
Critical: If your stimulation protocol has the name “Stim,” then the corresponding stimulation template file must have the name “Stim_1.tpl” and it needs to be stored in the folder where the PatchMaster *.pgf file is located, for example at “C:\Program Files (x86)\HEKA\PatchMaster\.” Each stimulation protocol must have its own stimulation template file.
Figure 6. Protocol for recording induced synaptic response. A. Stimulation protocol for stimulating the cell patched with probe 1 and postsynaptic response recording from cell patched with probe 2. “Stim-1” should be chosen as a stimulus in the “DA” section; in the “AD” section, stimulation signal and postsynaptic responses are recorded as Vmon-1 and Imon-2, respectively. B. Stimulation protocol for stimulating the cell patched with probe 2 and postsynaptic response recording from the cell patched with probe 1. Screenshot of the “Pulse generator” window. “Stim-2” should be chosen in the “DA” section; Vmon-2 and Imon-1 should be chosen in the “AD” for recording.
Pull glass patch pipettes with 3–5 mm short taper and fire-polish its tip using a microforge. Resistances of pipettes should be 2–4 MΩ when filled with the intracellular solution.
Place coverslip with neurons into the chamber and add 300 μL of extracellular solution.
Patch both neurons in the voltage clamp mode using whole-cell configuration. Holding potential (Vm) should be set to -70 mV for both cells.
Alternately, apply the stimulation protocols for both patched cells. Induced postsynaptic response will be recorded from the respective unstimulated cell (Figure 7).
Figure 7. Recorded postsynaptic current (black trace) from the paired postsynaptic neuron after stimulation of the paired presynaptic neuron with an action potential (blue trace). Screenshot of “Oscilloscope” window.
Paired-pulse stimulation of paired neurons
Prepare protocol for paired-pulse stimulation, as shown in Figure 8.
Figure 8. Stimulation protocol for paired-pulse stimulation of the cell patched with probe 1 and postsynaptic response recording from the cell patched with probe 2. Stimulation protocol represents a series of stimulations by two sequential depolarizations from -70 mV to 60 mV (5 ms duration each) with increasing inter-pulse interval. To stimulate cells on probe 2 with the response recording on probe 1, replace values in “DA” and “AD” sections (1 into 2, 2 into 1).
After application of a single stimulation alternately, apply the pair-pulse stimulation protocol to both presynaptic/postsynaptic cell configurations.
Data analysis
To analyze amplitudes of recorded induced postsynaptic responses, create the two functions “Minimum” and “Series time” in the window Analysis, as shown in Figure 9.
Note: The function “Minimum” is used to analyze postsynaptic currents in response to excitatory (glutamatergic) synaptic transmission. To analyze inhibitory (GABAergic) synaptic transmission, use the function “Maximum,” as the postsynaptic currents have opposite polarity of the peak current.
Figure 9. Analysis protocol for measuring current amplitude of postsynaptic response functions “Minimum” (A) and “Series time” (B). Screenshot of the “Analysis” window.
After application of the single stimulation protocol, read peak value of the synaptic current in Notebook window.
For the analysis of individual peak amplitudes after pair-pulse stimulation, chose the segment of the recorded trace to be analyzed (change Cursor Bounds (%) in Analysis window) and replay trace for analysis of each peak (Figure 10).
Note: Alternatively, analysis of the amplitude of recorded postsynaptic response can be performed using the FitMaster software.
Figure 10. Analysis of individual peak amplitudes after pair-pulse stimulation. Changing of cursor bounds is represented by the vertical lines on the Oscilloscope window.
General notes and troubleshooting
While performing experiments according to the current protocol, the experimenter may experience two types of problems related to the formation of neuronal pairs and the efficiency of viral infection. Both problems can be easily solved. If more than two neurons form networks on most of the poly-L-lysine spots (Figure 11), the plating density should be reduced (see step A2h). Increasing plating density is required if the majority of poly-L-lysine spots contain only one neuron.
Figure 11. Neuronal networks of cultured hippocampal neurons, formed by two (A, aim of the protocol) or three (B, plating density should be reduced) cells. Scale bar, 10 μm.
To successfully perform experiments on paired neurons in which one neuron was genetically modified, the efficiency of viral infection should be approximately 50%. If both neurons are virally infected or none are infected, it is necessary to decrease or increase, respectively, the concentration of the virus that is added to the freshly plated hippocampal neurons (step A2i).
Validation of protocol
The presented protocol was developed and successfully employed for the research work published in Geisler et al. (2019), in order to analyze the consequences of altered synaptic wiring induced by the α2δ2_ΔE23 isoform (section “Reduced synaptic transmission in aberrantly wired synapses,” Figure 12 in the article).
Acknowledgments
This study was supported by the Austrian Science Fund (FWF), grants P24079, F44060, F44150, DOC30-B30, and the Gesellschaft für Forschungsförderung Niederösterreich (NFB, grant LSC19-017). The presented protocol was developed and successfully employed for the research work published in Geisler et al. (2019).
Competing interests
We declare neither competing interests nor conflicts of interest.
Ethical considerations
Animal procedures were performed in compliance with EU regulations and national regulations and were approved by the Austrian Federal Ministry of Science, Research and Economy. MiMo Laboratory of Karl Landsteiner University is approved as user in accordance with §16 TVG 2012, license number 2021-0.412.631.
References
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Ablinger, C., Geisler, S. M., Stanika, R. I., Klein, C. T. and Obermair, G. J. (2020). Neuronal α2δ proteins and brain disorders. Pflugers Arch. - Eur. J. Physiol 472(7): 845-863.
Barral, J. and Reyes, A. (2017). Optogenetic Stimulation and Recording of Primary Cultured Neurons with Spatiotemporal Control. Bio Protoc 7(12): e2335.
Benskey, M. J. and Manfredsson, F. P. (2016). Lentivirus Production and Purification. Methods Mol Biol 1382: 107-114.
Bouteiller, J. M., Allam, S. L., Greget, R., Ambert, N., Hu, E. Y., Bischoff, S., Baudry, M. and Berger, T. W. (2010). Paired-pulse stimulation at glutamatergic synapses - pre- and postsynaptic components. Annu Int Conf IEEE Eng Med Biol Soc 2010: 787-790.
Deperrois, N. and Graupner, M. (2020). Short-term depression and long-term plasticity together tune sensitive range of synaptic plasticity. PLoS Comput. Biol 16(9): e1008265.
Dolphin, A. C. and Obermair, G. J. (2022). Regulation of Calcium Channels and Synaptic Function by Auxiliary α2δ Subunits. In: Zamponi, G. W. and Weiss, N. (Eds.) Voltage-Gated Calcium Channels. (pp. 93-114). Springer International Publishing.
Eroglu, C., Allen, N. J., Susman, M. W., O’Rourke, N. A., Park, C. Y., Ozkan, E., Chakraborty, C., Mulinyawe, S. B., Annis, D. S., Huberman, A. D., et al. (2009). Gabapentin receptor alpha2delta-1 is a neuronal thrombospondin receptor responsible for excitatory CNS synaptogenesis. Cell 139(2): 380-392.
Geisler, S., Schöpf, C. L. and Obermair, G. J. (2015). Emerging evidence for specific neuronal functions of auxiliary calcium channel α2δ subunits. Gen. Physiol. Biophys 34(2): 105-118.
Geisler, S., Schöpf, C. L., Stanika, R., Kalb, M., Campiglio, M., Repetto, D., Traxler, L., Missler, M. and Obermair, G. J. (2019). Presynaptic α2δ-2 Calcium Channel Subunits Regulate Postsynaptic GABA(A) Receptor Abundance and Axonal Wiring. J. Neurosci. 39(14): 2581-2605.
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Neuroscience > Cellular mechanisms > Synaptic physiology
Biophysics > Electrophysiology > Patch-clamp technique
Cell Biology > Cell signaling > Synaptic transmision
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Optogenetic Induction of Pyroptosis, Necroptosis, and Apoptosis in Mammalian Cell Lines
KS Kateryna Shkarina
PB Petr Broz
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4762 Views: 1247
Reviewed by: Rajesh RanjanTakashi AkeraEmmanuelle Berret
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Original Research Article:
The authors used this protocol in The Journal of Cell Biology Apr 2022
Abstract
Regulated cell death plays a key role in immunity, development, and homeostasis, but is also associated with a number of pathologies such as autoinflammatory and neurodegenerative diseases and cancer. However, despite the extensive mechanistic research of different cell death modalities, the direct comparison of different forms of cell death and their consequences on the cellular and tissue level remain poorly characterized. Comparative studies are hindered by the mechanistic and kinetic differences between cell death modalities, as well as the inability to selectively induce different cell death programs in an individual cell within cell populations or tissues. In this method, we present a protocol for rapid and specific optogenetic activation of three major types of programmed cell death: apoptosis, necroptosis, and pyroptosis, using light-induced forced oligomerization of their major effector proteins (caspases or kinases).
Keywords: Programmed cell death Apoptosis Pyroptosis Necroptosis Optogenetics
Background
Regulated cell death is a common feature of multicellular organisms and plays a key role in development and tissue homeostasis and in protecting the host against malignant growth and various pathogens. Research over the last two decades has identified over 12 different forms of regulated cell death (Galluzzi et al., 2018); however, their study is often complicated by the complex crosstalk and interconnectivity between the different cell death pathways (Bedoui et al., 2020). Additionally, the consequences of different types of cell death in the tissue still remain insufficiently understood, which is at least partially due to the challenges of selectively targeting single cells in multicellular populations, as well as to the pleiotropic effects of commonly used natural cell death triggers both on the dying cells and their neighbors.
In recent years, multiple strategies have been developed to specifically ablate cells both in vitro and in living animals. However, some of these methods (such as laser ablation or photosensitization) still lack specificity regarding the type of cell death to be induced (Tirlapur et al., 2001; Qi et al., 2012), while others [such as chemically inducible dimerization of apoptotic or necroptotic effector proteins (Oberst et al., 2010; Wu et al., 2014)] suffer from a limited spatiotemporal control and require a delivery of soluble ligands, thus limiting their in vivo applications. To overcome these limitations and expand the scope of the tools available for programmed cell death induction, we recently developed a set of optogenetically activated cell death effectors (optoCDEs) (Shkarina et al., 2022), which enable selective induction of three major types of programmed cell death: apoptosis, pyroptosis, and necroptosis. These tools consist of three modules: 1) a photoactuator domain Cry2olig (Cry2 E490G), which responds to blue light by rapid homo-oligomerization, 2) an mCherry tag, which enables the detection of the cells expressing optoCDEs and estimation of the relative construct expression levels, and 3) an effector module (Figure 1A–1C). For opto-caspases, the effector module corresponds to the protease domain (p20 and p10 subunits) of corresponding caspases; the endogenous linkers and cleavage sites essential for the caspase activation are retained, while CARD (in caspase-1, -4, -5, -9, and -11) and DED (caspase-8) domains, responsible for the endogenous upper-level protein–protein interactions and homo-oligomerization, are removed. In opto-RIPK3, the design is similar, while the RHIM motif (responsible for the upstream interaction with the RIPK1) is mutated. In optoMLKL, the effector domain orientation in relation to Cry2olig and mCherry is reversed to keep the MLKL N-terminus (responsible for membrane binding and disruption) exposed. The considerations behind the construct design and testing of the different construct versions are described in more detail in the original paper (Shkarina et al., 2022). The blue light stimulation triggers the rapid activation and oligomerization of Cry2olig, which in turn results in the proximity-induced activation of effector domains and subsequent processing of downstream substrates, culminating in cell death. While Cry2olig alone responds to the blue light within seconds (Taslimi et al., 2014), the timing of cell death induction is defined by the kinetics of effector activation as well as availability and efficiency of the processing and activation of downstream substrates (such as apoptotic executioner caspases, necroptotic effector GSDMD, or pyroptotic effector MLKL); the first morphological features of cell death can usually be detected within minutes after the beginning of illumination.
Figure 1. Schematic representation of the optogenetically activated cell death effectors (optoCDE) system. (A) OptoCDEs are inactive and monomeric in the dark state but are activated within seconds upon blue light illumination, which induces oligomerization of the Cry2olig photoactuator domain, activation of downstream substrates, and induction of cell death. (B) General architecture of major cell death effectors and design of optoCDE constructs. Cell death effectors used in the study generally consist of an adaptor domain (CARD or DED for caspases and RHIM for RIPK3) and an effector domain bearing protease (caspases), kinase (RIPK3), or membrane-disrupting (MLKL) function. In the optoCDEs, the effector domain is retained, while the adaptor is replaced with the Cry2olig-mCherry photoactuator module (mCherry is used to visualize optoCDE expression and/or clustering). (C) Schematic representation of major types of optoCDE constructs used in the study. The detailed description and evaluation of the optoCDE tools is available in the original paper (Shkarina et al., 2022). Figure adapted from Shkarina et al. (2022).
The optoCDEs can be applied in vitro, as described in this protocol, as well as in vivo to selectively kill specific cells or cell populations in a highly controlled and specific manner (Shkarina et al., 2022). Additionally, the precise control over the illumination parameters, such as light intensity and duration, provides new means for cellular and mechanistic studies of these forms of cell death, as well as probing cell survival mechanisms that limit cellular damage downstream of these effectors.
Materials and reagents
Cell culture and lentiviral transduction
Sterile serological pipettes (Falcon, catalog number: 357543)
Sterile micropipette tips (Starlab TipOne, catalog number: S1120-8810)
Tissue culture–treated cell culture flasks (TPP, catalog number: 90076)
10 cm Petri dishes (Falcon, catalog number: 351029)
Syringes (5 and 10 mL) (Braun Omnifix, catalog numbers: 4616103V and 4616057V)
Sterile tissue culture–treated 6-well flat-bottom plates (Eppendorf, catalog number: 0030720113)
Sterile 50 and 15 mL Falcon tubes (SPL life sciences, catalog numbers: 50015 and 50050)
Sterile 1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 11.3817.01)
Aluminum foil (Sigma, catalog number: 326852)
0.45 μm filters (Sarstedt, catalog number: 83.1826)
Polybrene (Merck, catalog number: TR-1003-G)
JetPRIME transfection reagent (Polyplus, catalog number: 101000027)
HEPES 1 M (Sigma, catalog number: H3375)
Dulbecco’s phosphate-buffered saline (DPBS) 1× (Thermo Fisher Scientific, catalog number: 10010023)
Dulbecco’s modified Eagle medium (DMEM) with GlutaMAX supplement (Thermo Fisher Scientific, catalog number: 10564011)
RPMI 1640 medium, with GlutaMAX supplement (Thermo Fisher Scientific, catalog number: 61870044)
Doxycycline hydrochloride (Sigma-Aldrich, catalog number: D3447)
Puromycin (Invivogen, catalog number: ant-pr-1)
Hygromycin B Gold (Invivogen, catalog number: ant-hg-1)
LPS-B5 ultrapure (Invivogen, catalog number: tlrl-b5lps)
PMA (Sigma-Aldrich, catalog number: P1585)
Fetal bovine serum (Bioconcept, catalog number: 2-01F10-I)
Live-cell imaging and cell death detection
8-well tissue culture–treated μ slides (ibidi, catalog number: 80826)
Collagen solution from bovine skin (Sigma-Aldrich, catalog number: C4243)
Opti-MEM reduced serum medium (Gibco, catalog number: 11058021)
CellTox Green (Promega, catalog number: G8741)
DRAQ7 (BioLegend, catalog number: 424001)
Annexin V Pacific Blue (BioLegend, catalog number: 640918)
Annexin V FITC (BioLegend, catalog number: 640906)
Annexin V Alexa Fluor 647 (BioLegend, catalog number: 640912)
CellEvent caspase-3/7 green (Thermo Fisher Scientific, catalog number: R37111)
Cell lysis and cytokine secretion analysis
Vision Plate 24, 150 micron, TC-treated, sterile (Life Systems Design, catalog number: 4ti-0241)
96-well flat-bottom plates
Lactate dehydrogenase (LDH) cytotoxicity detection kit (Sigma, catalog number: 11644793001)
Triton X-100 (Sigma, X100-500ML)
Human IL-1β ELISA kit (R&D, catalog number: DY401)
ELISA plates (Sigma-Aldrich, catalog number: M9410-1CS)
LDH stop solution [2 M acetic acid (Sigma, catalog number: 64-19-7)] in dH2O, store at 4 °C
Reagents and equipment for ELISA
Reagents and equipment for the Western Blot analysis
Fluorescence assay (Cisbio, catalog number: 62HIL1BPET)
Equipment
For cell culture
Tissue culture hood (such as HERASAFETM KS, Thermo Scientific)
Cell incubator (FormaTM Steri-CycleTM CO2 Incubator, Thermo Scientific)
Centrifuge (Eppendorf 5810R)
Light microscope (such as Leica DMI6000B)
Pipettes
For imaging
Point-scanning confocal (such as Zeiss LSM800 or Leica SP8)
For cell population–level assays
Light plate apparatus (Gerhardt et al., 2016) equipped with double row of 450 nm light LEDs
Spectrophotometer/ELISA plate reader
Multichannel pipettes
Western blot imager
Software
ZEN (ZEISS, https://www.zeiss.com/microscopy/en/products/software/zeiss-zen.html)
Iris (Jeff Tabor laboratory, http://taborlab.github.io/Iris/)
Fiji (NIH, https://imagej.net/software/fiji) (Version 2.3.0)
Software for the plate reader
Microsoft Excel (Microsoft, version 16.69.1)
GraphPad Prism (version 9.3.1.)
Procedure
Generation of stable cell lines expressing optogenetically activated cell death effectors (optoCDEs)
Lentiviral particle production
Prepare stocks of purified lentiviral plasmids encoding optoCDE constructs. We recommend using endotoxin-free midi- or maxi-prep kits for plasmid purification.
Twenty-four hours prior to transfection, plate HEK293T cells in a tissue culture–treated 6-well plate at 5 × 105 cells/well in 2 mL of fresh cell culture medium.
For each well, prepare the transfection mix, as summarized in Table 1, in 1.5 mL tubes. Mix gently and incubate at room temperature for 15 min.
Table 1. Transfection mix per well of a 6-well plate
Components Amount
pLVX-optoCDE 1.9 μg
PsPAX2 1.9 μg
VSVg 0.2 μg
JetPrime transfection reagent 5 μL
JetPrime Buffer 200 μL
Add the transfection mix dropwise to the cells. Ensure that the whole surface of the well is evenly covered (this can be assessed by monitoring the transient change of the medium color). Incubate the cells at 37 °C and 5% CO2.
At 6–12 h post transfection, replace the transfection mix with 2 mL of fresh cell culture medium for virus collection.
Note: Use the appropriate cell culture medium for the cell line to be transduced.
Incubate for an additional 36–48 h at 37 °C and 5% CO2.
Note: Over the course of virus production, the transfected cells might change the morphology, round up, and lift, as a consequence of viral particle release.
After 36–48 h, collect the virus-containing supernatants from transfected HEK293T cells into 15 mL Falcon tubes. At this step, the medium from the wells transfected with the same construct can be combined and further processed together.
Using a 5 or 10 mL syringe, gently pass the supernatants through a 0.45 μm syringe filter to remove cell debris and collect the filtrate into the clean 15 mL Falcon tube. Supplement the virus-containing supernatants with 20 μM HEPES (final concentration) to facilitate viral particle stability.
At this stage, the virus-containing supernatants can be either used directly or stored at -80 °C for several months.
Cell transduction
Seed the appropriate number of cells to be transduced.
i. For adherent cells: we recommend seeding 0.8 × 106–1 × 106 of cells per well of a 6-well plate, 6–24 h before transduction. Cells should be properly attached and cover 60%–70% of growth surface. The optimal seeding density can be adjusted to accommodate for the cell size and differences in the growth kinetics between different cell lines. We recommend using 2–5 mL/well of virus-containing medium for 6-well plates, 2 mL/well for 12-well plates, and 1 mL/well for 24-well plates.
ii. For non-adherent cell lines: cells can be collected, centrifuged, and resuspended in the virus-containing medium immediately prior to transduction. The second centrifugation step (spinfection) can be performed either in plates or in 50 mL Falcon tubes (in this case, the cells need to be resuspended and transferred to the plates or flasks following the spinfection).
iii. Include an additional well of mock-treated (polybrene-only treated) cells to control for cell viability and transduction efficiency.
Replace the culture medium with the virus-containing medium supplemented with 5 μg/mL polybrene to facilitate viral particle adhesion.
To facilitate the infection, centrifuge the cells with the virus at 3,000× g for 1 h at 37 °C. For the plate centrifugation, seal the sides of the plates with tape to prevent the accidental spillover of the supernatants during plate transfer and centrifugation. Suspension cells can also be centrifuged in 50 mL Falcon tubes. After the completion of this step, move the cells to the cell culture incubator.
After 6–12 h, aspirate the virus medium, wash cells 1–2 times with the pre-warmed DPBS, and add the appropriate volume of fresh growth medium. If cell density is too high, cells can be transferred into the larger vessel at this step.
Let the cells recover for 24–48 h to increase the cell number and allow for the proper expression of the selection markers and/or antibiotic resistance genes.
To select successfully transduced cells, replace the medium with fresh medium containing the appropriate antibiotic concentration.
i. Use the non-transduced wells as the positive control for selection efficiency.
ii. The optimal antibiotic concentration for each cell line can vary and has to be determined using an antibiotic titration.
iii. For most cell lines, we have successfully used the selection medium containing 1–5 μg/mL of puromycin or 50–100 μg/mL of hygromycin B Gold.
After 3–5 days, remove the antibiotic-containing medium and expand the surviving cells.
Cell line validation
To confirm the transduction efficiency, induce optoCDE expression by incubating the cells with 1 μg/mL doxycycline for 24–48 h (at this stage, cells need to be protected from light to prevent the optoCDE activation and cell death). Include the non-treated control (without doxycycline) to control for the leaky expression.
The percentage of mCherry-positive cells can be quantified using fluorescent microscopy or FACS. The efficiency of cell death induction can then be tested as described in sections B and C.
Note: OptoCDEs are fused with mCherry allowing easy assessment of construct expression.
Following selection and validation, the stable lines can either be frozen away in liquid nitrogen for storage or directly used for experiments.
Optogenetic induction and visualization of pyroptosis, necroptosis, and apoptosis using confocal microscopy
Cell preparation for imaging
One day before the experiment, seed the cells into the 8-well tissue culture–treated μ-slides at the concentration of 0.5 × 105–1 × 105 cells/well.
i. If using other type of cell culture plates, the cell concentration should be adjusted to achieve 30%–70% of the density at the day of experiment. We recommend using a lower cell density for experiments focused on the high-resolution visualization of the dynamic cellular events during different types of cell death in adherent cells.
ii. For poorly adherent cell types (such as HEK293T, MCF7, or equivalent), the slides can be additionally pre-coated with 5% collagen.
1) Prepare the appropriate volume of 5%–10% dilution of bovine skin collagen in sterile PBS. Mix thoroughly by inverting. Avoid vortexing, as this may lead to the collagen precipitation.
2) Add 100–150 μL of collagen solution per well. The volume of liquid should be sufficient to fully cover the growth surface.
3) Incubate the slides at room temperature for 5 min; then, aspirate the collagen and air dry the slides for 30–60 min in the hood. During aspiration, avoid touching the bottom of the slide (growth surface), as this might lead to the disruption of the collagen layer and reduce cell adhesion.
4) The coated slides can be stored at room temperature for several weeks.
To induce the construct expression in the cells, supplement the cell culture medium with 1 μg/mL doxycycline and incubate for 16–24 h. The longer induction time can be used to achieve higher expression levels but can also result in increased cytotoxicity.
To protect the cells from light following expression induction, construct a dark chamber by fully covering a petri dish with foil (Figure 2). Alternatively, cells can be placed in a plastic box or an alternative type of light-impermeable container, which allows air circulation and humidification.
Figure 2. Examples of the chambers that can be used to protect optogenetically activated cell death effectors (optoCDE)-expressing cells from light
On the day of experiment: pre-heat the microscope to 37 °C. Ensure that the temperature is stabilized before the start of the imaging.
Gently wash the cells with pre-warmed PBS and replace the cell culture medium with pre-warmed Opti-MEM or alternative imaging medium. At this stage, all the manipulations need to be performed under dim light or red-light conditions to avoid spontaneous optoCDE construct activation.
Optional: for cell death detection, the following reagents can be directly added to imaging medium:
i. Visualization of membrane permeabilization:
1) CellTox Green: 1:10,000
2) DRAQ7: 1:1,000
ii. Visualization of PS exposure
1) Annexin V Pacific Blue: 1:500
2) Annexin V FITC 1:1,000
3) Annexin V Alexa Fluor 647
iii. Apoptotic caspase activation
1) CellEvent caspase-3/7 green—it is recommended to pre-load the cells with the dye for 1 h before an experiment.
2) Alternatively, one can utilize genetically encoded fluorescent protein-based caspase-3/7 reporters, such as VC3AI or ZipGFP.
Microscope setup
Both 488 and 496 nm lasers can be used for Cry2olig activation. During continuous whole-field photoactivation experiments, Cry2olig stimulation can be coupled with simultaneous imaging of fluorophores in the green channel [in this case, it is preferable to set up this channel as the last in an acquisition sequence to obtain the non-stimulated (t = 0) images]. However, this is not possible when using pulsed activation and/or single-cell targeting; in this case, only red or far-red fluorescent dyes and proteins should be used for the visualization of cell death and other processes of interest.
Set up the time-lapse imaging. Keep in mind that Cry2olig activation will cumulatively depend on both the laser power and the frame rate, so both need to be adjusted accordingly (e.g., if using higher frame rate, it is recommended to decrease the 488 nm laser power, while longer frame intervals might require increased laser intensity for Cry2olig activation). As an example, Figure 3 shows the relationship between the blue laser intensity and the percentage of pyroptotic cells detected after 30 min of illumination.
Figure 3. Relationship between opto-(h)caspase-1 activation and illumination parameters. (A) Quantification of pyroptotic (DRAQ7+) cells at 15 and 30 min post transient blue light illumination, and (B) at continuous repetitive (every 15 s) stimulation with blue light of various intensity. The data corresponds to Figure 3A and 3B in the original paper (Shkarina et al., 2022).
Acquire the time-lapse series. An example of the typical pyroptotic, necroptotic, and apoptotic cell morphology is shown in Figure 4. Note that the kinetics and morphological characteristics of each type of cell death vary among cell types and might also depend on the optoCDE construct expression level.
Figure 4. Assessment of optogenetically activated cell death effectors (optoCDE)-induced cell death using microscopy. (A) Schematic representation of expected morphological changes in cells undergoing different types of cell death (apoptosis, pyroptosis, and necroptosis) in response to light-induced optoCDE activation. (B) Representative morphological features of HEK293T cells expressing pyroptotic [opto-(h)caspase-1], apoptotic [opto-(h)caspase-8], or necroptotic [opto-(h)RIPK3] effectors and stimulated with blue light. Red: mCherry-tagged optoCDEs; green: CellTox Green; blue: Annexin V. Due to the differences in cell death kinetics, the after illumination time point refers to 30 min for pyroptotic, 60 min for apoptotic, and 90 min for necroptotic cells. The images are derived from the original paper (Shkarina et al., 2022). (C) Representative images of cells expressing genetically encoded caspase-3/7 reporter VC3AI (green) and undergoing light-induced apoptosis approximately 1 h after illumination.
Additionally, when doing this type of experiments for the first time, perform a similar experiment with the same cell type expressing Cry2olig alone and monitor the signs of cell death and/or abnormal changes in the cellular behavior related to the phototoxicity. If such changes are observed, decrease the laser power and/or the frame rate.
Optogenetic induction of pyroptosis in single cells
Set up the region of stimulation (ROI) corresponding to the specific cytoplasmic region in the cell to be targeted using ZEN “Bleaching” and “Regions” mode.
i. To account for the light diffusion and minimize the influence on the neighboring cells, the ROI should not exceed 5–10 μm2.
ii. The optimal laser intensity and number of scanning iterations varies depending on construct expression level, cell type, and type of cell death, and has to be determined empirically.
iii. If the signs of cell death (such as membrane blebbing, DRAQ7, or Annexin V positivity) are observed in the neighboring cells, reduce the ROI size and/or laser intensity or number of scanning iterations.
Set up the time-lapse experiment. This should include several frames before the photoactivation as the baseline, after which the ROI stimulation is performed, and additionally 30–60 min or more after.
Acquire the time lapse. To monitor for cell death, the imaging medium can be supplemented with the far-red cell death dyes (Annexin V or DRAQ7). The example of such experiment is shown at Figure 5.
Figure 5. Single-cell optogenetic induction of pyroptosis. (A) Representative time-lapse images of HaCaT cells, where the cytoplasmic region of a selected single cell (inset) is selectively photoactivated with blue light [blue square represents region of illumination (ROI)] at 0 min. Note the morphological changes and gain of DRAQ7 signal (turquoise) in targeted cell but not in the neighboring cells. (B) Quantification of DRAQ7 intensity in the nucleus of the photoactivated cell. Figure adapted from Shkarina et al. (2022).
If signs of optoCDE activation and cell death are also detected in the neighboring cells, repeat the experiment with the reduced laser power and/or number of pulses or decrease the ROI area.
Sub-lethal opto-caspase-1 activation
Set up the ROI stimulation experiment as described above. In this experiment, the ROI can include the whole field of view or be limited to the single cell.
Reduce the 488 nm laser power to 0.1%–0.2% and number of iterations to 1–3.
To monitor membrane permeabilization, supplement the imaging medium with DRAQ7. Use higher dye concentration (1:100–1:500) and laser power/gain at this stage to detect low amount of membrane damage.
Perform the imaging and photoactivation. The transient low-level optoCDE activation will likely lead to three phenotypes: 1) the cells that will undergo cell death (acquire “high” DRAQ7 staining); 2) the survivor cells (might acquire moderate DRAQ7 staining and transiently display some early features of cell death, such as membrane blebbing or nuclear condensation, but are able to revert to the normal morphology after), and 3) the cells that are not affected by the stimulation (not gaining DRAQ7 signal and no change in morphology) (Figure 6). The ratio of these cells in the population might depend on the optoCDE expression level, as well as additional cell-intrinsic factors (such as the variability in expression of the downstream effectors or activity of membrane repair systems).
Adjust the stimulation parameters to achieve the desired optoCDE activation and cell survival ratio and repeat the experiment.
Figure 6. Sub-lethal induction of pyroptosis using transient opto-(h)caspase-1 activation in human keratinocytes. The confluent monolayer of HaCaT cells was transiently stimulated with low-intensity (0.2 mW/cm2/pulse, three pulses) blue light at 3 min after the beginning of time lapse, and the data was acquired for 60 min. (A) Representative images of the whole illuminated population before (at 0 min) and after (60 min). DRAQ7 (turquoise) is a membrane-impermeable DNA-binding dye used to visualize pyroptotic membrane permeabilization. (B) Close-up images of cells displaying three types of fates: pyroptotic (top), sub-lethal (middle), and both pyroptotic and non-responding cells (bottom). Note the strong DRAQ7 signal and loss of cytoplasmic mCherry in pyroptotic cells and low DRAQ7 positivity in the sub-lethally activated cells.
Optogenetic induction of pyroptosis in human macrophage-like cell lines
Important points
In this protocol, we utilize U937, human monocyte-like cell line, which can be differentiated into the macrophage-like phenotype using PMA treatment. However, a similar type of experiments can be performed with other cell lines.
Prior to seeding, transgenic U937 lines are grown in suspension in T75 flasks in complete RPMI medium. For optimal growth, the medium is exchanged every 2–3 days. Avoid growing the cells to too high density, as this will lead to the reduction of cell viability and might impact the differentiation.
The light plate apparatus was manufactured, assembled, and calibrated as described previously (Gerhardt et al., 2016). The programming of the devices is performed using Iris (http://taborlab.github.io/Iris/).
To determine the optimal illumination parameters for each type of cell death and for each cell type, we recommend testing a range of light intensities and illumination duration. When doing the experiment for the first time, always include the wild-type cells and/or cells expressing Cry2olig alone to control for the phototoxicity.
This protocol describes the analysis of pyroptosis induction using opto-(h)caspase-1; however, a similar procedure can be used to activate other optoCDE constructs.
Step-by-step protocol
Cell seeding and differentiation
Collect the desired volume of U937 cells into the 50 mL flasks and centrifuge at 300× g at room temperature for 5 min. Discard the medium.
Resuspend the cells in fresh RPMI.
Count the cells and resuspend to the final density of 250,000 cells/mL.
Add PMA to the final concentration of 5 ng/mL.
Seed in 24-well black tissue culture–treated plates (4titude). Include additional wells for the non-illuminated cells and the positive (total lysis) control.
After 24 h, remove the PMA-containing medium, wash once with pre-warmed PBS, and add fresh growth medium.
After 48 h, replace the growth medium with the induction medium containing 2 μg/mL doxycycline and incubate cells overnight to induce optoCDE expression.
Illumination
Set up the light plate apparatus in the cell incubator (Figure 6B). If using transient illumination times, these experiments can be performed on the bench before moving the plates to 37 °C.
Generate the custom illumination program using IRIS software. The example of an illumination layout is shown in Figure 7C. We recommend using duplicate wells for each illumination condition and including additional wells for the non-illuminated controls and total cell lysis (cells treated with 1% Triton X-100).
Figure 7. Light plate apparatus setup. (A) Photo of the self-made light plate apparatus device used for the lactate dehydrogenase (LDH)/ELISA and western blot assays. (B) Setting up the device in the cell culture incubator for long-term experiments. (C) Example of the illumination layout used for determining the optimal illumination parameters.
Download the program and transfer the files to the micro-SD card. Perform a short test illumination round to ensure that the device works properly.
Remove the cell culture medium from the cells and add 300 μL of pre-warmed Opti-MEM or equivalent serum-free phenol red free medium per well.
Place the plate into the light plate apparatus, cover the lid, and start the illumination. Upon the end of the program, the lights should turn off automatically.
Remove the plate from the light plate apparatus. Add 30 μL of 10% Triton X-100 to the positive control (total cell lysis) wells. Using a P1000 pipette, pipette 5–10 times to lyse the cells.
Centrifuge the plate at 500× g for 5 min at room temperature to pellet the cell debris.
Gently collect the supernatant from each well to the pre-labeled 1.5 mL microtubes. Avoid touching the well bottom to prevent debris collection. At this stage, 100 μL of supernatant can be used for lactate dehydrogenase (LDH) measurement, while the rest can be preserved at -20 °C for the cytokine measurement.
Assessment of cell lysis using LDH activity assay
Prepare the appropriate volume of the LDH reaction mix.
Using a multichannel pipette, add 30 μL of reaction mix per well of a flat-bottom 96-well plate. Avoid creating bubbles, as this will impact the assay efficiency and reading.
In each well, add 30 μL of the cell supernatants collected in the step 2h. Include two to three technical replicates per each experimental well. Minimize delays while pipetting, as this might impact the difference in the colorimetric reaction between the wells and lead to bias.
Mix by gently tapping the plate on the side. Avoid mixing by pipetting, as this will generate bubbles. If bubbles are produced during pipetting, they can be manually removed before reading the plates using either syringe needles or an inverted Bunsen burner.
Incubate the plate for 15–20 min at room temperature, protected from light.
Add 30 μL of stop solution per well and gently mix.
Measure the absorbance using a 490–492 nm filter.
Assessment of the IL-1β release
IL-1β release quantification following illumination is typically quantified using ELISA, performed according to the manufacturer’s protocol (available at R&D website).
We recommend using several dilutions (1:1, 1:2, 1:5) of the collected supernatants, as the amount of IL-1β in undiluted supernatants frequently exceeds the dynamic range of assay.
Alternatively, a FRET-based no-wash homogeneous time resolved fluorescence assay can also be utilized for quicker single-step IL-1β quantification.
Western blot analysis of optoCDE activation and cell death
Before the start of the experiment, prepare the following reagents:
i. Pre-labeled 1.5 mL microtubes for supernatant and lysate collection.
ii. Cell lysis (RIPA or equivalent) and loading buffer.
iii. Reagents for phenol-chloroform protein precipitation.
iv. Reagents for the polyacrylamide gel preparation (or pre-cast gels).
v. Western blot running, transfer, and wash buffers.
Perform the illumination of the plates as described in step 5a. Ensure that the cells are in the good condition before the start of experiment and are mCherry-positive (this can be assessed using epifluorescent microscopy).
Centrifuge the plates at 300–500× g for 5 min at room temperature to pellet the dead cells and collect the supernatants from each well into 1.5 mL microcentrifuge tubes.
Immediately add 50–70 μL of hot (95 °C) loading buffer to the remaining adherent/pelleted cells. At this stage, plates can be processed immediately or sealed with parafilm and stored at -80 °C for later processing.
Pipette the lysis buffer up and down 5–10 times and collect the lysates into the microtubes. If lysates become too viscous at this stage, they can be additionally sonicated.
Incubate for 5 min at 95 °C.
In parallel, perform the protein precipitation in the cell supernatants using the phenol-chloroform protein extraction method (Demarco et al. 2022). After the precipitation, the dried protein pellets can be either resuspended in 1× lysis/loading buffer and processed independently or combined with lysates of corresponding wells.
Load samples on the 10%–12% polyacrylamide gels.
Perform the gel running and western blot analysis of the samples following the protocol available in the host lab. The detailed protocol for western blot analysis of such samples can also be found in Demarco et al. (2022).
For the detection of inflammatory opto-caspase activation, we recommend using the following primary antibodies: rabbit anti-cleaved IL-1β (83186, CST; 1:1,000), which detects IL-1β by activated opto-(h)caspase-1; mouse anti-IL-1β (12242, CST; 1:1,000), which detects full-length unprocessed IL-1β in non-activated cells; rabbit anti-GSDMD (ab210070; 1:1,000; Abcam), which detects non-activated GSDMD in resting cells; rabbit anti-cleaved N-terminal GSDMD (ab215203; 1:1,000; Abcam), which detects active GSDMD cleaved by opto-(h)caspase-1; mouse anti-caspase-1 (clone Bally-1 AG-20B-0048-C100; 1:1,000; AdipoGen); mouse anti-mCherry (ab125096; 1:2,000; Abcam); and HRP-conjugated mouse anti-tubulin (ab40742; 1:5,000; Abcam), used as a loading control. The secondary HRP-conjugated or fluorescently labeled antibodies can be used according to the lab’s choice. A representative blot is shown in Figure 8.
Figure 8. Example of the western blot analysis of opto-caspase-1-induced pyroptosis in U937 cells. Experiment was performed as described above and treatment with nigericin, an NLRP3 inflammasome activator, was used as a positive control to detect IL-1β and GSDMD processing. Note the cleavage of opto-caspase-1 (as detected by the disappearance of full-length opto-caspase-1 band and accumulation of Cry2olig-mCherry), IL-1β, and GSDMD upon illumination. Figure adapted from Shkarina et al. (2022).
Data analysis
Analysis of microscopy images
The analysis of the microscopy images is described in the Methods section of the original publication (Shkarina, et al., 2022). For quantification of different types of cell death, we assessed several parameters: a) appearance of the characteristic morphological changes associated with the each type of cell death (cell rounding and swelling, nuclear condensation for pyroptosis and necroptosis, nuclear fragmentation and persistent membrane blebbing for apoptosis); b) Annexin V staining, as an indicator of membrane scrambling and phosphatidylserine exposure (for all three types of cell death); and c) uptake of specific dyes associated with the loss of membrane integrity, such as CellTox Green or DRAQ7 (for pyroptosis and necroptosis). Additionally, activation of the apoptotic caspases can be monitored using genetically encoded or chemical fluorogenic caspase reporters (such as VC3AI, described in the original paper, or CellEvent caspase-3/7 reporter system). The number of dying cells can be normalized to the total number of cells per field of view, or, alternatively, to the number of mCherry-positive cells in the population, to account for the construct expression and transduction efficiency.
Analysis of LDH assay and ELISA results
LDH release from pyroptotic (or generally necrotic) cells following illumination is quantified using the following equation:
(LDHsample - LDHnegative control)/(LDHpositive control - LDHnegative control) × 100
Where negative control is assay medium (e.g., Opti-MEM or other phenol red free medium) and positive control corresponds to the cells lysed with the 1% Triton X-100.
Quantification of IL-1β release is performed according to the manufacturer’s protocol. Note that the variation in initial (pre-illumination) cell density between the cell lines or conditions can have a strong effect on the amount of detected IL-1β. This can be corrected by normalizing IL-1β values to the ratio of maximum LDH values from lysed cells. An example of the LDH and IL-1β secretion data obtained from this type of experiments is shown in Figure 9.
Figure 9. Example of lactate dehydrogenase (LDH) release and IL-1β secretion upon illumination-induced opto-(h)caspase-1 activation. Nigericin (NLRP3 inflammasome activator) was used to monitor cell competency to endogenous inflammasome activation. Figure adapted from Shkarina et al. (2022).
Notes
General notes
When inducing different modes of cell death, it is important to take into consideration the selection of the appropriate cell line/type. We observed that the ability of cells to undergo different forms of cell death upon optoCDE activation varies among the different cell lines and cell types tested. This might depend on several factors:
Effector efficiency: we observed that, due to the differences in substrate processing kinetics and efficiency between different caspases, and also kinetic and mechanistic differences between separate cell death modalities, it might be necessary to adjust the levels of construct expression or stimulation to achieve similar cell death levels. As an example, in some cell lines, achieving an equal level of apoptosis might require higher expression of more prolonged stimulation for opto-caspase-8 than opto-caspase-9. Also, the same is applicable for opto-caspase-4 vs. opto-caspase-1 vs. opto-caspase-5, and for optoRIP3 vs. optoMLKL (although the last case can be explained by MLKL being the most distal cell death effector in necroptosis, while RIP3-induced necroptosis would be initiated more proximately and constrained by both endogenous MLKL levels and post-translational regulation).
Expression of downstream effectors, such as GSDMD for inflammatory (pyroptotic) caspases or MLKL for optoRIPK3. These proteins can be co-expressed with optoCDEs to achieve pyroptosis in the cell lines that are either naturally deficient (such as HEK cells) or express low levels (HeLa) of them endogenously.
Downstream regulatory mechanisms (such as phosphorylation or other types of post-translational modifications or protein–protein interactions), which might limit or promote optoCDE activation in some cell types.
Spontaneous or unwanted optoCDE activation due to overexpression or low-level activation due to visible light exposure might induce cytotoxicity or lead to the negative selection of expressing cells. Thus, it is essential to always protect the cells from visible light following the induction of optoCDE construct expression, or when using constitutive expression systems. All necessary cell handling, such as treatments or medium exchange, should be performed under dim light conditions, or using lab space and hoods equipped with red-light sources (lamps or LED strips). Additionally, we strongly recommend using inducible expression systems (such as Tet-ON) and tetracycline-free serum for stable cell line generation and maintenance.
Also, avoid checking the expressing cell lines under the microscope using violet, blue, or transmitted (white) light before the beginning of the experiment, as this will trigger the optoCDE construct activation. Usually, the mCherry expression in stable lines can be used as a good proxy measurement of the viability in unstimulated cells. Alternatively, visual assessment of cell density and morphology can be performed in separate wells, which can then be excluded from the experiment.
We recommend determining the optimal illumination parameters for each optoCDE construct and each cell line by testing several different blue light intensities and/or illumination duration. Additionally, it is essential to include wild-type cells and cells expressing Cry2olig alone to control for phototoxicity and potential non-specific effects of Cry2olig overexpression and photoactivation.
Acknowledgments
This work was supported by grants from the ERC (ERC2017-CoG-770988-InflamCellDeath), the Swiss National Science Foundation (175576 and 198005), the OPO Stiftung and Novartis to P.B. K.S. is a recipient of SNSF Postdoc.Mobility fellowship (P500PB_211096). This protocol was adapted from Shkarina et al. (2022).
Competing interests
The authors declare no competing interests.
References
Bedoui, S, Herold, M. J. and Strasser, A. (2020). Emerging connectivity of programmed cell death pathways and its physiological implications. Nat Rev Mol Cell Bio 21: 678–695.
Demarco, B., Ramos, S. and Broz, P. (2022). Detection of Gasdermin Activation and Lytic Cell Death During Pyroptosis and Apoptosis. In: Kufer, T.A., Kaparakis-Liaskos, M. (Eds.). Effector-Triggered Immunity. Methods in Molecular Biology. Humana Press, New York.
Galluzzi, L., Vitale, I., Aaronson, S. A., Abrams, J. M., Adam, D., Agostinis, P., Alnemri, E. S., Altucci, L., Amelio, I., Andrews, D. W., et al. (2018). Molecular mechanisms of cell death: recommendations of the Nomenclature Committee on Cell Death 2018. Cell Death Differ 25(3): 486–541.
Gerhardt, K. P., Olson, E. J., Castillo-Hair, S. M., Hartsough, L. A., Landry, B. P., Ekness, F., Yokoo, R., Gomez, E. J., Ramakrishnan, P., Suh, J., et al. (2016). An open-hardware platform for optogenetics and photobiology. Sci. Rep 6(1): e1038/srep35363.
Oberst, A., Pop, C., Tremblay, A. G., Blais, V., Denault, J. B., Salvesen, G. S. and Green, D. R. (2010). Inducible Dimerization and Inducible Cleavage Reveal a Requirement for Both Processes in Caspase-8 Activation. J. Biol. Chem 285(22): 16632–16642.
Qi, Y. B., Garren, E. J., Shu, X., Tsien, R. Y. and Jin, Y. (2012). Photo-inducible cell ablation in Caenorhabditis elegans using the genetically encoded singlet oxygen generating protein miniSOG. Proc. Natl. Acad. Sci. U.S.A. 109(19): 7499–7504.
Shkarina, K., Hasel de Carvalho, E., Santos, J. C., Ramos, S., Leptin, M. and Broz, P. (2022). Optogenetic activators of apoptosis, necroptosis, and pyroptosis. J. Cell Biol 221(6): e202109038.
Taslimi, A., Vrana, J. D., Chen, D., Borinskaya, S., Mayer, B. J., Kennedy, M. J. and Tucker, C. L. (2014). An optimized optogenetic clustering tool for probing protein interaction and function. Nat. Commun 5(1): e1038/ncomms5925.
Tirlapur, U. K., Konig, K., Peuckert, C., Krieg, R. and Halbhuber, K. J. (2001). Femtosecond near-infrared laser pulses elicit generation of reactive oxygen species in mammalian cells leading to apoptosis-like death. Exp Cell Res 263(1): 88–97.
Wu, X. N., Yang, Z. H., Wang, X. K., Zhang, Y., Wan, H., Song, Y., Chen, X., Shao, J. and Han, J. (2014).
Distinct roles of RIP1–RIP3 hetero- and RIP3–RIP3 homo-interaction in mediating necroptosis.Cell Death Differ 21(11): 1709–1720.
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Large-scale Purification of Type III Toxin-antitoxin Ribonucleoprotein Complex and its Components from Escherichia coli for Biophysical Studies
PM Parthasarathy Manikandan *
KN Kavyashree Nadig *
MS Mahavir Singh
(*contributed equally to this work)
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4763 Views: 415
Reviewed by: Alba BlesaValentine V Trotter Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nucleic Acids Research Feb 2022
Abstract
Toxin–antitoxin (TA) systems are widespread bacterial immune systems that confer protection against various environmental stresses. TA systems have been classified into eight types (I–VIII) based on the nature and mechanism of action of the antitoxin. Type III TA systems consist of a noncoding RNA antitoxin and a protein toxin, forming a ribonucleoprotein (RNP) TA complex that plays crucial roles in phage defence in bacteria. Type III TA systems are present in the human gut microbiome and several pathogenic bacteria and, therefore, could be exploited for a novel antibacterial strategy. Due to the inherent toxicity of the toxin for E. coli, it is challenging to overexpress and purify free toxins from E. coli expression systems. Therefore, protein toxin is typically co-expressed and co-purified with antitoxin RNA as an RNP complex from E. coli for structural and biophysical studies. Here, we have optimized the co-expression and purification method for ToxIN type III TA complexes from E. coli that results in the purification of TA RNP complex and, often, free antitoxin RNA and free active toxin in quantities required for the biophysical and structural studies. This protocol can also be adapted to purify isotopically labelled (e.g., uniformly 15N- or 13C-labelled) free toxin proteins, free antitoxin RNAs, and TA RNPs, which can be studied using multidimensional nuclear magnetic resonance (NMR) spectroscopy methods.
Key features
• Detailed protocol for the large-scale purification of ToxIN type III toxin–antitoxin complexes from E. coli.
• The optimized protocol results in obtaining milligrams of TA RNP complex, free toxin, and free antitoxin RNA.
• Commercially available plasmid vectors and chemicals are used to complete the protocol in five days after obtaining the required DNA clones.
• The purified TA complex, toxin protein, and antitoxin RNA are used for biophysical experiments such as NMR, ITC, and X-ray crystallography.
Graphical overview
Keywords: Toxin–antitoxin system ToxIN RNA–protein complex Type III TA system Co-purification
Background
Type III toxin–antitoxin (TA) systems are genetic modules found in several bacteria, consisting of a protein toxin and a noncoding RNA antitoxin (Fineran et al., 2009). Antitoxin RNA repeats directly bind to the protein toxin, inhibiting its activity by forming a TA ribonucleoprotein (RNP) complex under homeostatic conditions. The primary functions of type III TA systems are attributed to the phage inhibition mechanism in bacteria (Song and Wood, 2020; LeRoux and Laub, 2022; Lin et al., 2023). Type III TA systems are widespread and present in several pathogenic bacteria, including Staphylococcus aureus, Yersinia pseudotuberculosis, and Fusobacterium nucleatum (Blower et al., 2012), and several bacteria in the gut microbiome. Thus, understanding the structure and assembly of type III TA systems could help in devising novel antibacterial strategies by targeting these systems to free the toxins in the pathogenic bacteria, resulting in bacterial growth arrest or death.
The type III TA systems are classified into three families: ToxIN, CptIN, and TenpIN, based on toxin sequence homology (Blower et al., 2012). Among these classes, the ToxIN family (here, protein toxin and antitoxin RNA are called ToxN and ToxI, respectively) is the most studied in terms of its structure, assembly, and functions (Blower et al., 2011; Short et al., 2012; Guegler and Laub, 2021; Manikandan et al., 2022). In a recent study, we classified E. coli ToxIN TA systems into five clusters based on the sequence analysis of the toxin proteins (Manikandan et al., 2022). The structural and functional analysis of a CptIN TA complex from Eubacterium rectale has also been reported (Rao et al., 2015).
It is often difficult to purify free toxins from E. coli due to their inherent toxicity to the bacteria. To circumvent this issue, toxin and antitoxin are often co-expressed and co-purified as non-toxic TA complexes. This is followed by denaturing the TA complex (e.g., in the presence of 6 M guanidinium hydrochloride or 8 M urea) to separate the toxin and antitoxin. The free denatured toxin can then be refolded back to its natural form. However, in vitro refolding of proteins is not always successful. Alternatively, in some cases, active-site toxin mutants have been generated, which can be expressed in E. coli and purified (Samson et al., 2013). While several experiments can be performed using the catalytic mutant protein, a wild-type active protein is often still required to understand the toxins’ mechanism of action.
Here, by adapting the methods published earlier (Blower et al., 2011; Short et al., 2012; Rao et al., 2015; Manikandan et al., 2022), we have optimized a protocol to co-express and co-purify the ToxIN complexes and their components from three clusters of ToxIN family from E. coli (Manikandan et al., 2022) for their biophysical and structural investigation [using X-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy]. The method described here yields TA RNP complex, free protein toxin, and free antitoxin RNA of the ToxIN system with an approximate molar ratio of 3:2:2 (with an approximate yield of 7, 3, and 1.5 mg, respectively, from one litre of LB media). The availability of a robust purification method could potentially break the barrier between the discovery of new type III TA systems and their subsequent biophysical characterization to understand their mechanism of action.
Materials and reagents
Biological materials
E. coli BL21(DE3) competent cells (prepared in the lab following a standard protocol) (Sambrook and Russell, 2006)
E. coli DH5α competent cells (prepared in the lab following a standard protocol)
pColdTM II vector (Takara Bio, catalog number: 3362)
pET-21a(+) (Novagen, catalog number: 69740-3)
pETDuetTM-1 vector (Novagen, catalog number: 71146-3)
pRSFDuetTM-1 vector (Novagen, catalog number: 71341-3)
Reagents, chemicals, and kits
Acetic acid (glacial) (Sisco Research Laboratories, catalog number: 85801, CAS number: 64-19-7)
Acrylamide:Bis-acrylamide (19:1) for electrophoresis, 40% solution (Fischer BioReagentsTM, catalog number: BP1406-1, CAS number: 79-06-1), store at 4–8 °C
Agarose low EEO (Sisco Research Laboratories, catalog number: 36601, CAS number: 9012-36-6)
Ammonium persulfate (APS) (Sisco Research Laboratories, catalog number: 65553, CAS number: 7727-54-0)
Ampicillin (Sisco Research Laboratories, catalog number: 61314, CAS number: 69-52-3), store at 2–8 °C
Bradford reagent (Sigma, catalog number: B6916-500 mL), store at 2–8 °C
Bromophenol Blue indicator (Sisco Research Laboratories, catalog number: 11458, CAS number: 115-39-9)
Coomassie brilliant blue (Sisco Research Laboratories, catalog number: 93473, CAS number: 6104-58-1)
D/L-Dithiothreitol (DTT) (Sisco Research Laboratories, catalog number: 17315, CAS number: 3483-12-3), store at 0–4 °C
dNTPs (New England Biolabs, catalog number: N0447S), store at -20 °C
DpnI (New England Biolabs, catalog number: R0176), store at -20 °C
Ethanol (Changshu Hongsheng Fine Chemicals, analytical grade, UN No: 1170)
Ethylenediaminetetraacetic acid (EDTA) (Sisco Research Laboratories, catalog number: 50027, CAS number: 6381-92-6)
Formamide (Sisco Research Laboratories, catalog number: 71714 (062930), CAS number: 75-12-7)
Gel extraction kit (QIAquick®, Qiagen, catalog number: 28704)
Glycerol (Sisco Research Laboratories, catalog number: 62417, CAS number: 56-81-5)
Glycine (Sisco Research Laboratories, catalog number: 64072, CAS number: 56-40-6)
Hydrochloric acid (Sisco Research Laboratories, catalog number: 65955, CAS number: 7647-01-0)
Imidazole (Sisco Research Laboratories, catalog number: 61510-500G)
Isopropyl β-d-1-thiogalactopyranoside (IPTG) (Sisco Research Laboratories, catalog number: 67208, CAS number: 367-93-1), store at 0–4 °C
Kanamycin (Sisco Research Laboratories, catalog number: 99311, CAS number: 25389-94-0), store at 2–8 °C
Luria Bertani agar (LA), Miller (Himedia, catalog number: M1151-500G)
Luria broth (LB) (Himedia, catalog number: M575-500G)
Methanol for HPLC (SD Fine Chemical Limited, catalog number: 25217 L25)
NaCl (Sisco Research Laboratories, catalog number: 41721, CAS number: 7647-14-5)
NcoI (New England Biolabs, catalog number: R3193S), store at -20 °C
NdeI (New England Biolabs, catalog number: R0111S), store at -20 °C
PCR purification kit (Qiagen, QIAquick® PCR purification kit, catalog number: 28104)
Phusion High-Fidelity DNA polymerase (New England Biolabs, catalog number: M0530S), store at -20 °C
Plasmid extraction kit (QIAprep® spin Miniprep kit, Qiagen, catalog number: 27106)
Protease inhibitor cocktail (Roche, catalog number: 04693159001), store at 2–8 °C
Quick calf intestinal phosphatase (CIP) (New England Biolabs, catalog number: M0525S), store at -20 °C
Sodium dodecyl sulphate (SDS) (Sisco Research Laboratories, catalog number: 32096, CAS number: 151-21-3)
T4 DNA ligase (New England Biolabs, catalog number: M0202S), store at -20 °C
Tetramethylethylenediamine (TEMED) (Spectrochem private limited, catalog number: 012017), store at 2–8 °C
Toluidine Blue (Sisco Research Laboratories, catalog number: 22134, CAS number: 92-32-9)
Tris (Sisco Research Laboratories, catalog number: 71033-500G)
Urea (Sisco Research Laboratories, catalog number: 21113, CAS number: 57-13-6)
XhoI (New England Biolabs, catalog number: R0145S), store at -20 °C
β-Mercaptoethanol (Sisco Research Laboratories, catalog number: 83759, CAS number: 60-24-2)
Plastic and other materials
Centrifuge bottle 500 mL (Tarsons, catalog number: 544020)
Centrifuge tubes 15 mL (Tarsons, SpinwinTM Tube Conical bottom, catalog number: 520060)
Centrifuge tubes 50 mL (Tarsons, catalog number: 520061)
Membrane filters (0.22 μm) (Merck Life Science private limited, catalog number: GVWP04700)
Membrane filters (0.45 μm) (Merck Life Science private limited, catalog number: HVLP04700)
Microcentrifuge tubes 1.5 mL (Tarsons, catalog number: 500010)
Microcentrifuge tubes 2 mL (Tarsons, catalog number: 500020)
Oakridge centrifuge tubes (50 mL) (Thermo Scientific, catalog number: 3115-0050)
PCR tubes 0.2 mL flat cap (Tarsons, catalog number: 510051)
SnakeSkin dialysis bag 3.5 kDa cut off (dialysis bag) (Thermo Scientific, catalog number: 88242), store at 2–8 °C
Syringe filters (0.22 μm) (Sartorius Minisart®, catalog number: S6534-FMOSK)
Syringe filters (0.45 μm) (Sartorius Minisart®, catalog number: S6555-FMOSK)
Syringes 10 mL (Hindustan Syringes and Medical Devices Limited, Dispo Van, India)
Syringes 20 mL (Hindustan Syringes and Medical Devices Limited, Dispo Van, India)
Solutions
PCR reaction mixture for all the amplifications (see Recipes)
Double digestion of vector and insert using restriction enzymes (see Recipes)
Ligation reaction of vector and insert (see Recipes)
Lysis buffer (see Recipes)
Wash buffer (see Recipes)
Elution buffer (see Recipes)
Final wash buffer (see Recipes)
Dialysis buffer (see Recipes)
Ion exchange chromatography buffers (see Recipes)
Size exclusion chromatography buffer (see Recipes)
Separating gel for 12% SDS-PAGE (see Recipes)
Stacking gel for SDS-PAGE (see Recipes)
5× loading dye for SDS-PAGE (see Recipes)
Staining solution for SDS-PAGE (see Recipes)
10× running buffer for SDS-PAGE (see Recipes)
SDS-PAGE destaining solution (see Recipes)
Urea-acrylamide gel (see Recipes)
Formamide dye (see Recipes)
SDS-loading dye (5×) (see Recipes)
Tris-borate-EDTA (TBE) buffer (see Recipes)
Recipes
PCR reaction mixture for all the amplifications
Reaction component Volume (μL) (per 20 μL reaction)
5× Phusion HF buffer 4
10 mM dNTPs 0.4
10 μM forward primer 1
10 μM reverse primer 1
Template DNA (100 ng/μL) 1
Autoclaved MilliQ H2O 12.4
Phusion® High-Fidelity DNA Polymerase 0.2
Total volume 20
Double digestion of vector and insert using restriction enzymes
Reaction component Volume (μL) (per 50 μL reaction)
10× Cutsmart buffer 5
Enzyme 1 1
Enzyme 2 1
Vector/insert DNA 5 μg/2 μg
Autoclaved MilliQ H2O make up to 50 μL
Total volume 50
Ligation reaction of vector and insert
Reaction component Volume (μL) (per 20 μL reaction)
10× ligase buffer 2
Digested purified vector (10 ng/μL)
Insert (10 ng/μL)
5
5
T4 DNA ligase
Autoclaved MilliQ H2O
1
7
Total volume 20
Lysis buffer
Reagent Final concentration Amount
NaCl (4 M) 300 mM 37.5 mL
Tris-HCl (1 M, pH 7.5)
Imidazole (5 M, pH 8.0)
Glycerol (100%)
β-mercaptoethanol (14.28 M)
50 mM
10 mM
10%
2 mM
25 mL
1 mL
50 mL
70 μL
MilliQ H2O n/a 386.43 mL
Total n/a 500 mL
Wash buffer
Reagent Final concentration Amount
Lysis buffer n/a 49.9 mL
Imidazole (5 M, pH 8.0) 20 mM 100 μL
Total n/a 50 mL
Elution buffer
Reagent Final concentration Amount
Lysis buffer n/a 24 mL
Imidazole (5 M, pH 8.0) 200 mM 1 mL
Total n/a 25 mL
Final wash buffer
Reagent Final concentration Amount
Lysis buffer n/a 18 mL
Imidazole (5 M, pH 8.0) 500 mM 2 mL
Total n/a 20 mL
Dialysis buffer
Reagent Final concentration Amount
NaCl (4 M) 50 mM 25 mL
Tris-HCl (1 M, pH 7.5) 50 mM 100 mL
DTT 1 mM 308.48 mg
MilliQ H2O n/a 1,875 mL
Total n/a 2 L
Ion-exchange chromatography buffers
Buffer A: same as dialysis buffer.
Buffer B:
Reagent Final concentration Amount
NaCl (4 M) 1 M 125 mL
Tris-HCl (1 M, pH 7.5) 50 mM 25 mL
DTT 1 mM 77.12 mg
MilliQ H2O n/a 350 mL
Total n/a 500 mL
Size-exclusion chromatography buffer
Reagent Final concentration Amount
NaCl (4 M) 50 mM 6.25 mL
Tris-HCl (1 M, pH 7.5) 20 mM 10 mL
DTT 1 mM 77.12 mg
MilliQ H2O n/a 483.75 mL
Total n/a 500 mL
Separating gel for 12% SDS-PAGE (prepare solution volume depending on the gel cast size)
Reagent Amount
Acrylamide:Bis-acrylamide (19:1) (40% solution) 2.4 mL
1.5 M Tris, pH = 8.8 2 mL
10% SDS solution 80 μL
10% APS solution 80 μL
TEMED 8 μL
MilliQ H2O 3.432 mL
Total 8 mL
Stacking gel for SDS-PAGE (prepare solution volume depending on the gel cast size)
Reagent Amount
Acrylamide:Bis-acrylamide (19:1) (40% solution) 0.75 mL
1.5 M Tris, pH = 6.8 1.25 mL
10% SDS solution 50 μL
10% APS solution 50 μL
TEMED 5 μL
MilliQ H2O 2.9 mL
Total 5 mL
5× loading dye for SDS-PAGE
Reagent Amount
β-Mercaptoethanol 5%
Bromophenol Blue 0.02%
Glycerol 30%
SDS 10%
Tris-Cl (pH = 6.8) 250 mM
Staining solution for SDS-PAGE
Reagent Amount
Coomassie brilliant blue 1 g
Methanol 400 mL
Acetic acid 100 mL
MilliQ H2O 500 mL
Total 1 L
10× running buffer for SDS-PAGE
Reagent Amount
Tris base 30 g
Glycine 144 g
SDS 10 g
MilliQ H2O Make up to 1L
Total 1 L
SDS-PAGE destaining solution
Reagent Amount
Methanol 40 mL
Acetic acid (glacial) 10 mL
MilliQ H2O 50 mL
Total 100 mL
Urea-acrylamide gel (prepare solution volume depending on the gel cast size)
Reagent Concentration
Acrylamide:Bis-acrylamide (19:1) 15%
Urea 8 M
TBE buffer 1×
APS 1%
TEMED 0.1%
MilliQ H2O Make up the volume as per requirement
Formamide dye
Prepared using a CSHL protocol for formamide gel-loading buffer (Cold Spring Harb Protoc, 2013)
SDS-loading dye (5×)
Prepared using a CSHL protocol (Cold Spring Harb Protoc, 2008)
Tris-borate-EDTA (TBE) buffer
Prepared according to a CSHL protocol (Cold Spring Harb Protoc, 2010)
Equipment
Amicon Ultra-15 centrifugal filter unit, 3.5 kDa cutoff (Millipore Sigma, catalog number: UFC9030)
Anion-exchange column (HiTrapTM, Q FF, 5 mL)
Centrifuge (Kubota, Model-6500, serial number: K60115-G000)
Centrifuge for 50 and 1 mL tubes (Eppendorf, model: 5804R)
Dry bath (Bionova, model: SLM-DB-120)
Electrophoresis equipment (BIOBEE® Tech)
Electrophoresis power supply (Bio-Rad, PowerPacTM Basic, 041BR178399)
Fast protein liquid chromatography (FPLC), with fraction collector (GE Healthcare, ÄKTA prime plus)
Laminar airflow (local make)
Mastercycler (Eppendorf Flexi lid Nexus, 6333, serial number: 6333DQ408627)
Mixer (Eppendorf Mixmate 22331 Hamburg, serial number: 5353DN316975)
NanoDrop (Thermo Scientific, NanoDrop 2000c spectrophotometer)
Ni2+-NTA column (GE Healthcare HisTrapTM HP, 5 mL)
Peristaltic pump (Bio-Rad, Econo gradient pump, serial number: 491BR 1897, catalog number: 731-9002)
pH meter (Thermo ScientificTM Orion StarTM A111)
Rotor (Kubota, 6 × 50 mL- AG-506R)
Rotor (Kubota, 6 × 500 mL- AG-5006A)
S200 column (HiLoadTM 16/600 superdexTM 200pg, ID-0059)
SDS-PAGE setup (Invitrogen, Minigel tank)
Shaker and incubator (BioTek, NB-205VQ)
Sonicator (LABMAN Scientific Instruments, model: pro650)
Spectrophotometer (Eppendorf Biophotometer D30, serial number: 6133D0400689)
Tabletop centrifuge (Eppendorf centrifuge 5418, FA-45-18-11 S/N:20881)
Vacuum pump (Tarsons, Rockyvac 400)
Water filtration source (Millipore, Sigma)
Weighing balance, milligram sensitivity (Sartorius, BSA 623S-CW)
Procedure
Molecular cloning of type III antitoxin RNA
Perform the DNA synthesis and cloning of the identified type III TA operon into a high copy number E. coli plasmid vector through a gene synthesis service provider. Sub-clone the antitoxin repeats and its natural promoter into pRSFDuet-1 vector using standard cloning protocols described below (Figure 1A and Supplementary Table 1).
Design forward and reverse primers to amplify the antitoxin region along with the natural promoter and terminator, with addition of restriction enzyme sites NcoI and XhoI at the 5′ and 3′ ends of the insert. PCR amplify the insert from the plasmid containing type III TA operon, using Phusion DNA polymerase (a PCR reaction setup is indicated in Recipe 1 and Table 1).
Table 1. Thermocycling conditions for the PCR reaction
Step Temperature (°C) Duration No. of cycles
Initial denaturation 98 30 s 1
Denaturation 98 10 s 30
Annealing Ta 30 s
Extension 72 30 s per kb
Final extension 72 10 min 1
Hold 4 ∞ -
Purify the PCR-amplified insert using the PCR purification kit following the manufacturer’s protocol and elute DNA using nuclease-free water.
Digest ~1–2 μg of the PCR purified insert using the restriction enzymes NcoI and XhoI by incubating at 37 °C for 2 h by setting up a reaction as described in Recipe 2.
Digest ~5 μg of the pRSFDuet-1 vector using the same restriction enzymes in a 50 μL reaction. After digesting the vector, incubate the reaction mixture at 80 °C for 20 min to heat-inactivate the restriction enzymes. Treat the mixture with 0.3 μL of calf intestinal phosphatase (CIP) at 37 °C for 1 h to prevent vector self-ligation.
Purify the digested insert and vector using 1% agarose gel electrophoresis. Visualize the digested insert and vector bands from the ethidium bromide–stained gel by UV irradiation at 254 nm. Excise the gel bands containing the vector and insert and isolate the respective DNA fragments using a gel extraction kit following manufacturer’s protocol; elute DNAs using nuclease-free water.
Use 100 ng of each digested vector and insert DNAs (~1:7 ratio) to set up a 20 μL ligation reaction as described in Recipe 3. Incubate the reaction mixture at room temperature for 1 h followed by incubation at 4 °C for 5–12 h.
Transform 10 μL of the ligation reaction mixture in E. coli DH5α competent cells and plate them in LB agar plate containing kanamycin (0.05 mg/mL) antibiotic. A control ligation reaction and transformation could be performed without the insert DNA in the reaction mixture. The control reaction should result in significantly fewer (~10-fold) colonies than the ligation with the insert.
Isolate plasmid DNA from the colonies obtained upon transformation using a plasmid isolation kit and confirm the positive clones by sequencing.
Molecular cloning of type III toxin protein
Design forward and reverse primers to clone the type III toxin into an E. coli expression vector under an inducible promoter with a hexahistidine tag. The vector must be compatible for co-transformation with the pRSFDuet-1 vector with a different antibiotic resistance and a different origin of replication (e.g., pCold II, pRSFDuet-1, pET21a(+), etc.) In our study, we cloned the type III toxin into a pCold II vector under a cold-shock inducible promoter with an N-terminal hexahistidine tag between the restriction enzyme sites NdeI and XbaI.
Obtain the ligated plasmid containing toxin insert in the vector of interest by following cloning steps A2–A7 of antitoxin cloning using the toxin insert and appropriate vector DNA.
Co-transform 10 μL of the ligation mixture along with 100 ng of the antitoxin DNA containing plasmid in E. coli DH5α competent cells and plate on an LB agar plate containing both kanamycin (0.05 mg/mL) and ampicillin (0.1 mg/mL) antibiotics.
Isolate the plasmid DNA mixture (toxin and antitoxin) from the colonies obtained and confirm the positive toxin clones by sequencing using primers specific to the toxin cloning site. The obtained plasmid DNA mixture will be used further to transform E. coli cells to express and purify the type III TA complex. In our study, the plasmid DNA mixture contains antitoxin and toxin cloned in vectors, as shown in Supplementary Table 1.
Figure 1. A general strategy for cloning type III toxins in expression vectors.
(A) Schematic of cloning of antitoxin ToxI RNA in an expression vector. The antitoxin DNA was cloned along with its natural constitutive promoter in pRSFDuet-1 vector. (B) Schematic of cloning of toxin ToxN protein in an expression vector. The toxin DNA was cloned with an N-terminal hexahistidine tag in an IPTG inducible vector such as pCold II. (C) No colonies were obtained upon transformation of plasmids containing only toxin. (D) Positive colonies were obtained upon co-transforming plasmids containing toxin and antitoxin in different compatible vectors.
Transformation and inoculation
Place a vial of E. coli BL21 DE3 competent cells from -80 °C stock on ice to thaw for 15 min.
Add ~100 ng of the plasmid mixture (toxin and antitoxin) and incubate on ice for 20 min.
Provide heat shock by placing the vial in a dry bath at 42 °C for 45 s and immediately transfer the vial on ice and incubate for 5 min.
Add 250 μL of LB medium to the vial in a laminar airflow cabinet and incubate for 1 h in a shaker incubator maintained at 37 °C at 180 rpm.
Spread plate by using 150 μL of this culture on an agar plate containing double antibiotic [corresponding to the plasmid combination chosen; here, we used ampicillin (0.1 mg/mL) and kanamycin (0.05 mg/mL)] and incubate for 12–14 h to obtain the transformed colonies. We did not observe bacterial colonies post-transformation of only toxin plasmid (Figure 1C). However, the co-transformation of toxin and antitoxin plasmids resulted in bacterial colonies selected using dual antibiotics (Figure 1D).
Inoculate 100 mL of LB medium with both antibiotics with a single colony from the transformed plate. Incubate overnight at 37 °C at 180 rpm in a shaker incubator.
Depending on the plasmid in which the toxin and antitoxin are cloned (Table 1), one of the following steps can be followed:
Toxin cloned in pCold II: inoculate 10 mL of the overnight primary culture (in LB media) into a larger secondary culture (1 L of LB media) and incubate until OD600 reaches ~0.8–1.0. Incubate the culture at 15 °C without shaking for 30 min and induce by adding IPTG to a final concentration of 1 mM. Incubate at 180 rpm for 24 h at 15 °C for complex expression. Harvest the cells by centrifugation at 6,800× g for 15 min.
Toxin cloned in pET-21a(+): inoculate 10 mL of the overnight primary culture (in LB media) into a larger secondary culture (1 L of LB media) and incubate until an OD600 ~0.6–0.8. Induce by adding IPTG to a final concentration of 1 mM. Incubate the culture at 180 rpm for 4–5 h at 37 °C for complex expression. Harvest the cells by centrifugation at 6,800× g for 15 min.
Stop point: The cell pellet can be stored at -20 °C for up to a month before processing.
Cell lysis and Ni-NTA affinity chromatography
Resuspend the cells in lysis buffer (see Recipes) and add a tablet of protease inhibitor cocktail.
Lyse the cells by sonication with a pulse of 3 s on and 6 s off and an amplitude of 32%.
Centrifuge the lysed cells at 18,328× g for 45 min at 4 °C in the Oakridge centrifuge tubes.
Equilibrate Ni2+-NTA column connected to a peristaltic pump with lysis buffer. Filter the supernatant using a 0.45 μm syringe filter and load it onto the column at a 1.5 mL/min flow rate. After loading, wash the column with 50 mL of wash buffer (see Recipes). Elute the complex using elution buffer.
Collect three fractions: E1, the first 2 mL; E2, the next 15 mL; and E3, the final 5 mL. Store at 4 °C.
Wash the column with the final wash buffer to remove any bound protein to the column. Pass 25 mL of Milli-Q water and store the Ni2+-NTA column in 20% ethanol.
PAGE analysis of toxin and antitoxin: to analyse the flowthrough, wash, and elution fractions on 12% SDS-PAGE for the presence of toxin protein and on 15% Urea-PAGE for the presence of RNA antitoxin. Visualize the protein in SDS-PAGE gels with Coomassie brilliant blue staining solution followed by destaining using destaining solution (see Recipes). The RNA can be visualized by staining the urea-PAGE gels with 0.25% toluidine blue solution followed by destaining using water.
Dialysis
Collect all the elution fractions that contain the TA components for dialysis (usually, E2 has the TA components).
Dialysis: wash an appropriate length (which can hold the elution fraction volume containing the TA component) of the 6 M dialysis bag with a cutoff of 3.5 kDa with MilliQ H2O and equilibrate with dialysis buffer.
Carefully add the elution fractions into the bag and dialyse at 4 °C for 4–5 h.
Change the dialysis buffer and continue to dialyse for 12–14 h.
Ion-exchange chromatography
Equilibrate the anion-exchange column with 25 mL of ion-exchange buffer A.
Filter the dialysed complex using a 0.45 μm syringe filter and load it onto the anion-exchange column at a 1 mL/min flow rate connected to ÄKTA prime plus FPLC system. After loading, wash the column with 20 mL of ion-exchange buffer A.
Elute the complex by increasing the gradient of NaCl from 50 to 1,000 mM (using buffer A and buffer B), over a volume of 100 mL. This should yield separate fractions of ToxN protein (~0%–20% buffer B), TA RNP complex (~45%–55% buffer B), and antitoxin ToxI RNA repeat (~55%–75% buffer B) (Figure 2A), which can be confirmed by PAGE analysis of protein and RNA components.
Figure 2. Expression and purification of type III toxin–antitoxin (TA) components from ToxINEc-C1. (A) Anion-exchange chromatography profile of ToxINEc-C1 complex shows the purification of individual toxin, antitoxin, and complex components of ToxINEc-C1. (B–D) Size exclusion chromatography (SEC) profiles of ToxIEc-C1 RNA (B), ToxNEc-C1 protein (C), and ToxINEc-C1 complex (D). The SEC profiles suggest the presence of the purified components in a single oligomeric state. (E) SDS-PAGE analysis of SEC-purified ToxINEc-C1 complex. Gel is stained using Coomassie stain for protein and shows the presence of ToxNEc-C1 protein. (F) Urea-PAGE analysis of SEC-purified ToxINEc-C1 complex. Gel stained using toluidine blue dye for RNA shows the presence of ToxIEc-C1 RNA repeat.
Size exclusion chromatography (SEC)
After ion-exchange chromatography, collect the fractions containing toxin, antitoxin, and complex separately.
Pool each set of fractions separately and concentrate using Amicon® Ultra-15 centrifugal filter (3.5 kDa) to a final volume of ~4 mL.
Filter the concentrated samples using a 0.45 μm syringe filter and inject into Sephacryl S-200 or Superdex S200 columns, pre-equilibrated with SEC buffer. Figure 2B–2D show the SEC elution profiles of antitoxin RNA (Figure 2B), TA RNP complex (Figure 2C), and toxin protein (Figure 2D).
Analyse the fractions on SDS-PAGE and urea-PAGE. Figure 2E and 2F show the SDS-PAGE and urea-PAGE analysis of SEC-purified ToxINEc-C1 complex for the presence of toxin protein and antitoxin RNA, respectively.
The fractions can be pooled together and concentrated appropriately depending on the experiment that needs to be performed, such as NMR spectroscopy, crystallization, or ITC. Using this protocol, we could also successfully purify two other ToxIN complexes and their components (from cluster 4 and 5 of E. coli) (Manikandan et al., 2022) (Supplementary Figure 1 and Supplementary Figure 2).
General notes and troubleshooting
The complete type III TA operon should contain i) the predicted natural promoter (including the -35 and -10 box regions), ii) the antitoxin RNA repeats, iii) the predicted transcription terminator, and iv) the toxin protein coding region (codon optimized for E. coli expression). In our study, the type III TA operon from E. coli ToxIN Cluster 1 (ToxINEc-C1) including natural promoter was synthesized and cloned into a pUC57 vector by GenScript (USA).
It is advisable to run the fractions obtained after every step of purification (Ni2+-NTA affinity chromatography, ion-exchange chromatography, and size exclusion chromatography) on the SDS-PAGE and Urea-PAGE to visualize the protein and RNA, respectively. This also ensures the visualization of impurities in the fractions eluted and whether another round of purification is required before proceeding to the biophysical experiments.
All the buffers used in the protocol were filtered with 0.22 μm sterile filters and used within one week of preparation.
For measuring the concentration of protein, any of the standard methods could be employed, such as absorbance at 280 nm or Bradford’s assay.
Absorbance at 260 nm was used for measuring the concentration of RNA.
In cases where it was difficult to measure the protein concentration accurately, 1:1 ratio of the RNA:protein was assumed to obtain an approximate concentration of the RNP TA complex.
Since the protocol involves the purification of RNA and RNA–protein complexes, it is important to be careful while handling the samples and to keep the columns away from RNases to obtain a good yield.
The toxin and antitoxin of ToxIN systems used in our study were cloned in separate compatible vectors. It is also possible to clone them in the same vector under different promoters to express and purify the components.
The toxin protein and TA complex fractions can be stored at 4 °C (up to a week) and the antitoxin RNA fractions can be stored at -20 °C (up to a month) or -80 °C (for more than a month).
Acknowledgments
The authors acknowledge the financial support received from the Department of Biotechnology (DBT), India (grant number BT/COE/34/SP15219/2015) and IISc-DBT partnership program. M.S. acknowledges the Indo-Poland grant from the Department of Science and Technology (DST), India (DST/INT/POL/P-47/2020). Authors acknowledge the DST and DBT, India, for the NMR and ITC facilities at the Indian Institute of Science, Bangalore. The authors acknowledge funding for infrastructural support from the following programs of the Government of India: DST-FIST, UGC-CAS, and the DBT-IISc partnership program. P.M. acknowledges the research fellowship from CSIR, India. K.N. acknowledges the Prime Minister’s Research Fellowship from the Ministry of Education, India. M.S. is a recipient of STAR award (award number STR/2021/000015) from the Science and Engineering Research Board (SERB), DST, India. This method is validated in the original research article (Manikandan et al., 2022).
Competing interests
There is no competing interest.
References
Blower, T. R., Pei, X. Y., Short, F. L., Fineran, P. C., Humphreys, D. P., Luisi, B. F. and Salmond, G. P. C. (2011). A processed noncoding RNA regulates an altruistic bacterial antiviral system. Nat. Struct. Mol. Biol. 18(2): 185–190.
Blower, T. R., Short, F. L., Rao, F., Mizuguchi, K., Pei, X. Y., Fineran, P. C., Luisi, B. F. and Salmond, G. P. C. (2012). Identification and classification of bacterial Type III toxin–antitoxin systems encoded in chromosomal and plasmid genomes. Nucleic Acids Res. 40(13): 6158–6173.
Fineran, P. C., Blower, T. R., Foulds, I. J., Humphreys, D. P., Lilley, K. S. and Salmond, G. P. C. (2009). The phage abortive infection system, ToxIN, functions as a protein–RNA toxin–antitoxin pair. Proc. Natl. Acad. Sci. U.S.A. 106(3): 894–899.
Cold Spring Harb Protoc. (2008). SDS loading dye. doi: 10.1101/pdb.rec11577
Cold Spring Harb Protoc. (2010). TBE electrophoresis buffer (10X). doi:10.1101/pdb.rec12231
Cold Spring Harb Protoc. (2013). Formamide Gel-Loading Buffer. doi:10.1101/pdb.rec073510
Guegler, C. K. and Laub, M. T. (2021). Shutoff of host transcription triggers a toxin-antitoxin system to cleave phage RNA and abort infection. Mol. Cell 81(11): 2361–2373.e9.
LeRoux, M. and Laub, M. T. (2022). Toxin-Antitoxin Systems as Phage Defense Elements. Annu. Rev. Microbiol. 76(1): 21–43.
Lin, J., Guo, Y., Yao, J., Tang, K. and Wang, X. (2023). Applications of toxin-antitoxin systems in synthetic biology. Eng. Microbiol. 3(2): 100069.
Manikandan, P., Sandhya, S., Nadig, K., Paul, S., Srinivasan, N., Rothweiler, U. and Singh, M. (2022). Identification, functional characterization, assembly and structure of ToxIN type III toxin–antitoxin complex from E. coli. Nucleic Acids Res. 50(3): 1687–1700.
Rao, F., Short, F. L., Voss, J. E., Blower, T. R., Orme, A. L., Whittaker, T. E., Luisi, B. F. and Salmond, G. P. C. (2015). Co-evolution of quaternary organization and novel RNA tertiary interactions revealed in the crystal structure of a bacterial protein–RNA toxin–antitoxin system. Nucleic Acids Res. 43(19): 9529–9540.
Sambrook, J. and Russell, D. W. (2006). Preparation and Transformation of Competent E. coli Using Calcium Chloride. Cold Spring Harb. Protoc. 2006(1): pdb.prot3932.
Samson, J. E., Spinelli, S., Cambillau, C. and Moineau, S. (2013). Structure and activity of AbiQ, a lactococcal endoribonuclease belonging to the type III toxin-antitoxin system. Mol. Microbiol. 87(4): 756–768.
Short, F. L., Pei, X. Y., Blower, T. R., Ong, S. L., Fineran, P. C., Luisi, B. F. and Salmond, G. P. C. (2012). Selectivity and self-assembly in the control of a bacterial toxin by an antitoxic noncoding RNA pseudoknot. Proc. Natl. Acad. Sci. U.S.A. 110(3): E241-249.
Song, S. and Wood, T. K. (2020). A Primary Physiological Role of Toxin/Antitoxin Systems Is Phage Inhibition. Front. Microbiol. 11: e01895.
Supplementary information
The following supporting information can be downloaded here:
Supplementary Figure 1. Expression and purification of type III TA components from ToxINEc-Cluster 4.
Supplementary Figure 2. Expression and purification of type III TA components from ToxINEc-Cluster 5.
Supplementary Table 1. Table showing the details of source E. coli strains of the ToxIN systems studied and plasmid vectors used to express and purify different ToxIN complexes.
Article Information
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© 2023 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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Bi-directional Dual-flow-RootChip for Physiological Analysis of Plant Primary Roots Under Asymmetric Perfusion of Stress Treatments
CA Claudia Allan
BE Blake Elliot
VN Volker Nock
CM Claudia-Nicole Meisrimler
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4764 Views: 511
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Original Research Article:
The authors used this protocol in Frontiers in Plant Science Jan 2023
Abstract
Due to technical limitations, research to date has mainly focused on the role of abiotic and biotic stress–signalling molecules in the aerial organs of plants, including the whole shoot, stem, and leaves. Novel experimental platforms including the dual-flow-RootChip (dfRC), PlantChip, and RootArray have since expanded this to plant-root cell analysis. Based on microfluidic platforms for flow stream shaping and force sensing on tip-growing organisms, the dfRC has further been expanded into a bi-directional dual‐flow‐RootChip (bi-dfRC), incorporating a second adjacent pair of inlets/outlet, enabling bi-directional asymmetric perfusion of treatments towards plant roots (shoot-to-root or root-to-shoot). This protocol outlines, in detail, the design and use of the bi-dfRC platform. Plant culture on chip is combined with guided root growth and controlled exposure of the primary root to solute changes. The impact of surface treatment on root growth and defence signals can be tracked in response to abiotic and biotic stress or the combinatory effect of both. In particular, this protocol highlights the ability of the platform to culture a variety of plants, such as Arabidopsis thaliana, Nicotiana benthamiana, and Solanum lycopersicum, on chip. It demonstrates that by simply altering the dimensions of the bi-dfRC, a broad application basis to study desired plant species with varying primary root sizes under microfluidics is achieved.
Key features
• Expansion of the method developed by Stanley et al. (2018a) to study the directionality of defence signals responding to localised treatments.
Description of a microfluidic platform allowing culture of plants with primary roots up to 40 mm length, 550 μm width, and 500 μm height.
Treatment with polyvinylpyrrolidone (PVP) to permanently retain the hydrophilicity of partially hydrophobic bi-dfRC microchannels, enabling use with surface-sensitive plant lines.
• Description of novel tubing array setup equipped with rotatable valves for switching treatment reagent and orientation, while live-imaging on the bi-dfRC.
Graphical overview
Graphical overview of bi-dfRC fabrication, plantlet culture, and setup for root physiological analysis. (a) Schematic diagram depicting photolithography and replica molding, to produce a PDMS device. (b) Schematic diagram depicting seed culture off chip, followed by sub-culture of 4-day-old plantlets on chip. (c) Schematic diagram depicting microscopy and imaging setup, equipped with a media delivery system for asymmetric treatment introduction into the bi-dfRC microchannel root physiological analysis under varying conditions.
Keywords: Microfluidics Primary root Live imaging Arabidopsis thaliana Nicotiana benthamiana Solanum lycopersicum
Background
Plant survival in abiotic and biotic stress conditions is linked with the ability to respond to the external environment through signal transduction (Lamers et al., 2020). Data supports that drought and pathogen infection are two environmental stressors with an unequivocal impact on worldwide crop, forestry, and ecosystem loss (Farooq et al., 2012; Nazarov et al., 2020). Despite the severe impact, our knowledge on how abiotic and biotic stress impact plant defence is limited. Plant growth, development, and adaptation are the outcome of a signalling process within the plant (Nejat and Mantri, 2017). Calcium is a universal signal known to modulate gene expression, hormone release, movement (e.g., muscles, stomata), and cell death (Noman et al., 2021), amongst other processes. Recent research has shown that calcium interacts with reactive oxygen species signalling, specifically hydrogen peroxide, during abiotic and biotic stress (Sewelam et al., 2016; Toyota et al., 2018). Additionally, interplay exists with downstream signals including nitric oxide and phytohormones (Freschi, 2013). However, it remains unclear how these signalling processes interact to regulate root growth and adaptation towards environmental stress. Based on a microfluidic platform for flow stream shaping on tip-growing organisms (Stanley et al., 2018a and 2018b), a refined and enhanced bi-directional dual‐flow‐RootChip (bi-dfRC) platform was established (Allan et al., 2022 and 2023). The bi-dfRC incorporates a second adjacent set of microchannel inlets/outlets connected to the base of the root observation channel for flow reversal. By combining plant culture on chip, root extension and directionality of cellular signalling responses towards local stress conditions can be tracked via controlled exposure of the root to solute changes, in a variety of combinations and spatial orientations. This protocol highlights the broad culture capabilities of the bi-dfRC towards multiple plant species including Arabidopsis thaliana, Nicotiana benthamiana, and Solanum lycopersicum. On chip root culture can also accommodate touch-sensitive transgenic plant lines through hydrophobic retention of PVP-treated microchannels. By simply altering the microchannel dimensions, the culture of plantlet primary roots up to 40 mm × 550 µm× 500 μm (length, width, height) into the bi-dfRC becomes possible, highlighting the flexibility of the platform. These measurements are the largest of interest to date; however, the technology is available to adapt and manufacture even larger devices, given compatibility with desired imaging setups. For sizeable devices that become impractical for photolithography master mold fabrication, millifluidics may be utilised via 3D printing (Kitson et al., 2012). The knowledge delivered by microfluidic technology provides insight into plant signal transduction and facilitates a platform for future research on root and plant stress perception. This may lead to novel insight towards physiological processes for improved plant tolerance towards both abiotic stress events and pathogens.
Materials and reagents
Biological materials
Arabidopsis thaliana Columbia-0 (Col-0) wild-type seeds (in-house propagated)
Nicotiana benthamiana seeds (in-house propagated)
Solanum lycopersicum Money Maker seeds (Bunnings, Country Value, catalog number: 2961272)
Reagents
Negative-tone photoresist (DJMicrolaminates, Suex, catalog number: SUEX 100)
Photoresist developer (Microchemicals, catalog number: AZ326-MIF)
Chromium etchant solution (Sigma-Aldrich, catalog number: 651826)
Propylene glycol methyl ether acetate developer (Sigma-Aldrich, catalog number: 484431)
Isopropanol (Sigma-Aldrich, catalog number: I9516)
Acetone (Sigma-Aldrich, catalog number: 90872)
Methanol (Sigma-Aldrich, catalog number: 34860)
Anti-adhesion agent Trichloro (1H,1H,2H,2H-perfluorooctyl) silane (Sigma-Aldrich, catalog number: 48931)
Polydimethylsiloxane pre-polymer silicone elastomer base and silicone elastomer curing agent (Sigma-Aldrich, Sylgard 184, Electropar, catalog number: 761028-5EA)
Deionized water (Lab made)
Sudan blue dye (Sigma-Aldrich, catalog number: 306436)
Toluene (Sigma-Aldrich, catalog number: 244511)
Norland Optical Adhesive (AusOptic, Norland Products, catalog number: NOA-72)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
Ethanol (70%) (Sigma-Aldrich, catalog number: 1.00986)
Polyvinylpyrrolidone (Kollidon) (Sigma-Aldrich, catalog number: 02286)
Murashige and Skoog medium (Duchefa, catalog number: MFCD00240976)
Plant agar (GoldBio, catalog number: P1001.0100)
2-ethanesulfonic acid (MES) (Sigma-Aldrich, catalog number: M3671)
Janola Liquid Bleach (40%) (OfficeMax, catalog number: 2233630)
Potassium hydroxide (10 M) (Sigma-Aldrich, catalog number: 06103).
Solutions
Polyvinylpyrrolidone (22% w/v) (see Recipes)
Half-strength (½) Murashige and Skoog medium with 3.1 mM MES liquid medium (see Recipes)
Plant agarose (see Recipes)
Half-strength (½) Murashige and Skoog with 0.31 mM MES plant agarose (see Recipes)
Recipes
Polyvinylpyrrolidone (22% w/v)
Reagent Final concentration Amount
Kollidon 22% 2.22 g
Deionized water n/a 10 mL
Total 22% 10 mL
Half-strength (½) Murashige and Skoog medium with 3.1 mM MES liquid medium
Reagent Final concentration Amount
Murashige and Skoog ½ 6.084 g
MES 3.1 mM 3.026 g
Water (ultrapure) n/a 500 mL
Total 10× 500 mL
*Adjust pH to 5.6 using 10 M potassium hydroxide. For a 1× working concentration, dilute 1:10 ½ Murashige and Skoog with 3.1 mM MES in water (ultrapure). Autoclave at 121 °C for 1 h.
Plant agarose (1%)
Reagent Final concentration Amount
Plant agar 1% 5 g
Water (ultrapure) n/a 500 mL
Total 1% 500 mL
*Autoclave at 121 °C for 1 h.
Half-strength (½) Murashige and Skoog with 0.31 mM MES plant agarose
Reagent Final concentration Amount
Murashige and Skoog ½ 100 mL
MES 3.1 mM 100 mL
Water (ultrapure) n/a 300 mL
Plant agar 1% 500 mL
Total 1× 500 mL
*Add liquid media to pre-cooled molten plant agar and mix well.
Laboratory supplies
Photo-masks (Nanofilm, catalog number: 4 × 4 × 0.060 SL LRC 10M 1518 5K)
Single-side polished silicon wafers 4" (Prime grade, WaferPro, catalog number: J204001)
Polyethylene sheets (OfficeMax, Overhead Projector Transparency Film A4, catalog number: 1219839)
Poly (methyl methacrylate) ring (lab-made cast acrylic, Mulford Plastics)
100 mm × 100 mm × 3 mm thick steel slab (Lab made, Nzsteel)
26 mm × 60 mm glass microscope slides (Lab Supply, catalog number: MAR0101030)
Vacuum-sealable food storage bags (Noel Leeming, Sunbeam, catalog number: VS0520)
Seed raising soil mix (Mitre 10, Tui, catalog number: TSEE1533)
Vermiculite (Mitre 10, Garden Highlights, catalog number: GHVER05)
A4 and A3 paper (OfficeMax, catalog numbers: 2220938 and 2220350, respectively)
Photolithography mask aligner filter PL-360LP (Omega Filters, catalog number: W2927)
Plastic incubation chambers (Thermo Fisher Scientific, NuncTMOmniTrayTMSingle-Well Plate, catalog number: 242811)
Scalpel with No. 10 blade (OfficeMax, Swann Morton, catalog number: 2208318)
Hole punch Ø 1 mm, 3 mm, 5 mm (ProSciTech, catalog numbers: T983-10, T983-30, and T983-50, respectively)
10 mL glass test tube (Sigma-Aldrich, catalog number: CLS9944513)
Dome propagator (Mitre 10, Otaki Hydroponics, catalog number: TG3104013)
7.5 cm × 10 cm, 750 mL, and 15 L plastic growth pots (Mitre 10, IP Plastics, catalog numbers: 169840, 283255, and 283269, respectively)
Plastic seed collection base equipped with plastic sheath (Arasystem, catalog number: ASN002, Aracons 720)
90 mm round Petri dish (Thermo Fisher Scientific, catalog number: 101VR20)
8 cm metal sieve (Stevens, Capital Kitchen, catalog number: 6546448)
Toothpicks (Discount Office, Gilmours, catalog number: GL1010819)
1.5 mL microcentrifuge tube (Thermo Fisher Scientific, Eppendorf, catalog number: 0030125150)
Sealing film (Sigma-Aldrich, Parafilm, catalog number: HS234526C-1EA)
5 mL syringe (Amtech, catalog number: 302135)
Ethylene tetrafluoroethylene tubing OD 1/16" (Kinesis, catalog number: 1517L)
Platinum-cured silicon tubing (Darwin Microfluidics, catalog number: SHE-TUB-SIL-1*1)
Stainless steel 90° bent polydimethylsiloxane couplers (Lab made, Darwin Microfluidics, catalog number: PN-BEN-18G-20)
Flangeless fittings (¼-28, Kinesis, catalog number: XP-235x)
Quick Connect Luer adapters (Scientificlabs, Kinesis, catalog number: P-628)
4× 200 mm × 20 mm × 20 mm Aluminium V slot extrusion (Aluminium Extrusion Company)
4× 300 mm × 20 mm × 20 mm Aluminium V slot extrusion (Aluminium Extrusion Company)
4× 20 series Corner Bracket 3 way (Aluminium Extrusion Company)
1× 200 mm × 200 mm Poly (methyl methacrylate) sheet (Lab made, Sigma-Aldrich, catalog number: GF53167608)
4× 1 mm Aluminium L brackets (Lab made; Aluminium Extrusion Company)
20× 20 series T Slot Spring Nuts - M4 (Aluminium Extrusion Company)
Low Pressure Unions (Kinesis, catalog number: P-702)
High-performance liquid chromatography 4-way manual switching valves (Kinesis, catalog number: V-100D)
Equipment
Laser mask writer Heidelberg μPG101 (Heidelberg Instruments)
Convection oven (Thermo Fisher Scientific, Heratherm, catalog number: 51028112)
Oxygen plasma cleaner (Tergeo, PIE Scientific)
Sky-335R6 laminator (SkyDSB)
MA-6 Mask aligner (SUSS MicroTec)
Hot plate (Torrey Pines Scientific, catalog number: HS40)
Desiccator (Sp bel-art, catalog number: F42020-0000)
Vacuum pump (VABS Ltd, catalog number: TC-2000VS)
Fumigation hood (Thermoplastic Engineering, Xtracare VSS)
S2000 ultraviolet curing system (Polydispensing, OmniCure®)
Pressurised nitrogen gun (Vacuum, BLOVAC)
Guillotine/LC slicer (Lucy Clay, catalog number: V5)
Ultra sonication bath (Lab Supply, catalog number: ELM1005507)
Vacuum sealer (Harvey Norman, Foodsaver, catalog number: VS4500)
Growth chambers (Aralab, Fitoclima, catalog number: 600)
Laminar flow hood (LaboGene, ScanLaf)
Programmable Syringe Pump PHD2000 70-2001 (Darwin-microfluidics, Harvard Apparatus, catalog number: HA-70-3007)
Epifluorescence microscope (Zeiss, catalog number: AX10)
Software
L-Edit IC Mentor Graphics Siemens EDA (v2020.1, July 2020)
Fiji, ImageJ (2.9.0, September 2022)
ZEN 2 Blue Zeiss (v3.0, October 2022)
Procedure
Photolithography and replica molding
Design the bi-dfRCs in software (L-Edit IC, v2020.1) (Figure 1a–1c; Supplemental Files 1–3).
Note: Supplemental Files 1–3 are in .dxf format and can be viewed in the software eDrawings Viewer or modified in SolidWorks/Fusion360 (the latter is free for educational use).
Transfer the pattern onto a photomask blank via laser mask writer.
Position the photomask blank with AZ1518 photoresist layer onto the stage of a laser mask writer.
Turn on the laser mask writer exposure.
Develop the photomask in AZ326-MIF photoresist developer, for 1 min, at room temperature.
Rinse the photomask with deionized water to stop development and then dry with a nitrogen gun.
Note: Ensure that the photomask is thoroughly rinsed for at least 1–2 min during this process.
Transfer the etched features of the bi-dfRC into the photomask via chrome etching.
In an acid fume hood, submerge the fully developed photomask in chromium etchant solution, with manual agitation, for 1 min.
Rinse the photomask with deionized water to stop etching and then dry with a nitrogen gun.
To remove residual photoresist, rinse the photomask with acetone followed by methanol and isopropanol for 5 min each in a sonication bath.
Note: This process yields a photomask containing fully developed etched features of the bi-dfRC microchannels. This mask can be re-used multiple times.
Figure 1. Key dimensions of the bi-dfRC platform, including inlet diameters and length of the root guidance array in the observation channel (OC) for different plants. All diagrams have been scaled to size, for visualisation purposes. Schematic diagrams depicting the key dimensions of the A. thaliana bi-dfRC (a), the N. benthamiana bi-dfRC (b), and the S. lycopersicum bi-dfRC (c). Visual representation of the wafer designs/mold pattern presented in Supplemental Files 1-3 for (d)A. thalianabi-dfRCs, (e) the N. benthamiana bi-dfRCs, and the S. lycopersicum bi-dfRC (f).
Wafer lamination
Dehydrate a 4" single-side polished silicon wafer at 180 °C for 24 h in a convection oven.
Remove the silicon wafer using protective gloves and cool until safe to handle. Critical step: Avoid leaving wafer at room temperature for extended periods to avoid adsorption of moisture.
Clean the silicon wafer in a plasma cleaner.
Set the run cycle to a power of 100 W, pulse ratio of 255, and 5 standard cubic centimetres per minute (sccm) of oxygen for 10 min.
Laminate negative-tone dry film photoresist to wafer (Data Sheets/ResearchPapers, 2020).
Set the laminator rollers to 65 °C and speed 1.
To avoid displacement while laminating, align the wafer on an aluminium sheet with raised stoppers.
Partially cover the wafer with a first polyethylene sheet.
Remove bottom protective cover off the photoresist and place the exposed side onto the polyethylene, partially overlapping the wafer.
Cover with a second polyethylene sheet fixed to the aluminium sheet.
Pass the setup under the rollers, removing the first polyethylene sheet in the process. Critical step: Stop the first polyethylene sheet from entering between the rollers to prevent it from sticking to the wafer.
Remove laminated wafer and bake on a hotplate at 65 °C for 15 min.
Mold fabrication
Power on the mask aligner.
Note: Allow the mask aligner to warm up for ~30 min.
Using an ultraviolet intensity meter with a p365 nm sensor, check the ultraviolet intensity at the wafer holder.
Set a fitted dose of 166.6 mJ/cm2 at 365 nm, based on a dry film resist thickness of 100 μm, yielding a run time of 1,170 s (Data Sheets/ResearchPapers, 2021).
Set a soft-contact run to multi-exposure for 10 s consecutive intervals with 1 min breaks, for 17 repeats.
Note: This will prevent overheating of the photoresist, reducing mechanical stress within the cross-linked resist on the silicon wafer.
Load the bi-dfRC mask.
Position a photolithography mask aligner filter over the setup.
Remove the top protective cover off the photoresist.
Place the silicon wafer with the photoresist facing up onto the wafer holder of the mask aligner.
Apply vacuum to the wafer holder, secure, close, and then start the pre-set mask aligner run.
Post-exposure bake the exposed silicon wafer on a programmable hotplate, set to a cycle of 5 min at 65 °C, 20 min at 95 °C, and then 20 min at 20 °C. Critical step: Perform bake directly after exposure. Use a low ramping speed (15 °C/h) for the cool down step to reduce mechanical stress.
Place the silicon wafer feature-side down onto a stainless-steel mesh submerged in propylene glycol methyl ether acetate developer, for 30 min or until fully developed.
Wash the silicon wafer with fresh propylene glycol methyl ether acetate and isopropanol for 5 min.
Hardbake the silicon wafer on a programmable hotplate for 1 h at 125 °C and then 20 min at 20 °C. Critical step: Use a low ramping speed (15 °C/h) for the cool down step to reduce mechanical stress.
Note: After concluding photolithography, a silicon wafer with the bi-dfRC photo-developed structures is produced. The silicon wafer is used as a casting mold for soft lithography (Figure 2a–2c). Due to the different root sizes, one A. thaliana bi-dfRC wafer yields 10 microchannels, one N. benthamiana bi-dfRC wafer yields five microchannels, and one S. lycopersicum bi-dfRC wafer yields one microchannel.
Figure 2. Silicon wafer molds, with bi-dfRC photo-developed structures. Scale = 10 mm. (a) Silicon wafer for the A. thaliana bi-dfRCs comprise of 10 individual chip structures. (b) Silicon wafer for the N. benthamiana bi-dfRCs comprise of five individual chip structures. (c) Silicon wafer for the S. lycopersicum bi-dfRCs comprise of one individual chip structure.
Silane treatment of mold wafer
Treat the silicon wafer with anti-adhesion agent.
Place the silicon wafer feature-side up into a vacuum desiccator.
Add one drop of Trichloro (1H,1H,2H,2H-perfluorooctyl) silane in an open glass shot bottle into the desiccator.
Apply vacuum pumping for 5 min.
Switch off the vacuum pump and keep the bi-dfRC mold in the desiccator under vacuum for 30 min.
Notes:
i. This will coat the silicon wafer and prevent sticking of the elastomer during subsequent soft lithography, protecting the photo-resist layer for repeated use.
ii. Silane treatment was repeated following 10 subsequent uses of the wafer as a mold for soft lithography.
Soft lithography
Mix polydimethylsiloxane pre-polymer silicone elastomer base together with curing agent in a plastic container in a 10:1 (w/w) ratio.
Place the mixture into a desiccator and apply vacuum for 30–60 min until bubbles have disappeared. Critical step: This will remove air bubbles from the mixture while assisting with solution mixing.
Cast the wafer mold.
Cover a 100 mm × 100 mm × 3 mm thick steel slab with a fresh polyethylene sheet and then place the silicon wafer atop, feature-side up.
Note: Steel slab dimensions may vary but should fit inside the desiccator.
Position a round 1.5 mm tall poly (methyl methacrylate) ring around the perimeter of the wafer.
Note: This will create a seal around the edge of the wafer to avoid polydimethylsiloxane leakage during casting.
Pour the pre-degassed polydimethylsiloxane mixture onto the silicon wafer.
Place a fresh polyethylene sheet ¾ covering the setup, followed by another 100 × 100 × 3 mm thick steel slab.
Note: This setup reduces pressure build up and adds a counterweight to hold the silicon wafer in place, while circumventing leakage of the polydimethylsiloxane between the poly (methyl methacrylate) ring and wafer.
Transfer the setup into a desiccator and apply vacuum for ~40 min until bubbles have disappeared. Critical step: This will assist with the removal of additional air bubbles formed during pouring the polydimethylsiloxane.
Transfer the setup from the desiccator to a hotplate set to 80 °C for 2 h.
Note: The polydimethylsiloxane will cure into an elastic solid.
Remove the poly (methyl methacrylate) ring from the edges of the wafer by gently probing the edges with a scalpel and then peel the polydimethylsiloxane off the wafer.
Note: Place the pre-cured polydimethylsiloxane between two fresh polyethylene sheets.
Bake the pre-cured polydimethylsiloxane on a hot plate for an additional 2 h at 80 °C.
Note: The second heating cycle improves the curing of the polydimethylsiloxane.
Allow the polydimethylsiloxane to cool; then, vertically punch the inlet/outlet channels with hole punches (1 mm for media inlets/outlets, 1–2 mm for the root inlet, and 3–4 mm for media port for all chip dimensions).
Notes:
i. Punch root inlets on an 135° obtuse angle from the glass base aligned with the feature pattern for optimal root growth into the bi-dfRC microchannel.
ii. To avoid the polydimethylsiloxane tearing, twist the hole punch gently during extraction.
Using a guillotine, cut away excess polydimethylsiloxane and separate the etched microchannels as desired.
Note: Arabidopsis bi-dfRC fit two microchannel patterns to one 26 mm × 60 mm glass microscope slide, or five microchannels to one 76 mm × 83 mm coverslip. Nicotiana benthamiana and Solanum lycopersicum bi-dfRCs fit one microchannel to one 26 mm × 60 mm glass microscope slide.
Plasma bonding
Pre-clean glass microscope slides in acetone, followed by methanol and isopropanol for 5 min each.
Place the glass slides and polydimethylsiloxane chips (feature-side up) onto the sample holder of a plasma cleaner.
Note: Place a polyethylene transparent film under the polydimethylsiloxane to prevent bonding to the sample holder.
Place the sample holder into the plasma cleaner.
Set to run at 15 W power, pulse ratio of 50, and 5 sccm of oxygen for 1 min.
Note: This will activate the bonding surfaces of the polydimethylsiloxane and glass.
Remove the sample holder from the plasma cleaner and then bond the activated side of the polydimethylsiloxane chips to the glass coverslips by lightly pressing the exposed surfaces together.
Note: If sub-optimal bonding occurs, see Troubleshooting 1.
Place the bonded bi-dfRC on a hotplate at 80 °C for 2 h.
Notes:
This final heat cycle strengthens the bond between the polydimethylsiloxane and glass.
For storage, place the bi-dfRCs in a desiccator for 3 h and then vacuum seal shut using vacuum-sealable food storage bags.
Polyvinylpyrrolidone treatment (optional)
Note: This procedure is utilised to permanently retain the hydrophilicity of the bi-dfRC microfluidic channels, to culture surface sensitive transgenic plant lines on chip, including G-CaMP3 Ca2+sensor lines (Allan et al., 2022 and 2023).
Place the completed bi-dfRCs in the plasma cleaner for 30 W power, pulse ratio 50, and 5 sccm of oxygen for 3 min.
Remove bi-dfRCs from the plasma cleaner.
Add a drop of 22% w/v polyvinylpyrrolidone onto the inlet of the bi-dfRC and then allow the solution to passively fill the microchannels.
Leave the polyvinylpyrrolidone in the microchannels for ≥ 1 min and then wash three times with deionized water.
Dry the microchannels with a nitrogen spray gun.
Place the bi-dfRCs under vacuum desiccation for 2–3 h.
Using a vacuum sealer, store the bi-dfRCs into any sturdy plastic container in a vacuum sealable food storage bag for up to one year.
Epoxy dye treatment (optional, for chip visualisation) (Soffe et al., 2020)
Dissolve 2 mg of Sudan blue dye in 1 mL of toluene in a glass test tube.
Add 250 μL of Norland Optical Adhesive with manual agitation.
Ultrasonicate the glass test tube for 10 min.
Evaporate off the toluene blue in a fume hood.
Store solution at 4 °C until use.
Pre-treat bi-dfRC microchannels in a plasma cleaner on run power 15 W, pulse ratio 50, oxygen flow rate of 3 sccm for a duration of 60 s.
Place the post-activated bi-dfRCs into a desiccator.
Add one drop of Trichloro (1H,1H,2H,2H-perfluorooctyl) silane in a glass shot bottle into the desiccator.
Apply vacuum pumping for 5 min.
Sit the bi-dfRCs in the desiccator with the lid on for 1 h without vacuum.
Note: This will allow the silane to coat the polydimethylsiloxane.
Clean the polydimethylsiloxane surface and microchannels of the bi-dfRCs with isopropyl alcohol and deionized water, for 5 min.
Dry the bi-dfRCs surfaces and microchannels with a pressurised nitrogen gun.
Place the bi-dfRCs in a desiccator for 2 h.
Note: Desiccation prior to adding the dye will remove the residual water absorbed by the polydimethylsiloxane substrate.
Using a pipette, add 60 μL of the epoxy dye onto the root inlet of the bi-dfRCs.
Note: Dye will passively enter the bi-dfRC microchannels.
Clean excess epoxy dye from the outside surface of the polydimethylsiloxane using isopropyl alcohol.
Expose the epoxy dye filled bi-dfRCs using an ultraviolet spot curing system, inverting each bi-dfRC every 2 h for the first 6 h of exposure, with the chip placed approximately 200 mm away from the light source.
Notes:
This technique allows for even exposure while ensuring rapid curing and minimisation of leeching into the polydimethylsiloxane.
This protocol obtains high-quality images of the microfluidic device, with detailed illustration of the microchannels (Figure 3a–3c).
Figure 3. Epoxy dye (blue) filled microchannels of the bi-dfRCs, for visualisation. Scale = 5 mm. (a) Dyed microchannels of the A. thaliana bi-dfRCs. (b) Dyed microchannels of the N. benthamiana bi-dfRCs. (c) Dyed microchannel of the S. lycopersicum bi-dfRC.
Plant media preparation
Prepare half-strength (½) Murashige and Skoog medium with 3.1 mM MES in water (ultrapure) and adjust the pH of the solution to 5.6 (see Recipes).
Separately, prepare 1% plant agarose in water (ultrapure) (see Recipes).
Autoclave all media at 121 °C for 1–1.5 h.
Under a laminar flow, prepare ½ Murashige and Skoog with 0.31 mM MES plant agarose plates (see Recipes).
Note: Mixing of the stocks following autoclave allows for preparation of varying media concentrations, as desired. Alternatively, ½ MS/0.31 mM MES plant agar can simply be pre-prepared in a single bottle, prior to autoclave.
Under a laminar flow, pour 50 mL of media into each 90 mm diameter Petri dish and leave to solidify.
Seal the Petri dishes with parafilm.
Store at 4 °C until use.
Seed sterilisation and vernalisation
Add approximately 50 seeds into a 1.5 mL centrifuge tube.
Add 1 mL of 0.1% Triton X-100 for 4 min with manual agitation.
Note: For larger or rougher edged seeds (N. benthamiana and S. lycopersicum), add 40% bleach to the solution for 10–15 min.
Remove the detergent solution with a pipette without disrupting the seeds.
Add 70% ethanol for 2–5 min with manual agitation.
Remove the 70% ethanol without disrupting the seeds.
Wash the seeds four times with sterile water (ultrapure).
On the final wash, keep 500 mL of sterile water (ultrapure) in the tube.
Store the seeds in water at 4 °C for at least 12 h before use.
Note: Incubation in the cold induces vernalisation of the seeds, to accelerate flowering after planting.
Seed culture on agarose plates
Under a laminar flow, plate pre-sterilised and vernalised seeds onto ½ Murashige and Skoog with 0.31 mM MES plant agarose plates using a sterile toothpick.
Seal the Petri dishes with parafilm.
Incubate the agarose plates vertically, on a short-day growth cycle [8 h light (8 am), 16 h dark (4 pm)].
Note: Sub-culture A. thaliana onto the bi-dfRC following 5 days incubation, N. benthamiana following 10–14 days incubation, and S. lycopersicum following 4–5 days incubation. At this time, roots will be the optimal length to sub-culture into the bi-dfRC root inlet (n = 100, data not shown).
Plantlet culture on chip
Remove bi-dfRCs from the vacuum storage bag.
Under a laminar flow, surface sterilise the bi-dfRCs by exposure to ultraviolet light for 30 min.
Add 100 μL of 70% ethanol into the microchannels.
Note: This is a pre-sterilisation step to avoid microorganism growth during the subsequent incubation periods. If contamination prevails during incubation, see Troubleshooting 2.
Wash out the ethanol by adding 100 μL of sterile water (ultrapure) into the microchannel three times.
Replace the sterile water (ultrapure) with liquid ½ Murashige and Skoog with 0.31 mM MES media.
Using tweezers, place small pre-cut ½ Murashige and Skoog with 0.31 mM MES agarose squares (0.5 mm, width, length, and height) next to the root inlets of the bi-dfRCs.
Remove pre-cultured plants from incubation on agarose plates and subculture onto the bi-dfRC with tweezers.
Note: Position the primary root of the plantlet directly into the root inlet channel containing liquid ½ Murashige and Skoog with 0.31 mM MES media and then gently position the leaves onto the adjacent agarose square (Figure 4a–4f).
Figure 4. Plant culture onto the bi-dfRCs. Scale = 5 mm. (a) 9-day-oldA. thalianaroots cultured in the bi-dfRC observation channels. (b) 15-day-old N. benthamiana root cultured in the bi-dfRC observation channel. (c) 7-day-old S. lycopersicum root cultured in the bi-dfRC observation channel. Image series depicting the sub-culturing process of an A. thaliana root, firstly by aligning the plantlet at the pre-media-filled root inlet of the bi-dfRC (d), then gently situating the root tip into the root inlet (e), and finally resting the leaves onto a pre-cut agarose square (f).
Place bi-dfRCs with cultured plantlets into NuncTM OmniTrayTM Single-Well Plates (plastic incubation chambers) with 10 mL of sterile water (ultrapure).
Note: This will create a humid environment for successful plantlet growth.
Seal the plastic incubation chambers with parafilm.
Incubate the plates on short-day growth cycle [8 h light (8 am), 16 h dark (4 pm)] until imaging via microscopy.
Notes:
Place the plastic incubation chambers containing cultured bi-dfRCs on a 45-degree angle to promote root growth into the observation channel.
A. thalianaroots will be ready to image from 8–10 days incubation, N. benthamiana roots will be ready to image from 15–17 days incubation, and S. lycopersicum roots will be ready to image from 7–10 days incubation (including the off-chip pre-incubation on agarose plates) (n = 100) (data not shown). The developmental stage of the primary root should show a defined tip, elongation zone, and differentiation zone of the root (root growth spanning up to 50% of microchannel).
Syringe pump system and tubing array for media injection into the bi-dfRC microchannels
Connect two plastic syringes onto a PhD 2000 Programmable Syringe Pump 70-2001 (multi syringe carrier capability) and set flow rate to 20 μL/min.
Note: Maintaining steady flow rate is important for the retention of asymmetric fluid flow in the bi-dfRC observation channels.
Connect the desired length of 1/16" OD ethylene tetrafluoroethylene tubing to the plastic syringes via Quick Connect Luer adapters with ¼-28 Flangeless fittings.
Construct the fluidic setup.
Build the flow matrix by positioning the components, including 4× high-performance liquid chromatography grade manual switching valves V-100D (solution and directional change valves), 4× Low Pressure Unions for the waste (W1A–1B and W2A–2B), and 4× Low Pressure Unions for chip connection (Inlet: I1–I2 and outlet: O1–O2) (Figure 5a–5b).
Join the components together with the desired lengths of ethylene tetrafluoroethylene tubing. Critical step: It is essential that the two circuits are comprised of equal tubing distances (for both circuits, use 33 cm of tubing between syringes and solution change valves, 24 cm between solution change valves and directional change valves, 15 cm between directional change valves and inlet fittings, 33 cm from the inlet fittings to the bi-dfRC inlets, 33 cm from bi-dfRC outlets to outlet fittings, 15 cm between outlet fittings and directional change valves, and 15 cm to the waste; tubing length optimised for the mounted solution delivery array described below) to avoid unequal fluidic resistances, ensuring that the test solutions arrive at the bi-dfRC observation channel simultaneously. For issues with uneven asymmetric flow rate, see Troubleshooting 3.
Optional: For ease, place the fluidic setup onto a raised mount.
Screw the Manual Switching Valves into a pre-cut plastic Poly(methyl methacrylate) lid with defined holes/inlets for the high-performance liquid chromatography valves.
Note: This will allow the flow matrix to be stable and visible during imaging.
Raise the setup by attaching the plastic lid to a pre-constructed metal mount box.
To construct the mental mount box, fit together four 200 mm Aluminium V slot Extrusion and four 300 mm Aluminium V slot Extrusions (cut in half, 150 mm) with four 20 series Corner 3-way Brackets.
Note: Connect all extrusions together to form a 200 mm × 200 mm × 150 mm (L, W, H) cube.
Attach a pre-cut plastic Poly (methyl methacrylate) lid to the top of the base skeleton.
Attach the Low Pressure Unions (for waste and chip inlet/outlet) to the mount using four aluminium L brackets.
Secure the setup with 20 M4 20 series T Slot Spring Nuts.
Attach the four syringes mounted on the pump system to the flow matrix via the solution change high-performance liquid chromatography grade valves equipped with ¼-28 Flangeless fittings.
Note: The setup proposed here creates parallel circuits for asymmetric flow, combining the choice of fluid on each side of the root, in addition to directional control (Figure 5c–5d).
Figure 5. Tubing array set up for delivery of test solutions into the bi-dfRC observation channel. (a) Syringe pump and tubing array setup for delivery of test solutions into the bi-dfRC microchannels, harbouring solution and directional change valves. Set to deliver full treatment (green) through inlets A & B of the bi-dfRC at the differentiation zone. (b) Placement of all solution and directional change valves, waste (W) Low Pressure Unions (WIA-1B, W2A-2B) and Low Pressure Union inlets (I) 1-2 and outlets (O) 1-2 for delivery of test solutions into the bi-dfRC. (c) Schematic diagram depicting the delivery of a full treatment A (green) through inlets 1 and 2 of the bi-dfRC. (d) Schematic diagram depicting the delivery of an asymmetric one-sided treatment A (green) through inlet 1 and treatment B (red) through inlet 2 of the bi-dfRC. Rotating the solution change valves as desired will switch which treatment enters the bi-dfRC, allowing application of full or half treatments. Adjusting the directional change valves will switch what side of the bi-dfRC the treatment enters, changing inlets 1 and 2 to outlets.
Imaging
Using an epi-fluorescent microscope (or desired microscope), place the bi-dfRCs connected to the tubing array onto the microscope stage.
Focus on the sample with brightfield under a 5× lens (EC Plan-Neofluar 5×/0.15 M27).
Pre-wet/restore filling of the flow matrix.
Connect the syringe pump system equipped with the pre-solution-filled flow matrix to the bi-dfRC inlet/outlets.
Secure the bi-dfRC with tubing attached onto the microscope stage.
Note: For a known flow rate, refer to the following equation for the time it will take media to arrive at the bi-dfRC observation channel following solution change:
t=(πr2)d/Q
where t is the time, r is the tubing radius (internal), d is the tubing length, and Q is the flow rate.
Conduct the desired experiments.
Validation of protocol
Successful A. thaliana wild-type Col-0 and transgenic G-CaMP3 plant culture and root growth into the bi-dfRC microchannel was replicated under PVP-treated microchannels (Allan et al., 2023, Figure 1d). Successful culture of wild-type N. benthamiana and S. lycopersicum was shown in this protocol (Figure 4a–4c). Bi-dfRC molds can be supplied as part of a collaboration or contract work (please [email protected]).
General notes and troubleshooting
Troubleshooting
If sub-optimal bonding of the bi-dfRCs occurs between the polydimethylsiloxane substrate and glass slide base, pre-wet the inside of the plasma cleaner chamber with deionized water to increase humidity and/or run a pre-clean cycle firstly at 100 W power, pulse ratio of 255, and 5 sccm of oxygen for 15 min (nitrogen clean), secondly at 100 W power, pulse ratio of 255, and 5 sccm of oxygen for 10 min (clean), and lastly at 15 W power, pulse ratio of 50, and 5 sccm of oxygen for 1 min (polydimethylsiloxane bond cycle).
If the bi-dfRCs are contaminated during plant incubation on chip, pre-sterilise the bi-dfRCs in an autoclave at 121 °C for 1 h, using heat safe containers.
To further control for uneven asymmetric flow in the bi-dfRC microchannel, ensure that all valves and fittings are tight, and tubing is connected securely to the bi-dfRC inlets/outlets with no tears in the polydimethylsiloxane.
Acknowledgments
The authors thank Linda Chen for help with epoxy dye treatment and Gary Turner for fabrication of the raised mount to secure the tubing array. C. Allan is recipient of a PhD scholarship from the Biomolecular Interaction Centre, Christchurch. B. Elliot holds a MSc scholarship from The Brian Mason Scientific & Technical Trust. Further funding was provided to C. Meisrimler by the Department of Science, University of Canterbury and Royal Society Te Apārangi Catalyst funding CSG-UOC1902, and to V. Nock by Rutherford Discovery Fellowship RDF-19-UOC-019 and the Biomolecular Interaction Centre, University of Canterbury. The protocol has been derived from Allan et al. (2023).
Competing interests
The authors declare that they have no conflict of interest.
References
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Supplementary information
The following supporting information can be downloaded here:
Supplemental File 1. Design for A. thaliana bi-dfRC in Mentor Graphics, exported as a dxf file
Supplemental File 2. Design for N. benthamiana bi-dfRC in Mentor Graphics, exported as a dxf file
Supplemental File 3. Design for S. lycopersicum bi-dfRC in Mentor Graphics, exported as a dxf file
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Plant Science > Plant physiology > Plant growth
Cell Biology > Cell imaging > Microfluidics
Biological Sciences > Biological techniques
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4,765 | https://bio-protocol.org/en/bpdetail?id=4765&type=0 | # Bio-Protocol Content
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This protocol has been corrected. See the correction notice.
Peer-reviewed
Establishment of Human PD-1/PD-L1 Blockade Assay Based on Surface Plasmon Resonance (SPR) Biosensor
TP Tess Puopolo *
HL Huifang Li *
JG Justin Gutkowski
AC Ang Cai
NS Navindra P. Seeram
HM Hang Ma
CL Chang Liu §
(*contributed equally to this work, § Technical contact)
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4765 Views: 1082
Reviewed by: Kazem NouriToshitsugu Fujita Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Journal of Medicinal Chemistry May 2023
Abstract
Blockade of the programmed cell death protein 1 (PD-1)/PD-ligand 1 (PD-L1) axis is a promising strategy for cancer immunotherapy. Although antibody-based PD-1/PD-L1 inhibitors have shown remarkable results in clinical cancer studies, their inherent limitations underscore the significance of developing novel PD-1/PD-L1 inhibitors. Small molecule inhibitors have several advantages over antibody-based inhibitors, including favorable tumor penetration and oral bioavailability, fewer side effects, easier administration, preferred biological half-life, and lower cost. However, small molecule inhibitors that directly target the PD-1/PD-L1 interaction are still in the early development stage, partially due to the lack of reliable biophysical assays. Herein, we present a novel PD-1/PD-L1 blockade assay using a surface plasmon resonance (SPR)-based technique. This blockade assay immobilizes human PD-1 on a sensor chip, which interacts with PD-L1 inhibitors or negative PD-L1 binders with human PD-L1 protein at a range of molecular ratios. The binding kinetics of PD-L1 to PD-1 and the blockade rates of small molecules were determined. Compared to other techniques such as PD-1/PD-L1 pair enzyme-linked immunosorbent assay (ELISA) and AlphaLISA immunoassays, our SPR-based method offers real-time and label-free detection with advantages including shorter experimental runs and smaller sample quantity requirements.
Key features
• A SPR protocol screens compounds for their capacity to block the PD-1/PD-L1 interaction.
• Validation of PD-1/PD-L1 interaction, followed by assessing blockade effects with known inhibitors BMS-1166 and BMS-202, and a negative control NO-Losartan A.
• Analysis of percentage blockade of PD-1/PD-L1 of the samples to obtain the IC50.
• Broad applications in the discovery of small molecule–based PD-1/PD-L1 inhibitors for cancer immunotherapy.
Graphical overview
Keywords: PD-1 PD-L1 Blockade Surface plasmon resonance Cancer Immunotherapy
Background
Cancer is a global health burden resulting in high healthcare costs. Therefore, the search for effective therapeutics is of continued scientific interest. Recently, cancer immunotherapy, a strategy that utilizes the host’s own immune system to fight tumors, has become an effective treatment of cancers (Makuku et al., 2021). In cancer, the tumor cell microenvironment acts to inhibit immune checkpoints, which normally function to prevent uncontrolled proliferation (He and Xu, 2020). Programmed cell death protein 1 (PD-1), an immune checkpoint expressed by several types of immune cells, dampens the immune system upon programmed cell death ligand-1 (PD-L1) binding (Makuku et al., 2021). The interaction of PD-1/PD-L1 leads to the inhibition of phosphorylation of the T-cell-receptor (TCR) signaling intermediate, which terminates the TCR signaling cascade (Keir et al., 2008; Fife et al., 2009). Two signaling motifs in the cytoplasmic tail of PD-1 are the intracellular immunoreceptor tyrosine-based switch motif (ITSM) and the immunoreceptor tyrosine-based inhibitory motif. Upon PD-L1 binding to PD-1, ITSM is phosphorylated and recruits Src homology 2-containing tyrosine phosphatase, thereby inhibiting the phosphatidylinositol 3-kinase (PI3K)/Akt signaling pathway (Barclay et al., 2018; Ai et al., 2020). PI3K/Akt signaling pathway blockage further downregulates the mechanistic targets of rapamycin and inhibits protein synthesis and cell growth. PI3K/Akt signaling pathway blockage also inhibits the degradation of transcription factor FoxO1, which enhances the expression of PD-1 (Barclay et al., 2018; Ai et al., 2020).
The recognition of the PD-1 protein on the membrane of T cells by tumor cells results in the upregulation of PD-L1 (J. Liu et al., 2021). High expression of PD-L1 is one characteristic observed in many types of tumors including melanoma, lung cancer, and breast cancer (Mu et al., 2011; Fusi et al., 2015; Aguilar et al., 2019). PD-1/PD-L1 binding results in T-cell apoptosis (J. Liu et al., 2021). Blockade of the PD-1/PD-L1 axis results in tumor suppression due to interference between the tumor cell and the T cell (C. Liu et al., 2021; Makuku et al., 2021). Numerous studies have demonstrated that the blockage of PD-L1 or PD-1 is one of the most promising approaches for cancer immunotherapy (Zitvogel and Kroemer, 2012; Wu et al., 2018; Salmaninejad et al., 2019). Blocking the interactions of PD-L1 and PD-1 shuts off the inhibitory signaling pathways for T cells, reactivates the T cell–mediated anti-tumor responses by promoting T-cell proliferation, and enhances effector T-cell function (Salmaninejad et al., 2019; Tang and Zheng., 2018). Clinical data have demonstrated that the blockade of PD-1 or PD-L1 can boost T cell–mediated antitumor responses, generates durable clinical responses, and prolongs patient survival time (Ohaegbulam et al., 2015; Alsaab et al., 2017). Monoclonal antibodies against PD-1 (Pembrolizumab, Nivolumab, and Cemiplimab) or PD-L1 (Atezolizumab, Avelumab, and Durvalumab) have been approved by FDA for the treatment of a series of malignancies including breast cancer, bladder cancer, colorectal cancer, lung cancer, hepatoma, and melanoma (Massard et al., 2016; Kim, 2017; Xin Yu et al., 2020). Although these monoclonal antibodies demonstrated promising results with high clinical efficacy and immune-related adverse effects, immunogenicity and high costs are still major limitations of antibody-based immune checkpoint inhibitors (Hamanishi et al., 2015; Alsaab et al., 2017; Zinzani et al., 2017; Akinleye and Rasool et al., 2019). Alternatively, small-molecule inhibitors can overcome these advantages due to better tumor penetration and oral availability (Zhan et al., 2016).
Therefore, the discovery of small molecule inhibitors blocking the PD-1/PD-L1 interaction is a promising cancer therapy approach. Our group reported the evaluation of the PD-1/PD-L1 blockade (using a pair-ELISA technique) and the binding of compounds to either PD-1 or PD-L1 (Li et al., 2022). However, this method is not efficient for large-scale screenings of small molecule libraries for PD-1/PD-L1 inhibitors. Therefore, we developed a surface plasmon resonance (SPR)-based PD-1/PD-L1 blockade screening approach utilizing immobilized PD-1 (on the chip), PD-L1 (in solution), and known inhibitors (i.e., BMS-1166 or BMS-202, in solution). To exclude potential false positives, we included a negative PD-L1 inhibitor (NO-Losartan A) possessing a biphenyl group—a structural feature shared with the BMS-1166 and the BMS-202 compounds that were investigated together in this study. Notably, the SPR technique is a valuable complementary method to ELISA immunoassays that can also be used for the optimization of ELISA-based assays (Vaisocherová et al., 2009). SPR is an optical biosensor technology that employs the evanescent wave phenomenon to detect changes in the refractive index of a biosensor (Pattnaik, 2005). A light source illuminates the biosensor and prism, and as the analyte flows through the channel and binds to the target protein, the refractive index of the biosensor undergoes a shift. This interaction between analyte and protein is monitored in real-time, enabling precise measurement of the amount of bound protein as well as the rates of association and dissociation. The SPR assay has unique advantages over the ELISA-type assay. Rather than merely providing an endpoint, the SPR assay monitors the kinetics associated with the PD-1/PD-L1 blockade of small molecules in real time. We acknowledge that ELISA-type assays are more widely accessible and adaptable to different laboratory settings, and we recognize that our SPR assay requires specialized instrumentation and expertise, which may not be available in all laboratories. Furthermore, given that this blockade assay is solely based on in vitro experiments, it is imperative to perform functional assays and in vivo validation to confirm the potential of compounds exhibiting blockade effects against PD-1/PD-L1. However, we believe that this SPR-based protocol, which provides sufficient details, can facilitate the screening process of small molecule inhibitors that block the PD-1/PD-L1 interaction at a large scale.
In the present study, we utilized an SPR-based assay to determine the IC50 values of BMS-1166 and BMS-202, which were measured at 85.4 and 654.4 nM, respectively. BMS-1166 has been previously characterized with an IC50 value of 1.4 and 276 nM by homogeneous time-resolved fluorescence (HTRF) and cell-based assays (Jurkat cells expressing PD-1 in co-culture with CHO cells expressing PD-L1), respectively (Guzik et al., 2017). Previous investigations have reported IC50 values of BMS-202 at 18 and 96 nM utilizing different assays, including cell-based and HTRF approaches (Surmiak et al., 2021). Our results are comparable with previous findings and confirm the reliability and reproducibility of our SPR-based protocol.
Materials and reagents
Biological materials
Human PD-L1/B7-H1 protein, Fc Tag (ACROBiosystems, catalog number: D1-H5258)
Human PD-1/PDCD1 protein, Fc Tag, low endotoxin (ACROBiosystems, catalog number: PD1-H5257)
Reagents
BMS-1166 (Med Chem Express, catalog number: HY-102011)
BMS-202 (Med Chem Express, catalog number: HY-19745)
Amine Coupling kit [ethanolamine hydrochloride, dimethylaminopropyl-N’ethylcarbodiimide N-3-hydrochloride (EDC), and N-hydroxy succinimide (NDC)] (Global Life Sciences Solutions, Cytiva, catalog number: BR100050)
NO-Losartan A (Cayman Chemical Company, catalog number: Cay10006456)
Solutions
HBS-EP+ buffer 10× (Global Life Sciences Solutions, Cytiva, catalog number: BR100826)
Glycine 1.5 (Global Life Sciences Solutions, Cytiva, catalog number: BR100354)
Dimethyl sulfoxide (DMSO), anhydrous ≥ 99.9% (Sigma-Aldrich, catalog number: 276855)
DNase-free water (Fisher Scientific, catalog number: 188506)
Acetate 5.0 (Global Life Sciences Solutions, Cytiva, catalog numbers: BR100350, BR100351)
NaOH 50 mM (Global Life Sciences Solutions, Cytiva, catalog number: 100358)
PD-1 or PD-L1 protein solution, 500 μg/mL (PD-L1 protein solution equals to 2,000 nM) (see Recipes)
HBS-EP+ running buffer (250 mL Fisherbrand glass bottle) (see Recipes)
HBS-EP+ running buffer + 0.01% DMSO (250 mL Fisherbrand glass bottle) (see Recipes)
Recipes
PD-1 or PD-L1 protein solution, 500 μg/mL (PD-L1 protein solution equals to 2,000 nM)
Add 200 μL of DNase-free water to 100 μg of PD-1 or PD-L1 protein. (To prevent nucleic acid contamination, it is recommended to use DNase-free water for the preparation of a long-term stock solution of PD-1 or PD-L1 proteins. However, for assays utilizing fresh protein solutions, Milli-Q water is sufficient.)
HBS-EP+ running buffer (250 mL Fisherbrand glass bottle)
Add 20 mL of the HBS-EP+ buffer 10× to 180 mL of Milli-Q water.
HBS-EP+ running buffer + 0.01% DMSO (250 mL Fisherbrand glass bottle)
Add 25 mL of the HBS-EP+ buffer 10× to 224.975 mL of Milli-Q water.
Add 25 μL DMSO.
Laboratory supplies
Pipettes (ErgoOne Single Channel Pipette 2.5, 10, 200, 1,000 μL; USA Scientific, catalog numbers: 7100-0125, 7100-0510, 7100-2200, 7110-1000)
Pipette Tips (TipOne 10, 200, 1,000 μL; USA Scientific, catalog numbers: 1111-3800, 1110-1800, 1111-2821)
SCI-Fill motorized pipette filler (Scilogex, catalog number: 740200029999)
Serological pipettes 10 and 50 mL (Thermo ScientificTMNuncTM, catalog numbers: 02-923-204, 02-923-206)
96-well polystyrene microplates (Global Life Sciences Solutions, Cytiva, catalog number: BR100503)
Microplate foil, 96-well (Global Life Sciences Solutions, Cytiva, catalog number: 28975816)
Plastic vials 7 mm (Global Life Sciences Solutions, Cytiva, catalog number: BR100212)
Rubber caps, type 3 (Global Life Sciences Solutions, Cytiva, catalog number: BR100502)
Series S Sensor Chip CM5 (Global Life Sciences Solutions, Cytiva, catalog number: BR100530)
Fisherbrand reusable glass media bottles with cap 250 mL (Fisher Scientific, catalog number: FB800100)
Microcentrifuge tube 1.5 mL non-sterile (Cell Treat, Wilkem Scientific, catalog number: LCEL229441)
PCR tubes individual 0.2 mL flat cap (PureAmp, Wilkem Scientific, catalog number: LMTP3030)
Equipment
Biacore T200 SPR (Global Life Sciences Solutions, Cytiva, catalog number: 28975001)
Software and datasets
Biacore T200 analysis software (BIAevaluation version 4.1)
GraphPad Prism 9.1.2 (https://www.graphpad.com/updates/prism-912-release-notes)
Procedure
Immobilization of PD-1: Amine Coupling Method
Set immobilization method on the SPR instrument.
Note: All steps should follow the instrument’s manual.
Set temperature to 25 °C.
Set chip type as CM5.
Set flow cells per cycle as 1.
Under flow cell 1, check immobilize flow cell 1.
i. Set amine as the method.
ii. Set blank immobilization.
Under flow cell 2, check immobilize flow cell 2.
i. Set amine as the method.
ii. Set ligand as 40 μg/mL PD-1.
iii. Set aim for immobilization method with desired target level and the wash solution as 50 mM NaOH.
Set tubes R2 B1 as 40 μg/mL PD-1.
Set tubes R2 B2 as 50 mM NaOH.
Set tubes R2 B3 and R2 C3 as Ethanolamine.
Set tubes R2 B4 and R2 C4 as Empty.
Set tubes R2 B5 and R2 C5 as NHS.
Set tubes R2 B6 and R2 C6 as EDC.
Prepare 200 mL of HBS-EP+ running buffer solution (see Recipes).
Prepare Amine Coupling kit reagents and PD-1 protein in a reagent rack.
Prepare a stock solution of PD-1 at 500 μg/mL in DNase-free water (see Recipes).
i. Set at room temperature for 30 min to fully dissolve.
ii. Dilute PD-1 solution to 40 μg/mL in acetate 5.0 and add 160 μL to tube R2 B1.
Add 70 μL of 50 mM NaOH to tube R2 B2.
Add 140 μL of ethanolamine to tubes R2 B3 and R2 C3.
R2 B4 and R2 C4 remain empty.
Add 100 μL of NHS to tube R2 B5 and R2 C5 (NHS is included in the Amine Coupling kit).
Add 100 μL of EDC to tubes R2 B6 and R2 C6 (EDC is included in the Amine Coupling kit).
Place Tube A into the HBS-EP+ running buffer solution.
Eject the maintenance sensor chip and insert the CM5 chip.
Reopen the immobilization method, eject the rack, and insert reagent rack 2.
Run method for the estimated run time.
The following method will be performed:
Inject ligand solution for five pre-concentrations.
Establish a baseline with an injection of the HBS-EP+ running buffer solution.
Mix inject a 50:50 ratio of EDC + NHS with a contact time of 420 s and a flow rate of 10 μL/min to activate the chip surface with the modification of carboxymethyl groups to N-hydroxysuccinimide esters.
Continue baseline with an injection of the HBS-EP+ running buffer solution after chip modification. The baseline activation will observe a slight response unit (RU) effect.
Inject 40 μg/mL of PD-1 ligand to induce an electrostatic interaction that will couple the ligand to the chip surface. The ligand includes both immobilized and non-covalently bound proteins. At this stage, the PD-1 solution remains in contact with the CM5 sensor surface, resulting in a response that includes both immobilized and non-covalently bound PD-1. The N-hydroxysuccinimide esters present on the sensor chip surface react spontaneously with the primary amines on PD-1 to form stable and covalent links.
Immobilize the ligand prior to deactivation. This indicates that the ligand has surpassed the protein surface and the majority of the non-covalently bound ligand has been removed.
Deactivate remaining NHS-esters and remove unreacted esters through the injection of ethanolamine with NaOH utilizing a contact time of 420 s and a flow rate of 10 μL/min. The unreacted NHS-esters were deactivated using 35 μL of 1 M ethanolamine hydrochloride, which was adjusted to pH 8.5 with NaOH. Additionally, the deactivation process ensures the removal of any remaining electrostatically bound PD-1.
Notes:
i. A slight increase in RU is observed due to the change in the bulk refractive index.
ii. Stopping point if desired. Eject chip, gently wash chip surface with two drops of DI water, briefly let dry, and place at 4 °C. Replace running buffer with Milli-Q water for standby. The reproducibility of this immobilization protocol has been confirmed through multiple experiments conducted in our research group. It is important to note that when dissolving PD-L1 recombinant proteins (in lyophilized solid form), a minimum of 30 min of equilibrium time at room temperature is necessary to ensure full dissolution of the protein.
Validation of PD-1/PD-L1 interaction
Set PD-1/PD-L1 validation method on the SPR instrument.
Set temperature to 25 °C.
Under General Settings, set the data collection rate at 10 Hz, detection as multi, sample compartment temperature at 25 °C, and concentration unit as nM.
Under Assay Steps, set conditioning replicates to 20 times, startup to kinetics with replicates at 10 times, sample to kinetics with 1 replicate, and temperature at 25 °C.
Under Cycle Types, select new and enter Kinetics.
i. In commands, insert Capture, sample 1, and select the sample settings as high performance with a contact time of 120 s, dissociation time of 120 s, a flow rate of 10 μL/min, and a flow path of 1, 2, 3, 4. Under method variables, set property as a variable and select sample solution. Under evaluation variables, select evaluation purpose as kinetics/affinity and select the predefined variables as Conc and MW.
ii. In commands, insert Regeneration 1 and enter the Regeneration solution as Glycine 1.5 with a contact time of 30 s, a flow rate of 30 μL/min, and a flow path of 1, 2, 3, 4.
iii. In commands, insert Carry-over control 1 (injection of 30 s with a flow rate of 40 μL/min).
Under Cycle Types, select new and enter Conditioning. In commands, select capture and set it as Carry-over control 1 (injection of 30 s with a flow rate of 40 μL/min).
Under Variable Settings, select startup and select Define all values in the method. Enter values for the variables as sample buffer, under the sample 1 Sample Solution column header.
Under Variable Settings, select a sample and select Define all values at run time. The table should read the Command as Sample, and the Variable as Sample solution, Conc, and MW.
Select Verification to ensure the method has been verified and can be used to set up a run.
Review Overview assay steps including conditioning, startup, and sample and ensure the settings are as desired.
Select Setup Run.
Under Detection, select the flow path as 2-1, 4-3.
Input the sample solution: PD-L1 with concentrations from 0, 5, 10, 20 to 40 nM and a molecular weight of 51,300 Da.
Next, review the overview of assay steps for verification.
Select prime before the run.
Set each PD-L1 concentration as a separate sample well position in a 96-Well Microplate layout:
i. R1 A1 as PD-L1 0 nM.
ii. R1 A2 as PD-L1 5 nM.
iii. R1 A3 as PD-L1 10 nM.
iv. R1 A4 as PD-L1 20 nM.
v. R1 A5 as PD-L1 40 nM.
Set Glycine 1.5 for regeneration in the same plate layout.
i. R1 B1–B12.
Set sample buffer for a startup in desired well positions.
i. R1 C1–C4.
Prepare 200 mL of HBS-EP+ running buffer solution (see Recipes).
Prepare PD-L1 concentrations.
Prepare PD-L1 protein at 500 μg/mL (2,000 nM) (see Recipes).
Dilute to 40 nM (200 μL) in HBS-EP+ running buffer and add to the 96-well plate (R1 A5).
Add 100 μL of HBS-EP+ running buffer (R1 A1, R1 A2, R1 A3, and R1 A4) and perform a 2-fold serial dilution (100 μL) in the plate to 20, 10, and 5 nM.
Add 250 μL of the regeneration solution (Glycine 1.5) to R1 B1–B12.
Add 100 μL of HBS-EP+ running buffer to R1 C1 and 225 μL to R1 C2–C4.
Tightly cover the microplate with a microplate seal.
Remove the previously immobilized PD-1 CM5 chip from 4 °C.
Eject the maintenance chip and insert the PD-1 CM5 chip.
Place Tube A into the HBS-EP+ running buffer + 0.01% DMSO solution.
Open the established method, eject the rack, and insert a 96-well microplate.
Run method for the estimated run time.
After running the method, eject the chip and insert the maintenance chip.
Replace the running buffer with Milli-Q water for standby.
Note: The reproducibility of the PD-1/PD-L1 interaction step has been confirmed through multiple experiments conducted in our research group. This step is crucial in determining the binding affinity of PD-L1 at different concentrations, which, in turn, is necessary for identifying the optimal concentration of PD-L1 for subsequent blockade assays.
PD-1/PD-L1 blockade assay with established small molecule inhibitor: BMS-1166
Set the method on the SPR instrument.
Set temperature to 25 °C.
Set General Settings as the same as the validation protocol.
Under Assay Steps, set conditioning replicates to 20 times, set startup to conditioning with replicates at 10 times, sample to kinetics with 1 replicate, and temperature at 25 °C.
Set Cycle Types (both kinetics and conditioning) as the same as the validation protocol.
Under Variable Settings, select startup and select Define all values in the method. Enter values for the variables as sample buffer, under the sample 1 Sample solution column header.
Select Verification to ensure the method has been verified and can be used to set up a run.
Review Overview assay steps including conditioning, startup, and sample, and ensure the settings are as desired.
Select Setup Run.
Under Detection, select the flow path as 2-1, 4-3.
Input the sample solution: PD-L1 20 nM + BMS-1166 with concentrations from 0, 1, 5, 25, 125, 625, to 3,125 nM and a molecular weight of 51,300 Da.
Next, review the overview of assay steps for verification.
Select prime before run.
Set each PD-L1/BMS-1166 concentration as a separate sample well position in a 96-well microplate layout:
i. R1 A1 as PD-L1 20 nM + BMS-1166 0 nM.
ii. R1 A2 as PD-L1 20 nM + BMS-1166 1 nM.
iii. R1 A3 as PD-L1 20 nM + BMS-1166 5 nM.
iv. R1 A4 as PD-L1 20 nM + BMS-1166 25 nM.
v. R1 A5 as PD-L1 20 nM + BMS-1166 125 nM.
vi. R1 A6 as PD-L1 20 nM + BMS-1166 625 nM.
vii. R1 A7 as PD-L1 20 nM + BMS-1166 3,125 nM.
Set Glycine 1.5 for regeneration in reagent rack 2.
i. R2 A1.
Prepare 250 mL of HBS-EP+ running buffer solution (see Recipes).
Prepare PD-L1 solution at 20 nM in HBS-EP+ running buffer (1.5 mL microcentrifuge tube).
Add 1.5 μL of PD-L1 protein solution (2,000 nM stock to 148.5 μL of HBS-EP+ running buffer).
Prepare PD-L1 solution at 20 nM in HBS-EP+ running buffer + 0.01% DMSO (1.5 mL microcentrifuge tube).
Add 5 μL of PD-L1 protein solution 2,000 nM stock to 494.5 μL of HBS-EP+ running buffer.
Add 0.5 μL of 10% DMSO.
Prepare the BMS-1166 concentrations and the regeneration solution in a 96-well microplate.
Prepare a 31.25 mM stock of BMS-1166 in DMSO (10 μL) (in PCR tube).
Dilute the BMS-1166 stock to 3,125 nM in 100 μL of PD-L1 (20 nM) HBS-EP+ running buffer and add to the 96-well plate: R1 A7.
Add 80 μL of PD-L1 (20 nM) + HBS-EP+ running buffer + 0.01% DMSO solution to R1 A6–R1 A2 and perform a 5-fold serial dilution (20 μL) in the plate to 625, 125, 25, 5, and 1 nM.
Add 80 μL of PD-L1 (20 nM) + HBS-EP+ running buffer + 0.01% DMSO solution to R1 A1.
Add 3 mL of the regeneration solution (Glycine 1.5) to R2 A1.
Add 0.01% DMSO to HBS-EP+ to use as the running buffer.
Tightly cover the plate with a microplate seal.
Remove the previously immobilized PD-1 CM5 chip from 4 °C.
Eject the maintenance chip and insert PD-1 CM5 chip.
Place Tube A into the HBS-EP+ running buffer + 0.01% DMSO solution.
Open the established method, eject the rack, and insert the 96-well microplate.
Run method for the estimated run time.
After running the method, eject the chip and insert the maintenance chip.
Wash the chip with two drops of DI water and place at 4 °C.
Replace running buffer with Milli-Q water for standby.
PD-1/PD-L1 blockade assay with established small molecule inhibitor: BMS-202
Set the same method on the SPR instrument as the BMS-1166 method.
Set each PD-L1/BMS-202 concentration as a separate sample well position in a 96-well microplate layout:
i. R1 A1 as PD-L1 20 nM + BMS-202 0 nM.
ii.R1 A2 as PD-L1 20 nM + BMS-202 10 nM.
iii. R1 A3 as PD-L1 20 nM + BMS-202 100 nM.
iv. R1 A4 as PD-L1 20 nM + BMS-202 500 nM.
v. R1 A5 as PD-L1 20 nM + BMS-202 1,000 nM.
vi. R1 A6 as PD-L1 20 nM + BMS-202 5,000 nM.
Set Glycine 1.5 for regeneration in reagent rack 2.
i. R2 A1.
Prepare 250 mL of HBS-EP+ running buffer.
Prepare PD-L1 solution (20 nM) in HBS-EP+ running buffer (1.5 mL microcentrifuge tube).
Add 1.5 μL of PD-L1 protein solution (2,000 nM stock to 148.5 μL of HBS-EP+ running buffer).
Prepare PD-L1 solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO solution (1.5 mL microcentrifuge tube).
Add 5 μL of PD-L1 2,000 nM stock to 494.5 μL of HBS-EP+ running buffer.
Add 0.5 μL of 10% DMSO.
Prepare the BMS-202 concentrations and the regeneration solution in a 96-well microplate.
Prepare 50 mM stock solution of BMS-202 dissolved in DMSO.
In R1 A6, dilute the BMS-202 stock to 5,000 nM in 120 μL of PD-L1 solution (20 nM) in HBS-EP running buffer.
In R1 A5, perform a 5-fold dilution by adding 20 μL from R1 A6 with 80 μL of PD-L1 solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO.
In R1 A4, perform a 10-fold dilution by adding 10 μL from R1 A6 with 90 μL of PD-L1 solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO.
In R1 A3, perform a 10-fold dilution by adding 10 μL from R1 A5 with 90 μL of PD-L1 solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO.
In R1 A2, perform a 10-fold dilution by adding 10 μL from R1 A3 with 90 μL of PD-L1 solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO.
In R1 A1, add 90 μL of PD-L1 protein solution (20 nM) in HBS-EP+ running buffer + 0.01% DMSO.
Add 3 mL of the regeneration solution (Glycine 1.5) to R2 A1.
Add 0.01% DMSO to the HBS-EP+ running buffer.
Proceed with the same procedure as BMS-1166 to run BMS-202.
PD-1/PD-L1 blockade assay with negative control: NO-Losartan A
Set the same method on the SPR machine as the BMS-1166/BMS-202 methods.
Set each PD-L1/NO-Losartan A concentration as a separate sample well position in a 96-well microplate layout:
i. R1 A1 as PD-L1 20 nM + NO-Losartan A 0 nM.
ii. R1 A2 as PD-L1 20 nM + NO-Losartan A 1 nM.
iii. R1 A3 as PD-L1 20 nM + NO-Losartan A 5 nM.
iv. R1 A4 as PD-L1 20 nM + NO-Losartan A 25 nM.
v. R1 A5 as PD-L1 20 nM + NO-Losartan A 125 nM.
vi. R1 A6 as PD-L1 20 nM + NO-Losartan A 625 nM.
vii. R1 A7 as PD-L1 20 nM + NO-Losartan A 3,125 nM.
Prepare PD-L1 (20 nM), NO-Losartan A at different concentrations, HBS-EP+ running buffer, HBS-EP+ running buffer + 0.01% DMSO buffer, and regeneration solution, as described in the BMS-1166 protocol.
Proceed with the same procedure as BMS-1166/BMS-202 to run NO-Losartan A.
Note: In this protocol, NO-Losartan A was employed as a negative control due to the presence of a biphenyl group, which is a structural feature shared by the BMS-1166 and BMS-202 compounds tested. To ascertain the suitability of NO-Losartan A as a negative control, we conducted a preliminary investigation of its binding affinity with PD-L1. Our findings indicated that NO-Losartan A displayed negligible binding affinity towards PD-L1, confirming its suitability as a negative control in this protocol. For screening purposes, the selection of a negative control may depend on the specific composition of the compound library employed. It is essential to ensure that the chosen negative control exhibits no binding affinity towards the PD-L1.
Data analysis
Immobilization of PD-1:
Data were analyzed via the output from the SPR instrument indicating a low RU of the blank cell and a successful target RU of the PD-1 ligand.
Validation of PD-1/PD-L1 interaction:
Data were analyzed using the corresponding Biacore T200 analysis software (BIAevaluation version 4.1)
Under kinetics/affinity, select surface bound.
Select the curve as 2-1.
Perform a 1:1 binding mode with a constant fit to obtain the association rate (Ka), dissociation rate (Kd), and dissociation constant (KD).
Export analyzed curves into GraphPad Prism for graphical representation of data.
PD-1/PD-L1 blockade assay with established small molecule inhibitors (BMS-1166 and BMS-202) and negative control (NO-Losartan A):
Data were analyzed using the corresponding Biacore T200 analysis software.
Under kinetics/affinity, select surface bound.
Select the curve as 2-1.
Export curves into GraphPad Prism for graphical representation of data.
Blockade rate and IC50 value of each compound were analyzed using GraphPad Prism.
To determine the percentage blockade of each sample concentration, employ the subsequent formula: Percentage blockade (%) = [1 – (RU of PD-L1 incubated with the compound/RU of PD-L1 in the absence of the compound)] × 100.
The concentration of each compound and the blockade rate of each compound were imported into GraphPad Prism.
The XY analysis function was selected, and log (inhibitor) vs. response – Variable slope (four parameters) was chosen.
The IC50 values of each compound were obtained, and a goodness-of-fit assessment was performed. A coefficient of determination (R-squared) greater than 0.99 was required for a satisfactory fit.
Validation of protocol
Immobilization of PD-1 on SPR chip:
Flow cell 1 was immobilized as the blank with a final response (RU) of 103.1 (Figure 1A).
Flow cell 2 was immobilized with PD-1 ligand coated on the chip surface with a final response (RU) of 3688.5, indicating a successful reach of the target (Figure 1B).
Figure 1. Immobilization curves of (A) flow cell 1 (blank) and (B) flow cell 2 (PD-1). The first phase (phase 1) represents a stable baseline, while the second phase (phase 2) displays a responsive effect in the response unit (RU). In phase 3, a wash with ethanolamine hydrochloride (1 M; pH 8.5) was conducted. In phase 4, recombinant PD-1 protein (40 μg/mL) was injected and coupled to the surface matrix of flow cell 2, and running buffer was injected in the flow cell 1. In phase 5, any remaining electrostatically bound ligand was removed using ethanolamine hydrochloride (1 M; pH 8.5) to deactivate unreacted NHS-esters with a contact time of 420 s and a flow rate of 10 μL/min.
Validation of PD-1/PD-L1 interaction:
The binding interaction of 5, 10, 20, and 40 nM PD-L1 in solution to PD-1 on the chip surface was observed with a quantifiable response (Figure 2).
The analyzed binding parameters included an association rate (Ka) = 8.852 × 104 1/Ms, dissociation rate (Kd) = 0.01146 1/s, and dissociation constant (KD) = 1.295 × 10-7M.
Figure 2. Binding kinetics of PD-L1 to PD-1 at varying concentrations. The real-time surface plasmon resonance (SPR) response of the sensor chip to the binding reactions between PD-L1 (in solution) and PD-1 (immobilized on the chip) is displayed. The concentration series of PD-L1 used (5–40 nM; 2-fold dilutions) is shown. PD-L1 demonstrates distinct association (0–120 s) and dissociation phases (121–240 s) with PD-1, which are clearly visible.
PD-L1/PD-1 blockade assay with established small molecule inhibitor: BMS-1166
Blockade of the PD-1/PD-L1 binding interaction was observed by 1–3,125 nM BMS-1166 with 20 nM PD-L1 protein in 0.01% DMSO solution (Figure 3A). There was a concentration-dependent increase in percentage blockade of PD-1/PD-L1 with an IC50 of 85.4 nM (Figure 3B). The percentage blockade for 0, 1, 5, 25, 125, 625, and 3,215 nM BMS-1166 was 0%, 12.7%, 18.5%, 31.5%, 62.4%, 85.9%, and 94.2%, respectively (Figure 3C).
Figure 3. Blockade of PD-1/PD-L1 interaction with BMS-1166. (A) A representative real-time surface plasmon resonance (SPR) response to the binding reactions between PD-L1 (in solution) and PD-1 (immobilized on the chip) in the presence of BMS-1166, at various concentrations ranging from 1 to 3,215 nM, with a fixed concentration of 20 nM of PD-L1 in solution. (B) The sigmoidal binding profile of BMS-1166 on the log scale with an indicated IC50 of 85.4 nM. (C) BMS-1166 concentration-dependent increase in percentage blockade.
PD-1/PD-L1 blockade assay with established small molecule inhibitor: BMS-202:
Blockade of the PD-1/PD-L1 binding interaction was observed by 1–5,000 nM BMS1166 with 20 nM PD-L1 protein in a solution with 0.01% DMSO (Figure 4A). There was a concentration-dependent increase in percentage blockade of PD-1/PD-L1 with an IC50 of 654.4 nM (Figure 4B). The percentage blockade for 0, 10, 100, 500, 1,000, and 50,000 nM BMS-202 was 0%, 14.9%, 18.5%, 32.0%, 56.4%, and 67.1%, respectively (Figure 4C).
Figure 4. Blockade of PD-1/PD-L1 interaction with BMS-202. (A) BMS-202 at 0, 10, 100, 500, 1,000, and 5,000 nM with 20 nM PD-L1 in solution lowered the PD-1/PD-L1 binding response unit (RU), concentration dependently. (B) The sigmoidal binding profile of BMS-202 on the log scale with an indicated IC50 of 654.4 nM. (C) BMS-202 concentration-dependent increase in percentage blockade.
PD-1/PD-L1 blockade assay with negative control: NO-Losartan A:
No blockade of the PD-1/PD-L1 interaction was observed at 0, 1, 5, 25, 125, 625, and 3,125 nM NO-Losartan A (negative control) with 20 nM PD-L1 in 0.01% DMSO solution (Figure 5A). There was no observable binding profile for NO-Losartan A with a non-detectable IC50.
The percentage blockade for 0, 1, 5, 25, 125, 625, and 3,215 nM NO-Losartan A was 0%, 1.08%, 1.51%, 1.21%, 2.05%, 0%, and 0%, respectively (Figure 5B and 5C).
Figure 5. Failure to block the PD-1/PD-L1 interaction with NO-Losartan A. (A) PD-L1 was incubated with increasing concentrations of NO-Losartan A (0, 1, 5, 25, 125, 625, and 3,125 nM). The binding between PD-1 and PD-L1 was then assessed using a surface plasmon resonance (SPR) assay. (B) The inhibitory concentration (IC50) of NO-Losartan A could not be determined, as it failed to inhibit the PD-1/PD-L1 interaction. (C) Percentage of PD-1/PD-L1 interaction blockade by NO-Losartan A at all concentrations tested.
General notes and troubleshooting
General notes
This protocol outlines the procedure for the Biacore T200 system, which requires BIA evaluation software (version 4.1). While other SPR platforms from different brands can also be used, it should be noted that the specific steps may vary depending on the system’s operation. Key steps in this protocol include immobilizing a sufficient amount of PD-1 (more than 3500 RU) on the chip and running PD-L1 solution with a series of concentrations to obtain an observable RU.
Ensure there are no bubbles in any of the tubes. If there are bubbles, sonication can be performed to remove them.
Each control, BMS-1166, BMS-202, and NO-Losartan A were run separately. However, when running samples, they can be run in the same experiment as positive and negative controls. Furthermore, we evaluated the IC50 values of BMS-1166 (20–10,000 nM) and BMS-202 (20–10,000 nM) and found that they demonstrated similar IC50 values in Figure S1. Specifically, the IC50 value for BMS-1166 was 80.47 nM and the IC50 value for BMS-202 was 359.1 nM.
Technical replicates can be useful in this protocol; we recommend carefully considering the experimental design and the potential sources of variability when using SPR to investigate the blockade rates of different compounds using proteins from different sources. Depending on the nature of the experiment and the biological variability of the samples, it may be appropriate to include biological replicates to ensure accurate and reliable results.
The concentration of PD-L1 recombinant protein can vary depending on the RU. Although 20 nM of PD-L1 was used in this protocol and displayed an observable RU, a lower concentration may still be useful. However, it is not recommended to use PD-L1 with a concentration that leads to a RU of less than 10 RU, as this may result in an unfavorable signal-to-noise ratio. In addition, conducting technical replicates for this binding kinetics assay may not be necessary, as it is primarily used to confirm the suitability of the concentrations of PD-L1 for use in further blockade assays.
Use a final concentration of 0.01% DMSO, as this had the lowest percentage blockade effect on PD-1/PD-L1 binding (Table S1).
DMSO with different percentages significantly affected PD-L1 activities, as supported by the data displayed in the Supplementary materials. Try to use a minimum concentration of DMSO (e.g., 0.01% or lower) in the sample preparation and running buffer. Ensure the concentration of DMSO in the sample preparation and running buffer are equal to avoid buffer mismatch effects.
Troubleshooting
Unsuccessful immobilization can result from either the failure to immobilize PD-1 or a low final RU on the chip. This may be due to a low stock concentration of PD-1, as using a stock solution with a low concentration of PD-L1 in DNase-free water can affect the pH value of the working solution when PD-1 is dissolved in acetate 5.0.
Adjust the target response level (increasing it to 5,000 RU).
Change the running buffer or protein buffer (preparing a high concentration of PD-1 stock solution up to 1 mg/mL in DNase-free water).
Change to a different SPR chip (CM5 chip is used in this study, but the users can try the CM7 chip with a higher loading capacity to increase the amount of PD-1 on the chip).
Unsuccessful PD-1/PD-L1 interaction can result from an unsuccessful immobilization or a narrow concentration range of PD-L1.
a. Vary the protein concentrations (expand the PD-L1 concentration from nM to mM with a 10-fold dilution factor. This can help to identify the optimal concentration range for successful PD-1/PD-L1 interaction. Additionally, if unsuccessful immobilization is the cause, follow the steps outlined in the previous answer to address this issue).
Acknowledgments
Development of this protocol was supported in part by the instrumentation funded by the Rhode Island Institutional Development Award (IDeA) Network of Biomedical Research Excellence from the National Institute of General Medical Sciences of the National Institutes of Health under grant number P2GM10340. The protocols described here are adapted from our previous work (Jing et al., 2023).
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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Supplementary information
The following supporting information can be downloaded here:
Figure S1. Inhibition of PD-1/PD-L1 interaction by BMS-202 and BMS-1166
Table S1. Effects of DMSO 0, 0.01, 0.05, 0.1, 0.5, and 1% on PD-1/PD-L1 Binding
Supplementary materials
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Correction Notice: Assessment of Chemosensory Response to Volatile Compounds in Healthy, Aged, and Neurodegenerative Caenorhabditis elegans Models
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Roberto Grau
Published: Jun 5, 2023
DOI: 10.21769/BioProtoc.4766 Views: 156
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The word “individuals” has been added to the sentence "Here, we present a modified protocol to assess avoidance or attraction responses to diverse stimuli in healthy individuals and Caenorhabditis elegans models associated with neurodegenerative diseases" in the abstract section of “Assessment of Chemosensory Response to Volatile Compounds in Healthy, Aged, and Neurodegenerative Caenorhabditis elegans Models" (https://bio-protocol.org/e4650).
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A Novel Method for Measuring Mitochondrial Respiratory Parameters in Wheat Paleae (Paleae Superior) Using the XF24 Analyzer
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Abstract
Understanding the influence of secondary metabolites from fungi on the mitochondria of the host plant during infection is of great importance for the knowledge of fungus–plant interactions in general; it could help generate resistant plants in the future and in the development of specifically acting plant protection products. For this purpose, it must first be possible to record the mitochondrial parameters in the host plant. As of the date of this protocol, no measurements of mitochondrial respiration parameters have been performed in wheat paleae. The protocol shown here describes the measurements using the XF24 analyzer, which measures the rate of oxygen consumption in the sample by changes in the fluorescence of solid-state fluorophores. This procedure covers the preparation of samples for the XF24 analyzer and the measurement of mitochondrial parameters by adding specific mitochondrial inhibitors. It also shows the necessary approach and steps to be followed to obtain reliable, reproducible results. This is a robust protocol that allows the analysis of mitochondrial respiration directly in the wheat paleae. It demonstrates an important add-on method to existing screenings and also offers the possibility to test the effects of early infection of plants by harmful fungi (e.g., Fusarium graminearum) on mitochondrial respiration parameters.
Key features
• This protocol offers the possibility of testing the effects of early infection of plants by pathogens on mitochondrial respiration parameters.
• This protocol requires a Seahorse XF24 Flux Analyzer with Islet Capture Microplates and the Seahorse Capture Screen Insert Tool.
Graphical overview
Keywords: Oxygen consumption rate Mitochondrial respiration Wheat paleae Plant culture Seahorse XF24 Flux Analyzer
Background
The filamentous ascomycete Fusarium graminearum is the main pathogen causing Fusarium head blight (FHB) in wheat (McMullen et al., 1997). The consequences of host plant infection lead to significant quantitative and economic losses (Goswami and Kistler, 2004), as the infection results in incompletely formed grains. During infection, Fusarium graminearum produces a variety of secondary metabolites, low-molecular-weight molecules that are generally not necessary for the growth or developmental processes of the fungus but result in contamination of the grain with harmful mycotoxins (Shwab and Keller, 2008). Contaminated grain thus becomes unusable as food for humans and animals.
One of these mycotoxins is butenolide, which was first isolated in 1967 and shown to be toxic to mitochondrial respiration in rat cardiac muscle tissue (Wang et al., 2009). Further studies revealed that mitochondria of muscle tissue are affected. This includes swelling, disruption of the double membrane, and disruption of mitochondrial respiration (Wang et al., 2007 and 2009; Pei et al., 2013). Whether plant mitochondria are also affected by butenolide has not yet been investigated in detail. However, the impairment of animal mitochondria by butenolide could be an indication of a possible effect of this metabolite on plant mitochondria. Therefore, understanding the function of these substances on mitochondria during infection is of great importance for the knowledge of fungus–plant interaction in general and should help to produce resistant plants in the future.
Because the measurement of mitochondrial respiration in cells from plant cultures may not fully reflect the real situation, we describe here a method for measuring mitochondrial respiration parameters in wheat paleae (paleae superior) using an XF24 Extracellular Flux Analyzer. To date, no measurements of mitochondrial respiratory parameters have been performed in wheat paleae. The XF24 analyzer is a respirometer based on a multi-well plate that measures oxygen consumption rate (OCR) by changes in the fluorescence of solid-state fluorophores (Gerencser et al., 2009). To obtain a mitochondrial respiration profile, mitochondrial agents are automatically injected through various ports during testing (Divakaruni et al., 2014). This procedure involves preparation of samples for the XF24 analyzer and the measurement of mitochondrial parameters by addition of specific mitochondrial inhibitors.
The analysis of mitochondrial respiration in these samples is an important addition to existing studies and also offers the possibility to test the effects of early infection of plants with harmful fungi on mitochondrial respiration parameters. This could contribute to the development of resistant plants as well as specifically acting plant protection products. In addition to questions on resistance to fungal attack, this protocol can be used to investigate questions relating to mitochondrial activity in wheat in more detail or to use mitochondrial parameters as a supplement to classical phenotyping.
Materials and reagents
Centrifuge tubes 15 and 50 mL (Sarstedt, catalog numbers: 62.554.001, 62.547.254)
Petri dishes (Sarstedt, catalog number: 83.3902)
Pipette tips 10, 20, 200, and 1,000 μL (Sarstedt, catalog numbers: 70.1130, 70.116, 70.760.002, 70.762)
Sterile filter 0.2 μm (Sarstedt, catalog number: 83.1826.001)
Wheat paleae (kept on 1.6% agarose gel at 4 °C for a maximum of 36 h) (prevention of dehydration during transport)
KCl (Carl Roth, catalog number: 6781.3)
KH2PO4 (Carl Roth, catalog number: 3904.2)
Na2HPO4·12H2O (Carl Roth, catalog number: N350.1)
NaCl (Carl Roth, catalog number: 3957.3)
NaOH (Carl Roth, catalog number: 6771.3)
Seahorse XF calibrant solution (Agilent Technologies, catalog number: 100840-000)
Seahorse XF Cell Mito Stress Test kit [oligomycin, FCCP (Carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone), and rotenone/antimycin A] (Agilent Technologies, catalog number: 103015-100); store at -20 °C
Agarose standard (Carl Roth, catalog number: 3810.3)
Dulbecco’s modified Eagle’s medium, high glucose (Sigma-Aldrich, catalog number: D7777-10L), storage at 4 °C
10 M NaOH (for adjustment of pH values) (see Recipes)
1.6% agarose gel (see Recipes)
1× PBS (see Recipes)
XF assay medium (see Recipes)
Equipment
Scalpel, stainless steel (Carl Roth, catalog number: T997.1)
Seahorse Capture Screen Insert Tool (Agilent Technologies, catalog number: 101135-100)
Seahorse XF24 Islet Capture FluxPak containing sensor cartridges and Islet Capture Microplates (Agilent Technologies, catalog number: 101174-100)
Tweezers, stainless steel (Carl Roth, catalog number: 2687.1)
Incubator at 37 °C without CO2 (GFL, catalog number: 4010)
pH Meter FiveEasyTM F20 (Mettler Toledo, catalog number: 30266626)
Pipettes, Eppendorf Research® Plus 10, 20, 200, and 1,000 μL (Eppendorf, catalog numbers: 3123000020, 3123000039, 3123000055, 3123000063)
Seahorse XF24 Flux Analyzer (Agilent Technologies, model: XF24, catalog number: 100737-100)
Software and datasets
Seahorse XF24 Flux Analyzer Software (instrument software) (Agilent Technologies, Version 1.8.1.1)
Wave (analysis software) (Agilent Technologies, Version 2.6.1)
GraphPad Prism (statistical software) (GraphPad Software, Inc., Version 9.5.1)
Procedure
Day before the assay
Hydrate the sensor cartridge in Seahorse XF calibrant solution according to the manufacturer’s instructions one day prior to measurement in a non-CO2 incubator.
Turn on the XF24 analyzer to preheat the system to 37 °C.
Day of the assay
Sample preparation
The previously cultivated wheat flowers (see Notes) are separated from a spike with a scalpel and the wheat paleae is then isolated from the flowers (Figure 1).
Wash the wheat paleae five times in 1× PBS.
After this step, the wheat paleae can be used directly for measurement or stored in 1.6% agarose at 4 °C until further use. The storage time at this temperature is seven days maximum (see Notes).
Figure 1. Structure of a wheat flower. P = Paleae, L = Lemma, K = Caryopsis, G = Gluma. Scale bar = 2 mm. Figure from Glasenapp (2017).
Assay
Prepare the XF assay medium and heat to 37 °C.
Take the wheat flower from the 1× PBS or agarose gel and wash it again five times in 1× PBS to remove any remaining impurities or residues of agarose (Figure 2A).
After removing the lower as well as the upper part of the wheat paleae, wash again in 1× PBS (Figure 2C).
Put the wheat paleae with its concave side (side with the wings) down, on a net [Figure 2B, 2C, and 2G (7–9)].
It is important to ensure that the wings of the wheat paleae side are facing down towards the capture screen. This orientation is essential for the measurement, as the side facing the capture screen gets detected by the sensor (Figure 2G).
Add 450 μL of XF assay medium into each well of the Islet Capture microplate.
Incubate the Islet Capture microplate in the incubator (without CO2) at 37 ° C for 45 min to adjust the wheat paleae to the new XF assay medium.
Load the injection ports of the sensor cartridge as programmed (see the programming of the XF controller), follow the instructions of the XF Analyzer, and start the calibration of the sensor cartridge.
When calibration is complete, place the Islet Capture microplate in the XF Analyzer and start the measurement.
Data analysis is performed with the Seahorse analysis software Wave.
Figure 2. Microscopic image of wheat paleae in the XF24 Islet Capture microplate. The wheat paleae (A) must be placed on the net such that its upper side with its so-called wings points downwards in contact with the capture screen. The sample must then be placed in the multi-well plate, using the capture screen insert tool (B, C). Top view (D) and view from below (E) of wheat paleae inserted into the multi-well plate. The processed wheat paleae should cover the entire surface of the capture screen to ensure that the same area is always measured (D–F). The measurement of oxygen consumption takes place over the surface within the ring marked with the arrow (F). Schematic setup of the XF24 measuring system with an Islet Capture microplate (G). Scale bar = 2 mm.
Programming the XF controller
This section describes the individual program steps of the XF controller for measuring the OCR as shown in Figures 3 and 4. The stress reagents are diluted to a final concentration of 4 μM with XF assay medium. The concentration of oligomycin, FCCP, and rotenone/antimycin A should be determined in a preliminary experiment for the respective experimental conditions. The injection of the stress reagent should be simultaneously carried out in all wells. All measurements are done in several cycles. Each cycle involves 4 min mixing, 2 min waiting, and 3 min OCR and extracellular acidification rate (ECAR).
Workflow of the XF controller
Calibrate probes.
Time of Delay: 15 min.
First Loop: 3 times (Basal state).
Mixing: 4 min
Time of Delay: 2 min
Measurement: 3 min
Injection of Port A (50 μL of oligomycin, 4 μM final concentration).
Use oligomycin to block the ATP synthase.
Then, the decrease in OCR corresponds to the amount of oxygen used for ATP synthesis.
Second Loop: 6 times (decrease of ATP-linked respiration)
Mixing: 4 min
Time of Delay: 2 min
Measurement: 3 min
Injection of Port B (55 μL of FCCP, 4 μM final concentration).
The decoupling of the respiratory chain by FCCP allows the maximal respiration and the spare respiratory capacity to be determined independently of the proton gradient.
Third Loop: 3 times (potentially uncoupled respiration).
Mixing: 4 min
Time of Delay: 2 min
Measurement: 3 min
Injection of Port C (60 μL of rotenone/antimycin A, 4 μM final concentration).
In the last Loop, complexes I and II are blocked by the addition of rotenone and antimycin A. The remaining oxygen consumption thus depends only on the non-mitochondrial respiration.
Fourth Loop: 6 times (non-mitochondrial respiration).
Mixing: 4 min
Time of Delay: 2 min
Measurement: 3 min
End of Program.
Data analysis
Data can be analyzed using GraphPad Prism 9.5.1 (GraphPad Software, Inc.). Values are presented as mean ± SEM. Five replicates per experimental group are recommended.
Validation of protocol
This method is routinely used in our laboratory. N = 20+. Data are analyzed using Prism 9.5.1 (GraphPad Software, Inc.). Values are presented as mean ± SEM or individual values. Five replicates per experimental group are recommended. Background wells (distributed over the plate to exclude temperature effects) and wildtype wheat paleae with and without inhibitors are used as controls. Also, different orientations of wheat paleae were applied in the experimental setup. Prior to the use of this assay, extensive establishment and validation work was carried out over a period of several weeks. This protocol was published in the PhD thesis “Studies on early infection of wheat Paleae by Fusarium graminearum” by the Hamburg University and additionally presented at international congresses (Glasenapp, 2017).
Representative data
Measurement of OCR in wheat paleae according to the proceeding steps (Figure 3) shows the typical course after addition of the four stress reagents oligomycin, FCCP, and rotenone/antimycin A as depicted by the black curve in Figure 4.
Figure 3. Exemplary XF24 measurement using the XF Cell Mito Stress Test kit (Agilent Technologies). Shown is the oxygen consumption rate (OCR; pmoles/min) as a function of time (minutes). First, the basal respiration is recorded. The addition of oligomycin determines proton leakage and ATP-linked respiration. FCCP is used to measure maximal respiration. Antimycin A and rotenone block respiration of the mitochondria, to determine non-mitochondrial respiration. Figure created after the template of Agilent Technologies (Agilent Technologies. Retrieved October 15, 2019, from https://www.agilent.com/en/products/cell-analysis/seahorse-xf-consumables/kitsreagents-media/seahorse-xf-cell-mito-stress-test-kit#additionalinformation).
Figure 4. Oxygen consumption rate (OCR) in wheat paleae. Scheme depicting the main stages after injecting the four active ingredients in three injecting steps. This makes it possible to determine the efficiency of the individual complexes within the respiratory chain in wildtype wheat paleae (WT) (n = 10, mean ± SEM) and wheat paleae infected with Fusarium graminearum (n = 3, mean ± SEM).
Notes
The measurement of OCR in wheat paleae described here was highly reproducible. Also, different orientations of wheat paleae were applied in the experimental setup. Prior to the use of this assay, extensive establishment and validation work was carried out over a period of several weeks.
The concentration of oligomycin, FCCP, and rotenone/antimycin A should be determined in a preliminary experiment for the respective experimental conditions. The indicated concentrations may differ for other samples. In preliminary studies, the concentration of 4 μM was determined. Lower concentrations show, in contrast to cell cultures, no or little effects.
Because wheat paleae are a complex cell system in which the substances simply take longer to penetrate the entire sample, a higher concentration and more measurement points, as stated in the Seahorse manual, must be used.
After removing the wheat paleae from the rest of the plant, these can be embedded in 1.6% agarose gel and stored at 4 °C. The samples should be stored for a maximum of seven days. It is important to perform the OCR measurement during this period, as the respiration parameters decrease significantly after.
The orientation of the wheat paleae in the measurement chamber is important because the wings are the most accessible for measuring mitochondrial respiration. Thus, the wings should be oriented in the direction of the sensors (Figure 2G). This is confirmed in samples where the wings are removed or placed facing away from the sensor (Figure 5).
Figure 5. Oxygen consumption rate (OCR) in wheat paleae of different orientations. Shown are the curves of wheat paleae with different orientation or removal of the wings in the Seahorse XF24 Flux Analyzer (n = 5, mean ± SEM).
The wheat cultivar Nandu (Lochow-Petkus, Bergen-Wohlde, Germany) used for the studies was cultivated in a growth chamber for approximately six weeks at 18–20 °C with 70% relative humidity and a photoperiod of 16 h until flowering.
Based on the measured OCR values (Figure 4), further mitochondrial parameters such as basal respiration, ATP production, but also maximum respiration or proton leak can be analyzed using the Seahorse software (Wave) (Figure 3).
The only limitation of this protocol is the requirement of Seahorse XF24 Flux Analyzer with Islet Capture microplates and the Seahorse Capture Screen Insert Tool equipment.
The Islet Capture microplate includes 24 wells, leaving four wells empty to be used as background.
Recipes
10 M NaOH
40 g of NaOH
add 100 mL of ddH2O
1.6% agarose gel
1.6 g of agarose
add 100 mL of ddH2O
1× PBS
8 g of NaCl
0.20 g of KCl
2.88 g of Na2HPO4·12H2O
1.24 g of KH2PO4
add 1 L of ddH2O, pH 7.4
XF assay medium
0.675 g of Dulbecco’s modified Eagle’s medium, high glucose
add 50 mL of ddH2O, sterile filtering, pH 7.4
Acknowledgments
The authors thank Dr. Anika Glasenapp for the kind supply of samples. They also thank Dr. Wilhelm Schäfer for scientific advice. This study was supported by the BMBFFHprofUnt2012 “MitoFunk” (03FH022PX2) and by the Baden-Württemberg Ministry of Science, Research and Art. This protocol was modified based on the previous work from Schniertshauer et al (2019).
Competing interests
One of the authors has a consulting contract with MSE Pharmazeutika GmbH, Bad Homburg, Germany.
References
Divakaruni, A. S., Paradyse, A., Ferrick, D. A., Murphy, A. N. and Jastroch, M. (2014). Analysis and Interpretation of Microplate-Based Oxygen Consumption and pH Data. Methods Enzymol. 547: 309–354.
Gerencser, A. A., Neilson, A., Choi, S. W., Edman, U., Yadava, N., Oh, R. J., Ferrick, D. A., Nicholls, D. G. and Brand, M. D. (2009). Quantitative Microplate-Based Respirometry with Correction for Oxygen Diffusion. Anal. Chem. 81(16): 6868–6878.
Goswami, R. S. and Kistler, H. C. (2004). Heading for disaster: Fusarium graminearum on cereal crops. Mol. Plant Pathol 5(6): 515–525.
Glasenapp, A. (2017). Studies on early infection of wheat Paleae by Fusarium graminearum. PhD thesis. Hamburg University.
McMullen, M., Jones, R. and Gallenberg, D. (1997). Scab of Wheat and Barley: A Re-emerging Disease of Devastating Impact. Plant Disease 81(12): 1340–1348.
Pei, J., Fu, W., Yang, L., Zhang, Z. and Liu, Y. (2013). Oxidative Stress Is Involved in the Pathogenesis of Keshan Disease (an Endemic Dilated Cardiomyopathy) in China. Oxid. Med. Cell. Longevity 2013: 1–5.
Shwab, E. K. and Keller, N. P. (2008). Regulation of secondary metabolite production in filamentous ascomycetes. Mycol. Res. 112(2): 225–230.
Schniertshauer, D., Gebhard, D. and Bergemann, J. (2019). A New Efficient Method for Measuring Oxygen Consumption Rate Directly ex vivo in Human Epidermal Biopsies. Bio-protocol 9(5): e3185.
Wang, Y. M., Peng, S. Q., Zhou, Q., Wang, M. W., Yan, C. H., Wang, G. Q. and Yang, H. Y. (2007). The oxidative damage of butenolide to isolated erythrocyte membranes. Toxicol. In Vitro 21(5): 863–869.
Wang, Y. M., Liu, J. B. and Peng, S. Q. (2009). Effects of Fusarium Mycotoxin Butenolide on Myocardial Mitochondria In Vitro. Toxicol. Mech. Methods 19(2): 79–85.
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Improved Methods for Acetocarmine and Haematoxylin Staining to Visualize Chromosomes in the Filamentous Green Alga Zygnema (Charophyta)
NR Nina Rittmeier
AH Andreas Holzinger
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4768 Views: 402
Reviewed by: Dennis J NürnbergMercedes Nieves MoriónShuhei Ota
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Original Research Article:
The authors used this protocol in bioRxiv Feb 2023
Abstract
Genome sizes of Zygnema spp. vary greatly, being unknown whether polyploidization occurred. The exact number of chromosomes in this genus is unknown since counting methods established for higher plants cannot be applied to green algae. The massive presence of pectins and arabinogalactan proteins in the cell wall interferes with the uptake of staining solutions; moreover, cell divisions in green algae are not restricted to meristems as in higher plants, which is another limiting factor. Cell divisions occur randomly in the thallus, due to the intercalary growth of algal filaments. Therefore, we increased the number of cell divisions via synchronization by changing the light cycle (10:14 h light/dark). The number of observed mitotic stages peaked at the beginning of the dark cycle. This protocol describes two methods for the visualization of chromosomes in the filamentous green alga Zygnema. Existing protocols were modified, leading to improved acetocarmine and haematoxylin staining methods as investigated by light microscopy. A freeze-shattering approach with liquid nitrogen was applied to increase the accessibility of the haematoxylin dye. These modified protocols allowed reliable chromosome counting in the genus Zygnema.
Key features
• Improved method for chromosome staining in filamentous green algae.
• Optimized for the Zygnema strains SAG 698-1a (Z. cylindricum), SAG 698-1b (Z. circumcarinatum), and SAG 2419 (Zygnema ‘Saalach’).
•This protocol builds upon the methods of chromosomal staining in green algae developed by Wittmann (1965), Staker (1971), and Fujii and Guerra (1998).
• Cultivation and synchronization: 14 days; fixation and permeabilization: 24 h; staining: 1 h; image analysis and chromosome number quantification: up to 20 h.
Keywords: Acetocarmine Haematoxylin Hydroxyquinoline Chromosomes Zygnema Zygnematophyceae Charophyta Light microscopy
Background
Background on chromosome counting in Zygnema
Zygnematophyceae are the immediate sister group to land plants, thought to have colonized land 550 million years ago (Wodniok et al., 2011; Leebens-Mack et al., 2019). Data on nuclear DNA content are available for Zygnema (Čertnerová, 2021; Feng et al., 2021 and 2023), and genome sizes for different strains of Zygnema spp. have been established, but these show great variation (Feng et al., 2023). Chromosomes of Zygnema have previously been illustrated by transmission electron microscopy (Bakker and Lokhorst, 1987), although without giving actual numbers. The actual chromosome numbers of Zygnema vary drastically, ranging from 14 to 82 (Kurssanow, 1911; Miller 1973; Guiry et al., 2022). Thus, it is unclear if polyploidization has happened. Traditional counting methods established for higher plants are hampered in green algae by pectins [massive homogalacturonan accumulations have been reported by Herburger et al. (2019)] and arabinogalactan proteins in the cell wall. Even though numerous protocols have been established for chromosomal staining in green algae (Godward, 1948; Wittmann, 1965; Prasad and Godward, 1966; Staker, 1971; Gerlach, 1977; Fujii and Guerra, 1998), they still face difficulties handling Zygnema spp., as their chromosomes are small, sticky, and mostly only able to be counted in their mid- to late-prophase. Furthermore, conventional staining methods lead to over-staining of the cell wall and DNA-dense areas in the cytoplasm known as karyoide in Zygnema, Charophyta (Kopetzky-Rechtperg, 1934).
Modification of the methods from Wittmann (1965), Staker (1971), and Fujii and Guerra (1998)
For all three Zygnema ssp., two different modified staining procedures were used: (1) acetocarmine (Figure 1A–1E) and (2) haematoxylin (Figure 1A, 1F–1L). To achieve full mitotic synchronization in Zygnema, a light/dark cycle (10:14 h) was applied following Staker (1971), i.e., with a longer dark period than our standard cultivation cycle (light/dark, 16:8 h).
Figure 1. Summary of the experimental procedure for acetocarmine (B–E) and haematoxylin (F–L) staining in Zygnema circumcarinatum.(A) Erlenmeyer flask with young culture; (B) 8-hydroxyquinoline prefixation; (C, F) fixation in Carnoy’s fluid, illustrating the gradual bleaching from time 0, 1, and 12 h (from left to right); (D) staining with acetocarmine; (E) projected Z-stack rendered by Helicon Focus software used for counting the chromosomes (circle: chromosomes marked with green crosses in ImageJ; inset: symbolic representation of individual images for Z-stacks); (G) algal biomass mounted in acetic acid and squished between two slides; (H) slides dipped in liquid nitrogen; (I) HCl treatment; (J, K) aceto-haematoxylin-iron alum staining; (L) light microscopic image used for quantifying the number of chromosomes (circle). Scale bars = 10 µm.
For the staining of the chromosomes, Staker (1971) used propriocarmine, which was replaced with 1% acetocarmine in the current protocol; also, the Zygnema filaments were not chopped or randomized, which would lead to complete destruction. Moreover, prior to the fixation in Carnoy’s fluid, cells were treated with 8-hydroxyquinoline to depolymerize microtubules, resulting in sticky and condensed metaphase chromosomes (Bukhari, 2004). Haematoxylin staining by Wittmann (1965) was used, but the treatment with chloralhydrate and slide heating steps was omitted and replaced with hydrolysis by 5 N HCl (Fujii and Guerra, 1998), resulting in contrast improvement between chromosomes and cytoplasm. Changes only occurred with the algal filaments being placed between two microscopic slides before liquid nitrogen treatment with a freeze-shattering method (Wasteneys et al., 1997); the HCl treatment time was reduced to 10 min to assure the integrity of the Zygnema filaments, which disintegrate after prolonged treatment.
Materials and reagents
Biological material
Zygnema cylindricum strain SAG 698-1a (Figure 2A; Feng et al., 2021; isolated 1929 by Czurda V.; deposited 1954 by Pringshein E.G.), collected from a ditch at meadow Poselteich (Polenský Rybnik; 50°33′09.7″N 14°40′09.7″E) near Hirschberg (Dosky) in Czech Republic, Europe.
Figure 2. Chromosomes (circles) visualized by light microscopy in different Zygnema strains. (A, D, G) Z. cylindricum; (B, E, H) Z. circumcarinatum; and (C, F, I) Z. Saalach. (A, B, C) Living cells without staining; (D, E, F) acetocarmine staining; (G, H, I) haematoxylin staining. Insets in E, G, and I show manual drawings of chromosomes counted with ImageJ; counted chromosome numbers: n = 20 for Z. cylindricum/Z. circumcarinatum and n = 30 for Z. Saalach. Scale bars = 20 µm.
Zygnema circumcarinatum strain SAG 698-1b (Figure 2B; isolated 1929 by Czurda V.; deposited 1954 by Pringshein E.G.), collected from a ditch at meadow Poselteich (Polenský Rybnik; 50°33′09.7"N 14°40′09.7″E) near Hirschberg (Dosky) in Czech Republic, Europe. The results of the chromosome counting for this strain have been recently published (Feng et al., 2023)
Zygnema ‘Saalach’ (SAG 2419; 47°47′8.70″N, 12°56′42.66″E; 440 m above sea level; Figure 2C), collected near Salzburg, Austria (Herburger et al., 2015)
Reagents
8-Hydroxyquinoline (Merck, catalog number: 148-24-3)
Acetocarmine (Morphisto, catalog number: 10411)
Liquid nitrogen
5 N HCl (Merck, catalog number: 258148)
Bi-distilled water (A. bidest)
Bold’s basal culture medium (BBM), pH 5.5 (Bischoff and Bold, 1963)
Glacial acetic acid (Merck, catalog number: A6283)
100% ethanol (Sigma, catalog number: 493546)
45% acetic acid (Merck, catalog number: A6283)
Haematoxylin (Merck, catalog number: H9627)
Ammonium iron (III) sulfate (Sigma, catalog number: 221260)
Solutions
2 mM 8-Hydroxyquinoline (see Recipes)
Carnoy’s fluid (see Recipes)
Aceto-haematoxylin-iron alum (see Recipes)
Recipes
2 mM 8-Hydroxyquinoline
Reagent Final concentration Quantity
8-Hydroxyquinoline 2 mM 29 mg
A. bidest 100% 1,000 mL
Total 1,000 mL
This solution can be kept in the dark at room temperature (RT) for up to one year and can last for up to 100 preparations.
Carnoy’s fluid
Reagent Final concentration Quantity
Glacial acetic acid 100% 125 mL
Ethanol (absolute) 100% 375 mL
Total 500 mL
This solution should be prepared immediately before use and can last for up to 50 preparations.
Aceto-haematoxylin-iron alum
Reagent Final concentration Quantity
Acetic acid 45% 100 mL
Haematoxylin 0.4% 400 mg
Ammonium iron (III) sulfate 0.1% 100 mg
Total 100 mL
This solution can be kept at 4 °C for up to half a year and can last for up to 300 preparations.
Equipment
Gas burner
Fridge (4 °C)
Growth chamber (Panasonic, MLR-352-PE equipped with 2 Panasonic FL40SS·ENW/37 fluorescent tubes)
Light microscope [Zeiss Axiovert 200M microscope equipped with a 100×, 1.3 NA objective lens (Carl Zeiss AG)] with a Zeiss high-resolution AxioCam HRm Rev.3 camera
Glassware: 250–500 mL Erlenmeyer flask (Analyticsshop.com, ID1121226361), 10 mL glass vials (Merck, 27151), 10 mL test tubes (Merck, Z741001), culture dish (Analyticsshop.com, ø 100 mm), glass jar (for storage)
Glass Pasteur pipettes (Analytics Shop.com, BR747715)
LLG-Syringe filters, CA, 0.20 m, ø 13 mm (Lab logistics Group, 14140027207)
1 mL Syringe, Omnifix®-F (Bio-apo.at, 00569881)
2 mL tubes
Lint-free paper
Metal rack
Microscopic slides and coverslips
Wooden clip
Pair of fine-pointed tweezers
Liquid nitrogen container with lid
Leather gloves
Long tongs
Spray skirt
Safety goggles
Stopwatch or timer
Software
Helicon Focus (HeliconSoft Ltd.)
ImageJ (1.53v)
Procedure
Acetocarmine staining
Synchronization of mitotic activity
Place pure algal biomass into a 250–500 mL Erlenmeyer flask containing 150–250 mL of BBM (Figure 1A).
Grow cultures in a light/dark regime of 10:14 h at 20 °C and 50 μmol photons m-2·s-1 in the light period for two to three weeks to obtain log-phase cultures.
Harvesting of algal biomass
Collect algal biomass at the beginning of the dark cycle in the laminar flow hood (to maintain sterility of the cultures).
Place the sample with the fine-pointed tweezers into 2 mL tubes containing 1 mL of A. bidest at RT.
Fix collected samples immediately after collection.
Fixation
Transfer the samples to 10 mL glass vials containing 5 mL of 2 mM 8-hydroxyquinoline, leave at RT for 1–2 h under the fume hood, and then transfer to 4 °C for 1–2 h (Figure 1B).
Note: The necessary time has to be tested out for each species. For SAG 698-1a and SAG 698-1b, 1 h each was enough, while for SAG 2419, 2 h was needed.
Remove 8-hydroxyquinoline completely with a glass Pasteur pipette and wash the sample with 5 mL of A. bidest three times for at least 1 min under the fume hood.
Immediately submerge samples in 5 mL of Carnoy’s fluid and leave at RT for 12 h until all chlorophyll is removed and the samples are visibly bleached (Figure 1C).
Staining
Collect the current bleached sample with the fine-pointed tweezers and place in the test tube containing 5 mL of 1% acetocarmine.
Hold the test tube with the wooden clip over the gas burner at low flame by not constantly keeping it in the flame and boil the algal biomass in acetocarmine for 5 min under the fume hood (Figure 1D).
Pour the acetocarmine-boiled algal biomass into the culture dish. Select the stained filaments with fine-pointed tweezers, place them onto a microscopic slide with a small droplet of acetocarmine, and place a coverslip on top.
Microscopical analysis
Visualize the stained chromosomes with a light microscope (Figure 1E).
Take 50–100 images per area in Z-direction.
Note: Either use an automated focus or capture images manually with a distance of 0.2–0.4 µm.
Render stacked models with the software Helicon Focus (HeliconSoft Ltd.).
Count the chromosomes with ImageJ.
Notes:
i. The following tools in ImageJ should be used to process the stacked images; for details, see the following YouTube video by Kevin Foley: https://m.youtube.com/watch?v=D1qBaFwuF4E.
Process - Subtract Background
Image - Adjust - Threshold
Process - Binary - Fill Holes
Process - Binary - Convert to Mask
Process – Binary- Watershed
Analyze - Analyze Particles
ii. The number of chromosomes counted is based on a minimum of three biological replicates. In Figure 2D, 2E, and 2F, representative samples are illustrated. For each biological replicate, the chromosomes of at least 10 samples were counted (technical replicates).
Haematoxylin staining
Synchronization of mitotic activity
Place pure algal biomass into 250–500 mL tubes containing 200 mL of BBM (Figure 1A).
Grow cultures in a light/dark regime of 10:14 h at 20 °C and 50 μmol photons m-2·s-1 in the light period for at least two weeks.
Harvesting of algal biomass
Collect algal biomass at the beginning of the dark cycle.
Place the sample with fine-pointed tweezers into 2 mL tubes containing 1 mL of A. bidest at RT.
Fix collected samples immediately.
Fixation
Place harvested material into the 10 mL glass vial containing 5 mL of Carnoy’s fluid and incubate for 2 h or until sample is completely bleached at RT (Figure 1F).
Decant the Carnoy’s fluid and replace with 5 mL of 70% ethanol (samples can be stored in this mixture at 4 °C for up to half a year).
Wash samples in 5 mL of A. bidest three times for at least 1 min before transferring the bleached material to a microscopic slide (mount in 45% acetic acid) and squashing with another slide (Figure 1G).
Liquid nitrogen treatment
Fill the liquid nitrogen container halfway up with liquid nitrogen (wear safety goggles, leather gloves, and a spray skirt for the whole procedure) and place lid on top.
Open the container and use the long tongs to dip the two slides for ten seconds into the liquid nitrogen (Figure 1H).
Remove the slides from the liquid nitrogen, pull them apart carefully while still frozen, and let them air dry for at least 30 min.
Note: The slides can be pulled apart by hand or, if proven difficult, a spatula can be used.
Let the leftover liquid nitrogen evaporate under the fume hood.
HCl treatment
Plunge the dried slide with the samples attached into 5 N HCl for at least 10 min under the fume hood (Figure 1I).
Remove the slide and let it air dry for another 30 min.
Stain the dried slides immediately or keep them in a glass jar at -20 °C for up to half a year.
Staining
To stain the material on the microscopic slides, mount it with one droplet of the aceto-haematoxylin-iron alum (Figure 1J) and place a coverslip on top.
Note: A syringe equipped with a syringe filter is used to minimize the fallout particles of the solution, which could lead to contamination of the sample.
Full saturation is reached after an incubation time of 5 min (Figure 1K); the excessive dye can be removed with a lint-free paper.
Microscopical analysis
Visualize the stained chromosomes with a light microscope (Figure 1L).
Take up to 100 images per area in Z-direction.
Render stacked models with the software Helicon Focus (HeliconSoft Ltd.).
Count the chromosomes with ImageJ (see Section A, step 5d).
Note: The numbers of chromosomes counted are based on a minimum of three biological replicates; in Figure 2G, 2H, and 2I, representative samples are illustrated. Per biological replicate, the chromosomes of at least 10 samples were counted (technical replicates).
Acknowledgments
We would like to acknowledge that this protocol is adapted from the previous work of Wittmann (1965), Staker (1971), and Fujii and Guerra (1998). We would also like to acknowledge the help of Clemens Maylandt, Charlotte Permann, and Gregor Pichler, University of Innsbruck, in improving the protocols. The study was supported by Austrian Science Fund (FWF) project P 34181-B to A.H.
References
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Bukhari, Y. M. (2004). A simple method of chromosome preparation for Acacia and Prosopis (Mimosaceae). Hereditas 126(2): 195–197.
Čertnerová, D. (2021). Nuclei isolation protocols for flow cytometry allowing nuclear DNA content estimation in problematic microalgal groups. J. Appl. Phycol. 33(4): 2057–2067.
Feng, X., Holzinger, A., Permann, C., Anderson, D. and Yin, Y. (2021). Characterization of two Zygnema strains (Zygnema circumcarinatum SAG 698-1a and SAG 698-1b) and a rapid method to estimate nuclear genome size of Zygnematophycean green algae. Front. Plant Sci. 12: e610381.
Feng, X., Zheng, J., Irisarri, I., Yu, H., Zheng, B., Ali, Z., de Vries, S., Keller, J., Fürst-Jansen, J. M., Dadras, A., et al. (2023). Chromosome-level genomes of multicellular algal sisters to land plants illuminate signaling network evolution. bioRxiv: e526407.
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Godward, M. B. E. (1948). The Iron Alum Acetocarmine Method for Algæ. Nature 161(4084): 203.
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Herburger, K., Xin, A. and Holzinger, A. (2019). Homogalacturonan Accumulation in Cell Walls of the Green Alga Zygnema sp. (Charophyta) Increases Desiccation Resistance. Front. Plant Sci. 10: e00540.
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Kurssanow, L. (1911). Über Befruchtung, Reifung und Keimung bei Zygnema. Flora Allg. Bot. Ztg. 104(1): 65–84.
Leebens-Mack, J. H., Barker, M. S., Carpenter, E. J., Deyholos, M. K., Gitzendanner, M. A., Graham, S. W., et al. (2019). One thousand plant transcriptomes and the phylogenomics of green plants. Nature 574(7780): 679–685.
Miller, R. (1973). A developmental and physiological comparison of two mating strains of Zygnema circumcarinatum (Cuzrda). The University of Arizona.
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A Semi-throughput Procedure for Assaying Plant NADP-malate Dehydrogenase Activity Using a Plate Reader
KB Kevin Baudry
EI Emmanuelle Issakidis-Bourguet
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4769 Views: 322
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Original Research Article:
The authors used this protocol in Plant Physiology Aug 2022
Abstract
Chloroplast NADP-dependent malate dehydrogenase (NADP-MDH) is a redox regulated enzyme playing an important role in plant redox homeostasis. Leaf NADP-MDH activation level is considered a proxy for the chloroplast redox status. NADP-MDH enzyme activity is commonly assayed spectrophotometrically by following oxaloacetate-dependent NADPH oxidation at 340 nm. We have developed a plate-adapted protocol to monitor NADP-MDH activity allowing faster data production and lower reagent consumption compared to the classic cuvette format of a spectrophotometer. We provide a detailed procedure to assay NADP-MDH activity and measure the enzyme activation state in purified protein preparations or in leaf extracts. This protocol is provided together with a semi-automatized data analysis procedure using an R script.
Keywords: NADP-malate dehydrogenase NADP-MDH Enzyme activity assay Thioredoxin TRX R script Spectrophotometry
Background
Chloroplast NADP-dependent malate dehydrogenase (NADP-MDH) catalyzes the reduction of oxaloacetate to malate using NADPH. To be active, this enzyme needs to be reduced by thioredoxins (TRX), ubiquitous thiol-disulfide oxidoreductases (Issakidis et al., 1994; Collin et al., 2003). In C3 plants, NADP-MDH is involved in the export of reducing power from the chloroplast to the cytosol via the malate valve (Scheibe, 2004). TRX-dependent activation of NADP-MDH makes the link between the chloroplast electron transport chain, the redox state of the chloroplast, and the other cell compartments (Scheibe and Dietz, 2012; Heyno et al., 2014). Hence, the redox state of NADP-MDH is considered as a proxy for the plant leaf cellular redox state. NADP-MDH redox state in protein preparations or in plant extracts is deduced from the ratio between initial/extractable (i.e., activity of the enzyme or extract, without pre-treatment) and maximal activity/enzyme capacity (i.e., activity of the enzyme or in the extract, after reductive activation by TRX) (Issakidis et al., 1994; Keryer et al., 2004).
NADP-MDH activity can be easily assayed spectrophotometrically by monitoring oxaloacetate-dependent NADPH oxidation at 340 nm (Jacquot et al., 1995). Here, we developed a plate-adapted protocol to assay NADP-MDH activity of a large number of samples at the same time, associated with a semi-automatized data analysis procedure using a user-friendly R script. Compared with the classic method in a 1 mL spectrophotometer cuvette, the plate format allows increasing the experimental replicates and/or tested samples or conditions, for a gain of time (estimated divided by three), and in data precision and reliability, at a lower cost (divided by five). We implemented this method to measure NADP-MDH activity in Arabidopsis leaf protein extracts and using purified preparations of recombinant sorghum NADP-MDH. Our method is applicable to measure the activity of virtually any plant species.
Materials and reagents
General materials and reagents
Oxaloacetic acid (OAA) (Sigma-Aldrich, catalog number: O-4126)
β-Nicotinamide adenine dinucleotide 2’-phosphate reduced (NADPH) (Roth, catalog number: AE14.3)
Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D9779)
Tris pH 7.9
Classic PCR plate 96 wells (Thermo Fisher Scientific, catalog number: AB0700)
Plate 96 wells (Genetix X6011 96well)
Recombinant TRX m type (stored at -20 °C) (Collin et al., 2003)
Materials and reagents specific for in vitro assay
Recombinant NADP-MDH (stored at -20 °C) (Issakidis et al., 1994)
Materials and reagents specific for ex planta assay
4–5-week-old Arabidopsis plants
Metallic beads (3 mm diameter)
Protease inhibitor cocktail for plant (Sigma-Aldrich, catalog number: P9599)
Tris pH 6.8
QubitTM Protein and Protein Broad Range (BR) Assay kits (Thermo Fisher Scientific, catalog number: Q33212)
Solutions
Extraction buffer for ex planta assay (see Recipes)
Activation medium for ex planta assay (see Recipes)
Activation medium for the in vitro assay (see Recipes)
Reaction medium (see Recipes)
Equipment
Tecan infinite m200 PRO plate reader (Tecan) with a 230–1,000 nm wavelength range
Multichannel pipettes, 12 channels able to pipette 2–50 μL (Eppendorf)
General-purpose tweezers (Fisher Scientific, catalog number: 17-467-231)
Thermal cycler for incubation (Applied Biosystems 2720 Thermal Cycler)
Qubit 2.0 fluorometer (Thermo Fisher Scientific, Q32866) (for ex planta assay) (see Note 1)
Refrigerated microcentrifuge (Thermo Scientific, catalog number: 75-772-441)
Tissue Lyser II bead mill (Qiagen, catalog number: 85300)
Software
R (https://www.r-project.org/) version 3.6.3 or later
RStudio (https://www.rstudio.com/) version 2022.07.1+554 or later
Excel (Microsoft)
Notepad++ (https://notepad-plus-plus.org/) or any software to read text files
Magellan version 7.2 (https://lifesciences.tecan.com/software-magellan)
Procedure
An overview of the procedure and data analysis workflow is presented in Figure 1.
For NADP-MDH ex planta activity, follow sections A and C. For recombinant NADP-MDH activity (in vitro) assay, follow sections B and C. As a general advice, because this protocol uses plates and can measure up to 12 samples at the same time, triplicate assays can be easily performed in parallel for each sample type and for each experimental condition (initial or maximal activity for ex planta activity, activation kinetics for recombinant enzyme).
Figure 1. Overview of the procedure and data analysis workflow. Grey circles refer to corresponding procedure sections. Blue circles refer to corresponding data analysis sections. For NADP-MDH ex planta activity, follow procedures A and C and then analysis A, B and C. For NADP-MDH in vitro assay, follow procedures B and C and then analysis A, B and D.
Arabidopsis NADP-MDH extraction and activation, ex planta assay (alternative for B)
Extraction from leaf samples:
Using tweezers, collect one young adult leaf from a 4–5-week-old Arabidopsis plant in a collecting tube containing two metallic beads. Flash freeze samples in liquid nitrogen (see Note 2).
Grind sample in collecting tube by shaking using a bead mill (1 min at a frequency of 30 vibrations per second). Bead mill racks must be pre-chilled in liquid nitrogen before use.
Quickly add 150 μL of extraction buffer (see Recipe 1).
Vortex for 15 s.
Centrifuge at 12,000× g for 10 min at 4 °C.
Transfer at least 120 μL of supernatant into a well of a PCR plate and keep on ice. Avoid pipetting debris since it may interfere with the measurement in the plate reader. This supernatant is hereafter called the crude extract.
Measure initial NADP-MDH activity from crude extracts. We suggest measuring it at least three times for each sample (see Section C, step 4a.i.). Since the activity slows down rapidly after extraction due to spontaneous oxidation of the extract, these measurements should be performed as soon as possible.
NADP-MDH activation:
For each sample, prepare 12 μL of activation medium (see Recipe 2) and transfer this medium into a well of a PCR plate.
Using the multichannel pipette, transfer 50 μL of crude extract into the PCR plate wells containing the activation medium. Mix by pipetting up and down three or four times. Avoid forming bubbles. If bubbles form, a short spin of the PCR plate can help to remove them.
Incubate the plate at 21 °C for 20 min in a thermal cycler.
Measure maximal NADP-MDH activity (see Section C, step 4a.ii.).
Protein quantification:
Dilute crude extract at 1/20 in Tris pH 6.8 buffer.
Use 2 μL of diluted sample to assay the protein concentration with the Qubit Protein and Protein Broad Range (BR) Assay kit following the manufacturer’s protocol.
Activation of recombinant NADP-MDH, in vitro assay (alternative for A)
Prepare M1 and M2 medium (see Note 3 and Recipe 3)
Prepare 15 μL of M1 medium per activation kinetics and transfer into a PCR plate.
Prepare 15 μL of M2 medium per activation kinetics and transfer into another row of the same PCR plate.
Incubate the plate at 21 °C for 5 min in a thermal cycler.
Activation kinetics:
Using a multichannel pipette, mix the 15 μL of M2 medium with the 15 μL of M1 medium. Mix by pipetting up and down three or four times, avoiding forming bubbles, and start the timer.
Immediately measure the t0 NADP-MDH activity (see Section C, step 4b).
If bubbles have formed at the previous step, a short spin down can help to remove them; in any case, be careful not to pipette them at further time points.
Keep incubating at 21 °C and let the thermal cycler with the lid open.
At each time point of the activation kinetics, measure the NADP-MDH activity (see Section C, step 4b). We suggest measuring activity at 0, 3, 6, 9, 12, 15, 20, and 25 min of activation.
NADP-MDH activity monitoring
Create and save the program for the Tecan plate reader (see Note 4).
Prepare reaction medium (see Recipe 4) and transfer 200 μL of this medium into each well of a Genetix plate (see Recipe 4).
Prewarm at 30 °C the plate containing the reaction medium in the Tecan.
Monitor the NADP-MDH activity (see Note 5).
For ex planta assay:
i. Measure initial NADP-MDH activity, as soon as possible after extraction. For this purpose, add 20 μL of crude extract to 200 μL of prewarmed reaction medium and mix as recommended in Note 4. Start OD340nm monitoring for 1 min. We recommend performing three measurements for each sample.
ii. After 20 min activation at 21 °C, measure maximal NADP-MDH activity. Add 20 μL of activated extract to the 200 μL of prewarmed reaction medium and mix as recommended in Note 4. Start OD340nm monitoring for 1 min. We recommend performing three measurements for each sample.
For in vitro assay: at each time point of the activation kinetics (0, 3, 6, 9, 12, 15, 20, and 25 min), withdraw 3 μL of activation mixture, add them to the 200 μL of prewarmed reaction medium, and mix as recommended in Note 4. Start monitoring OD340nm for 1 min.
Export raw data to an Excel sheet through Magellan software interface.
Data analysis
In this protocol, NADP-MDH activity is defined as the estimated slope calculated by a linear model on the four first data points. We use R to estimate slopes and automatize the analysis, since the procedures can quickly generate a lot of data. The R script was developed to be user friendly. The user does not need to be skilled in programming. We propose a criterion to filter reliable slopes from problematic ones. Further data processing, such as replicate averaging, plotting and statistical analysis, can be conducted in R or Excel. This part, being out of the scope of the present paper, is not detailed here.
For NADP-MDH ex planta activity, follow sections A, B, and C. For NADP-MDH in vitro assay, follow sections A, B, and D.
Raw data formatting
Open exported raw data with Excel. All the OD values obtained are on the same sheet. An example of an Excel sheet containing raw results is provided as supplemental data and its layout is commented in Figure 2.
Figure 2. Example of raw data exported in Excel. Red, green and blue boxes show OD values measured in Genetix plate row A, B and C respectively. Red, green and blue dashed boxes show time stamps for Genetix plate row A, B and C respectively.
Copy the values obtained from all measured rows of the Genetix plate into as many separated tabulated text files. To do this, you need to copy time stamps and monitored values for each plate row from the Excel sheet to a new tabulated text file (extension .txt). In these files, the first column must be time stamps in seconds, then OD values for columns 1–12 (or fewer). The first row is the header containing column names (could be numbers or specific names). During the 1 min monitoring, the plate reader can acquire 9–11 OD measurements, thus the text file should have 10–12 rows (including the column/sample names). An example of text file is provided in Table 1; it corresponds to the values obtained for a single plate row (row C in the provided dataset). At the end of this step, you should have as many text files as measured rows.
Table 1. Example of text file containing exported raw data. In this dataset, the plate reader measured nine points during 1 min acquisition; the first column contains time stamps in seconds and the first row contains column names. Here, samples are named from 1 to 12.
Time 1 2 3 4 5 6 7 8 9 10 11 12
0 0.8235 0.9455 0.8758 0.859 0.8209 0.9507 0.8806 0.8498 0.8001 0.9164 0.8633 0.8185
7 0.744 0.9145 0.8366 0.7988 0.7369 0.9195 0.8346 0.7875 0.7259 0.8869 0.8178 0.7561
14 0.6787 0.8889 0.7946 0.7416 0.6732 0.8887 0.7906 0.7295 0.6594 0.8591 0.7721 0.6968
21 0.6211 0.8621 0.7506 0.6868 0.6134 0.8572 0.7483 0.6764 0.5998 0.832 0.7326 0.6444
28 0.5724 0.8359 0.7151 0.6373 0.5668 0.8313 0.7098 0.6259 0.5452 0.802 0.695 0.595
35 0.5292 0.8104 0.6767 0.5939 0.5206 0.8033 0.6723 0.5815 0.5004 0.7775 0.6594 0.553
42 0.4909 0.7865 0.6443 0.5543 0.4819 0.7779 0.6396 0.5437 0.4634 0.7511 0.6253 0.5146
50 0.4617 0.7618 0.6134 0.5186 0.4513 0.7564 0.608 0.5083 0.4364 0.7291 0.5959 0.4787
57 0.4321 0.7396 0.5858 0.4884 0.4234 0.7317 0.5783 0.4766 0.409 0.7067 0.569 0.4507
Slope estimation on R
Open RStudio.
Create and save the script to analyze data in order to estimate the NADP-MDH activity defined as the initial slope and some other values (see Note 6).
Analyze data for the first text file:
Adapt line 3 in the script. You must type the name of text files between the quotation marks. As an example, if the text file name is rowC.txt, the line 3 should be: data_name="rowC.txt" (see Note 6).
Execute all the lines of the script. The script generates two outputs: a pdf file with curves of each monitored NADP-MDH activity, and a text file containing estimated slopes.
Open the pdf file and check every single curve. The curve should be bended or linear and continuously decreasing, without abrupt breaks. Example of expected curves and problematic curves are presented in Figure 3. Most unexpected shapes could be due to homogenization and/or bubble issues. It could be frequent at the beginning, but as soon as the gesture is mastered, it becomes rare.
Figure 3. Example of NADP-MDH activity curves. Evolution of OD340nm over time showing the NADPH consumption by MDH. Blue line is the linear model calculated on the basis of the 4 first time points. Value printed in the top right corner is the coefficient of variation of the estimated slope. A. Example of expected curves. B. Example of problematic curves. Curves 1 and 2 have a first value lower than the second (indicated by circles). Curves 3, 5 and 6 have abrupt breaks (indicated by arrowheads).
Check whether the calculated linear model correctly fits the curve or not. In addition to the curves’ quick check, we suggest using a criterion based on the slope coefficient of variation. We suggest discarding the slope value when this coefficient is higher than 15% (see Note 7).
Open the output text file. This file contains different values recorded for different curves: the estimated slope, its associated standard error calculated by the linear model, the coefficient of variation, and the calculated r of the linear model. An example of this output is presented in Table 2.
Table 2. Example of output text file. Extract of the output text file obtained with the dataset in Table 1. Slope: value of the estimated slope; coeffvar: value of the coefficient of variation; sd: calculated standard error of the slope value; rsquared: value of calculated linear model r.
rowC rowC_1 rowC_2 rowC_3 rowC_4
slope -0.00960714285714286 -0.00393999999999999 -0.00596571428571429 -0.00819714285714285
coeffvar 5.19414804028292000 2.94294762693293000 1.82244229793482000 1.48983689600294000
sd 0.00049900922244147 0.00011595213650116 0.00010872170051680 0.00012212405870378
rsquared 0.99463312400325000 0.99827080714040100 0.99933618176150200 0.99955627418466600
Proceed similarly for all text files.
Activity analysis for ex planta assay (alternative for D)
Calculate average initial activity and average maximal activity of the three measurements for each sample.
Calculate initial activity and maximal activity per microgram of protein.
Calculate activation ratio defined as the ratio of initial activity with maximal activity.
Perform statistical analysis using experimental replicates (see Note 8).
Activity analysis for in vitro analysis (alternative for C)
Plot activation kinetics with the calculated average activity.
Perform statistical analysis of experimental replicates (see Note 8).
Validation of protocol
This protocol was used to generate the data published by Baudry et al. (2022). The protocol reliability and reproducibility are illustrated in Figure 4 by a dataset row C, corresponding to raw data of NADP- MDH assays for four different samples of recombinant MDH performed in triplicate.
Figure 4. Example of replicate variation. NADPH consumption curves and deduced slope values obtained from three replicates of four different conditions, using recombinant MDH. A. NADPH consumption curves; symbols represent condition average values at each kinetics time point. B. Estimated slope absolute values of curves presented in A, symbols represent condition average values.
Notes
Protein quantitation can be performed using any other protein assay with a suitable range of sensitivity and using a standard spectrophotometer.
Leaf sampling: since NADP-MDH activation state undergoes fast variations with light intensity and along the light photoperiod, all leaf samples must be collected at the same time and in situ of plant culture. We recommend rapidly collecting leaves well exposed to the light and at the middle of the photoperiod. Leaf samples can be stored for several weeks at -80 °C after snap-freezing in liquid nitrogen.
M1 and M2 activation medium: recipes provided for M1 and M2 medium are standard conditions for full activation of NADP-MDH in vitro after 20 min incubation. This must be validated by checking that maximal activity has been obtained (activation plateau).
Tecan programming: to prewarm reaction medium and follow OD340nm for 1 min row by row, the Tecan must be programmed with its dedicated software Magellan. The programming is simple and consists in adding action boxes. The following program can be used for both types of assays. The user request function allows to pause the program until the user clicks on the pop-up, and allows to wait for the incubation times or during user actions:
Box Plate: select the type of plate.
Box Move Plate: check Out.
Box User Request: type a text to inform that user has to place the plate containing the reaction medium and prewarming is going to start.
Box Move Plate: check In.
Box Temperature: check On and set the temperature to 30.0 °C.
Box Wait for temperature: set minimum and maximum temperature to reach (recommendation: 29.5 and 31.0).
Box User Request: type a text to inform user that the temperature is reached.
Box User Request: type a text to inform user that Tecan is ready for measurement.
Box Move Plate: check Out.
Box User Request: type a text to inform user to mix sample in row A for measurement.
Box Move Plate: check In.
Box Part of Plate: select row A.
Box Kinetic Cycle: set Duration on 1 min.
Box Absorbance: set wavelength to 340 nm and number of flashes to 1.
Repeat steps 4h–4n for other plate rows (B to H).
Box Move Plate: check Out.
Recommendation to properly start NADP-MDH activity monitoring: using the multichannel pipette, quickly mix the sample with the assay medium. We strongly recommend mixing by stirring the mixture with the pipette tip making three or four turns. Do not mix by pipetting up and down to avoid making bubbles. Because the plate reader measures the OD vertically, bubbles can disrupt the measurement and make the data unusable. If bubbles form, you can try to quickly push them to the side; however, you run the risk of missing the complete row measurement by spending too much time on it, thus missing the beginning of the reaction. After mixing, as quickly as possible, hit the OK button on the software pop-up to move the plate in the plate reader and start monitoring.
R script to analyze raw data and calculate NADP-MDH activity: the following R script can be used to analyze data (plot curves of experimental data, slope estimation, coefficient of variation calculation, and export all these data as pdf and text files).
Line 3 (data_name="rowA.txt") must be adapted for every file to be analyzed.
Although not recommended, the window parameter (line 16) that defines which points are included in the linear model for the slope calculation can be modified for better fitting of some curves. In that case, the same parameter value should be used for all the samples (see Note 7).
<script>
##Data Import ####
#enter the data text file's name
data_name="rowA.txt"
#output file names creation
data_name.short=strsplit(data_name,split="\\.")[[1]][1]
pdf_name=paste0(data_name.short,".pdf") #output pdf name
outfile_name=paste0("Slope_",data_name.short,".txt") #output txt file name
#import raw data, change the column names and check that time stamp column is numeric
rawdata=read.table(data_name, header=T, sep="\t", check.names=F, comment.char="") colnames(rawdata)=c("time",paste(data_name.short,colnames(rawdata)[-1],sep="_"))
rawdata$time=as.numeric(gsub("s$","",as.character(rawdata$time)))
##Data Analysis ####
#parameters setting
window=c(1:4) #point used to estimate the slope, suggested default value = c(1:4)
MIN=min(rawdata[,-grep("time",colnames(rawdata))],na.rm=T) #lowest OD value, graphical parameter
MAX=max(rawdata[,-grep("time",colnames(rawdata))],na.rm=T) #highest OD value, graphical parameter
#slope estimation and plot
Slope.df=data.frame(matrix(nrow=4,ncol=ncol(rawdata)-1)) #data.frame to save slope estimation results
colnames(Slope.df)=colnames(rawdata[,-1])
row.names(Slope.df)=c("slope","coeffvar","sd","rsquared")
pdf(pdf_name) #export plots as a pdf
par(mfrow=c(3,4)) #12plots per page
for (i in 2:ncol(rawdata)) {
#slope estimation
LMtmp=lm(rawdata[window,i]~rawdata[window,"time"]) #estimation of the slope during the set window with a linear model
LMsum=summary(LMtmp)
Slope.df[c(1,3,4),(i-1)]=c(LMsum$coefficients[2,1:2],LMsum$r.squared) #save estimated slope, slope std error and the lm's r.squared
Slope.df[2,(i-1)]=100*abs(LMsum$coefficients[2,2]/LMsum$coefficients[2,1]) #coefficient of variation = sd/slope
#plot
plot(x=rawdata$time, y=rawdata[,i], ylim=c(MIN,MAX), xlim=c(0,60), pch=20, type="o", ylab="OD_340nm", xlab="time (s)", main=colnames(rawdata)[i]) #raw data plot
abline(LMtmp,col="blue") #plot the estimated slope
legend("topright","",bty="n",title=paste0(round(Slope.df[2,(i-1)],2),"%")) #add coefficient of variation value
#rm tmp data
rm(LMsum,LMtmp)
}
dev.off()
##Data Output ####
final.df=cbind.data.frame(data=rownames(Slope.df),Slope.df) #table to export as a txt file
colnames(final.df)[1]=data_name.short
write.table(final.df,outfile_name,row.names=F,col.names=T,quote=F,sep="\t") #print as a txt file a table containing the recorded slopes and others values
</script>
Slope coefficient of variation used: in this protocol, we define the measured NADP-MDH activity as the initial slope calculated on the basis of acquired data. For reliable results, the linear model used to estimate the slope should properly fit the curve. During the linear modeling, R estimates the intercept and the slope of the model and provides standard error value for both parameters. Our proposed coefficient of variation is the ratio of the slope standard error with the estimated slope value. We suggest discarding the slope value when this coefficient is higher than 15% to ensure reliability of the value. This value has been defined empirically, since most curves with a coefficient value higher than 20% (when the slope is calculated with four points) show an unexpected shape.
In Figure 3B, for five out of six example curves we might want to calculate the slope using only three points instead of four, as suggested. Removing the first time point cannot be considered for curves 1 and 2; since we try to measure the initial velocity, these curves should be discarded from the analysis. Removing the fourth point could be considered as long as all the slopes of the dataset are calculated on three points and the overall curve appearance is good (i.e., without an abrupt break). As an example, one can envisage doing such modeling on three points for curve 4 but not for curves 3 and 5. Curve 6 should be discarded from the dataset since it shows many break points.
According to the experimental design, different tests can be done. For example, for a design with one factor at two modalities (e.g., wild type vs. mutant), t-tests can be done; for one factor with more than two modalities (e.g., wild type, mutant 1, and mutant 2), one-way ANOVA should be done instead of t-test; for a design with more than one factor (e.g., wild type vs. mutant, both in mock and treatment conditions), a two-way (or more) ANOVA should be done. Although these tests cannot be done for kinetics, they can be done on calculated kinetic slopes, activity at a precise time point, etc.
Recipes
Following buffers, A, B, and C must be freshly prepared and kept on ice until used, since their oxidation could alter the results.
Extraction buffer for ex planta assay
100 mM Tris pH 6.8
Protease inhibitor cocktail for plant (diluted 100 times)
Activation medium for ex planta assay
50 mM DTT
50 μM recombinant TRX m type
Activation medium for the in vitro assay
The NADP-MDH activation mixture is as following: 0.24 μg/μL recombinant NADP-MDH; 10 mM DTT; 10 μM recombinant TRX m type; 30 mM Tris pH 7.9. To start the activation reaction, the two media M1 and M2 are mixed (by pipetting up and down three or four times) 1:1 vol:vol.
M1 contains only the NADP-MDH in Tris buffer:
0.48 μg/μL NADP-MDH
30 mM Tris pH 7.9
M2 contains all reagents, except NADP-MDH, in Tris buffer:
20 mM DTT
20 μM recombinant TRX m type
30 mM Tris pH 7.9
Reaction medium
160 μM NADPH
750 μM OAA
30 mM Tris pH 7.9
Prewarm at 30 °C.
This medium must be freshly prepared and cannot be stored more than half a day.
Acknowledgments
K.B. research was supported by a French Ph.D. fellowship from “Ministère de la Recherche et de l’Enseignement Supérieur.” IPS2 benefits from the support of the Labex Saclay Plant Sciences-SPS (ANR-17-EUR-0007). This protocol was used to produce the data published in the following research article: Baudry et al. (2022).
Competing interests
The authors declare no conflict of interest.
Ethics considerations
Any of these experimental procedures involved neither human nor animal subjects.
References
Baudry, K., Barbut, F., Domenichini, S., Guillaumot, D., Thy, M. P., Vanacker, H., Majeran, W., Krieger-Liszkay, A., Issakidis-Bourguet, E., Lurin, C., et al. (2022). Adenylates regulate Arabidopsis plastidial thioredoxin activities through the binding of a CBS domain protein. Plant Physiol. 189(4): 2298–2314.
Collin, V., Issakidis-Bourguet, E., Marchand, C., Hirasawa, M., Lancelin, J. M., Knaff, D. B. and Miginiac-Maslow, M. (2003). The Arabidopsis Plastidial Thioredoxins. J. Biol. Chem. 278(26): 23747–23752.
Heyno, E., Innocenti, G., Lemaire, S. D., Issakidis-Bourguet, E. and Krieger-Liszkay, A. (2014). Putative role of the malate valve enzyme NADP–malate dehydrogenase in H2O2 signalling in Arabidopsis. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369(1640): 20130228.
Issakidis, E., Saarinen, M., Decottignies, P., Jacquot, J., Crétin, C., Gadal, P. and Miginiac-Maslow, M. (1994). Identification and characterization of the second regulatory disulfide bridge of recombinant sorghum leaf NADP-malate dehydrogenase. J. Biol. Chem. 269(5): 3511–3517.
Jacquot, J. P., Issakidis, E., Decottignies, P., Lemaire, M. and Miginiac-Maslow, M. (1995). [25] Analysis and manipulation of target enzymes for thioredoxin control. In: Lester Packer (Ed.). Meth. Enzymol (pp. 240–252). Academic Press.
Keryer, E., Collin, V., Lavergne, D., Lemaire, S. and Issakidis-Bourguet, E. (2004). Characterization of Arabidopsis Mutants for the Variable Subunit of Ferredoxin:thioredoxin Reductase. Photosynth. Res. 79(3): 265–274.
Scheibe, R. (2004). Malate valves to balance cellular energy supply. Physiol. Plant. 120(1): 21–26.
Scheibe, R. and Dietz, K. J. (2012). Reduction-oxidation network for flexible adjustment of cellular metabolism in photoautotrophic cells. Plant Cell Environ. 35(2): 202–216.
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© 2023 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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Caste Transition and Reversion in Harpegnathos saltator Ant Colonies
CO Comzit Opachaloemphan *
FC Francisco Carmona-Aldana *
HY Hua Yan
(*contributed equally to this work)
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4770 Views: 404
Reviewed by: Khyati Hitesh ShahKevin Haight Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Sep 2022
Abstract
Living organisms possess the ability to respond to environmental cues and adapt their behaviors and physiologies for survival. Eusocial insects, such as ants, bees, wasps, and termites, have evolved advanced sociality: living together in colonies where individuals innately develop into reproductive and non-reproductive castes. These castes exhibit remarkably distinct behaviors and physiologies that support their specialized roles in the colony. Among ant species, Harpegnathos saltator females stand out with their highly plastic caste phenotypes that can be easily manipulated in a laboratory environment. In this protocol, we provide detailed instructions on how to generate H. saltator ant colonies, define castes based on behavioral and physiological phenotypes, and experimentally induce caste switches, including the transition from a non-reproductive worker to a reproductive gamergate and vice versa (known as reversion). The unusual features of H. saltator make it a valuable tool to investigate cellular and molecular mechanisms underlying phenotypic plasticity in eusocial organisms.
Key features
• H. saltator is one of few ant species showing remarkable caste plasticity with striking phenotypic changes, being a useful subject for studying behavioral plasticity.
• Caste switches in H. saltator can be easily manipulated in a controlled laboratory environment by controlling the presence of reproductive females in a colony.
• The relatively large size of H. saltator females allows researchers to dissect various tissues of interest and conduct detailed phenotypic analyses.
Keywords: Ant Harpegnathos saltator Worker Gamergate Revertant Caste transition Caste reversion Phenotypic plasticity
Background
Eusocial insects (ants, bees, wasps, and termites) live in colonies in which all individuals have a high genetic relatedness, while only one or few individuals reproduce (Chapman et al., 2000; Trontti et al., 2005; Loope, 2015). The reproductive (queens and also kings in termites) and non-reproductive (workers) castes in a colony display different phenotypic traits including differential behavior, reproduction, metabolism, and lifespan, a phenomenon known as polyphenism (Simpson et al., 2011; Kapheim et al., 2012). Eusocial insects offer a valuable experimental model to compare gene expression between castes and to identify the key genes as causal factors determining differential phenotypes at the organismal level (Colgan et al., 2011; Bonasio et al., 2012).
Harpegnathos saltator (commonly known as the Indian jumping ant) is an ant species originating from India. These ants display an unusual feature: individuals can transition from non-reproductive workers to reproductive pseudo-queens (also called gamergates) (Liebig et al., 2000; Monnin and Peeters, 2008; Sieber et al., 2021a). When the queen is absent from a colony, workers can switch their caste and show queen-like traits: minimal movement, staying inside the nest, active ovaries, and production of eggs. In contrast, some workers show active movement in the nest to take care of the progeny, while others hunt and forage. Furthermore, workers have small ovaries and rarely produce eggs.
A colony with gamergates is as stable as a queen colony (Opachaloemphan et al., 2021; Penick et al., 2021). Additionally, gamergates can be experimentally converted back to workers (to distinguish, they will be referred to as revertants) (Penick et al., 2021). This process is associated with the reversion of phenotypic traits: revertants display worker-like physiology, behavior, and aging (Penick et al., 2021; Yan et al., 2022). With a high-quality genome (Bonasio et al., 2010; Shields et al., 2018), multiple studies have been performed in H. saltator to analyze caste-specific transcriptomes and methylomes at tissue and/or single-cell levels (Bonasio et al., 2012; Sheng et al., 2020; Sieriebriennikov et al., 2021). H. saltator is one of the three species (along with Oocerea biroi and Solenopsis invicta) in which reverse genetics was applied in ants using the CRISPR-Cas9 system (Trible et al., 2017; Yan et al., 2017; Chiu et al., 2020). This experimental paradigm has helped to identify cellular and molecular processes required for the establishment of caste identity (Gospocic et al., 2021; Opachaloemphan et al., 2021) and determine the role of neuropeptides, hormones, and transcription factors in regulating caste-specific behaviors, and the role of insulin signaling in reproduction and longevity (Penick et al., 2014; Gospocic et al., 2017 and 2021; Yan et al., 2022). Here, we provide a detailed protocol explaining the experimental framework we have used on H. saltator to induce caste transition and reversion, the identifiable and quantifiable parameters to define each caste, and the techniques for tissue dissection and staining.
Materials and reagents
Oil-based paint markers (Uni-Paint, catalog number: 63721); store at room temperature
Entomology featherweight forceps (BioQuip, Featherweight Forceps, catalog number: 4748)
Dissection forceps (Dumont #55, catalog number: 11295-51)
Wood flour (System Three Resins, System Three); store at room temperature and dry
Dental plaster (Kulzer, Modern Materials Lab Stone Blue Type III); store at room temperature
Small plastic container (Pioneer Plastics, catalog number: 028C, inner dimensions: 9.52 cm × 9.52 cm × 7.77 cm)
Medium plastic container (Pioneer Plastics, catalog number: 079C, inner dimensions: 18.89 cm × 13.49 cm × 9.53 cm)
Crickets (Ghann’s Crickets, Live Crickets " and ¼", catalog number: C14 and C38, respectively)
Cricket chow (Ghann’s Crickets, Ghann’s Cricket chow, catalog number: FD-GCC); store at room temperature. Mix with apples and potatoes for feeding crickets
Cover glass for the chamber (dimensions: 10.0 cm × 9.0 cm × 0.5 cm)
Fluon (Fluorogistx, TeflonTM PTFE DISP 30 Fluoropolymer Dispersion, catalog number: D14783090); store at room temperature
2" Poly foam brush (Jen Manufacturing, catalog number: JNSN846)
Multipurpose foam mats [American Floor Mats, Soft Floors Interlocking Tiles, dimensions: 9.0 cm × 7.5 cm × 1.0 cm with an additional area (bulge) of 2.3 cm × 1.5 cm × 1.0 cm for the nest entrance]
16% paraformaldehyde (PFA) (Electron Microscopy Sciences, catalog number: 15710); store at room temperature
Triton X-100 (Sigma, catalog number: X100); store at room temperature
Alexa FluorTM 488 Phalloidin (Thermo Fisher, Alexa FluorTM, catalog number: A12379); store at 4 °C and protected from light
DAPI dye (Tocris, catalog number: 5748); store at -20 °C
Aluminum foil (Reynolds Wrap)
Single sided iSpacer 0.5 mm deep (SubJin Lab Optical Clearing Innovation, iSpacer, catalog number: IS008)
Microscope slides (Fisher Scientific, catalog number: 12-550-143)
Microscope cover glass (Fisher Scientific, catalog number: 12-542A)
Mounting solution (Thermo Fisher, Slow-FadeTM Gold Antifade mountant, catalog number: S36936); store at room temperature and protected from light
Black dissection Petri dish (Pyrex, catalog number: DD-90-S-BLK)
Sodium chloride (NaCl) (Fisher Scientific, catalog number: S271); store at room temperature
Potassium chloride (KCl) (Fisher Scientific, catalog number: 217); store at room temperature
Sodium phosphate dibasic (Na2HPO4) (Fisher Scientific, catalog number: S374); store at room temperature
Potassium phosphate monobasic (KH2PO4) (Fisher Scientific, catalog number: P285); store at room temperature
Hydrochloric acid (HCl) (Fisher Scientific, catalog number: A144S); store at room temperature
1× PBS pH 7.4 (store at room temperature, see Recipes)
Equipment
Temperature-controlled incubator (Percival Scientific, catalog number: I-36VL)
Tube rotator (Thermo Scientific, Tube Revolver, catalog number: 11-676-341)
Software
Fiji (ImageJ, https://imagej.net/software/fiji/) (Schindelin et al., 2012)
Procedure
Nest preparation
Prepare a mix of 1,200 mL of deionized H2O and 1,900 g of dental plaster powder. Add the powder gradually while stirring. Dissolve any clumps and avoid forming bubbles. Add a small quantity of plaster powder or water to achieve the best consistency. This quantity is enough to prepare four medium-sized (Figure 1A) or eight small containers.
Pour the mix into the containers, forming a layer 2.5 cm high for medium containers (Figure 1B) or 1.5 cm for small containers. Remove any air bubbles after pouring by tapping the bottoms of the containers against a solid surface (Sieber et al., 2021b). This needs to be done right after pouring the mix, as it does not take long to solidify.
While the wet plaster is drying, place a cover glass on top of a foam mat and place them near the rear center of the nest (Figure 1C). Push down the mat a few centimeters deep into the plaster to form the chamber (only for medium containers, Figure 1D) (Sieber et al., 2021b). The cover glass should not be submerged into the plaster, and the mat should not reach the bottom of the plastic container.
On the next day, remove the cover glass and the mat using a spatula. Use the spatula to remove extra plaster and make uniform borders around the chamber. Leave the nest to dry for one week.
Once the nest is dry, prepare a solution with equal parts Fluon and H2O together. Mix by gently swirling to avoid bubbles. Hold the nest upside down and use a foam brush to apply a continuous layer throughout all the walls of the container, starting 1 inch above the plaster surface (Figure 1D). Avoid bubbles and contact between Fluon and the plaster. Leave the nest to dry by placing it upside down.
After Fluon dries, place the cover glass over the chamber (Figure 1B and 1D). Place the lid on the container. The nest is now ready to use. Alternatively, store the nest at room temperature.
Note: Use the same procedure for small containers. Skip step A3 and A4.
Figure 1. Nest preparation. (A) Plastic containers are used to make ant nests. (B) Dental plaster is used to make the nest in the medium plastic container forming a 2.5 cm thick layer with a deep area for the chamber in the center. (C) A foam mat (see Materials and Reagents) is used during nest preparation to cast the chamber in the plaster. (D) A cover glass is used to cover the chamber. A layer of Fluon is applied to the walls of the container, which prevents the ants from crawling out.
Caste transition (from workers to gamergates) and caste reversion (from gamergates to revertants)
Generate gamergates: set up a transition colony containing 30 age-matched workers (approximately 2–4 weeks post eclosion) reared in a temperature-controlled incubator (approximately 25 °C).
Collect 30 young workers (a.k.a. callows) that can be distinguished by their yellow cuticle in contrast to the brown cuticle of older workers (Figure 2A). They should be collected from a healthy and mature colony, which contains eggs, larvae, pupae, at least 100 adults, and, importantly, contains no dueling or policing events.
Figure 2. Paint marks to identify ants. (A) Recently eclosed ants, also called callows (1), are identified by their yellow cuticle, as opposed to mature ants with dark brown cuticle (2). Oil-based paint markers are used to label each ant with a unique combination of colors (3). (B) Marks are painted while holding the ants with featherweight forceps. (C) The transition colony is set with 30 workers, each marked with a unique combination of colors.
Label each callow with oil-based paint markers. Gently hold the ant by the thorax using featherweight forceps and paint each ant with a dot of different color on its thorax. Some ants can be painted with two or three dots, so the combination of colors can be used to distinguish individuals (Figure 2B).
Note: Handle callows gently, as they have a very soft cuticle that can be easily damaged.
Keep the labeled callows in an empty box for at least 5 min for the paint to dry before returning them back to their original colony. Keep them in that colony for one week.
Prepare a new medium-sized nest box by adding deionized water to wet the plaster, as mineral ions in water can affect the plaster’s absorption ability. Watering the plaster is necessary to maintain humidity in the colony. Wait for water to be absorbed. Transfer the labeled callows to the new nest (Figure 2C).
Behavioral observations during the transition from worker to gamergate: to distinguish the prospective gamergates from workers, we monitor for gamergate-specific behaviors, such as antennal dueling, egg-laying, and dominance (see below). On the other hand, workers perform foraging, cleaning, and colony defense.
Feed with five pre-stung crickets (previously stung crickets by workers of another mature colony) per transition colony every two days or three times per week.
While feeding, remove rotten crickets and any colony waste.
Observe the frequency of antennal dueling for at least 15 min each time, three times per week (Figure 3A and Video 1). The first observation of dueling typically occurs within the first 3–7 days after the colony is set up. This is counted as day 1 of the worker-to-gamergate transition.
Note: Intensive and consistent dueling behavior for at least one week is an early indication of the prospective gamergate. Use their painted color code to track gamergate- and worker-specific behaviors of all individuals in the colony. Individuals that show foraging behavior (stinging crickets and moving them into the chamber) and no dueling are workers.
Figure 3. Differential phenotypes of prospective gamergates and workers during transition. (A) Constant duelers actively interact between themselves, forming a gamergate circle (dashed line). (B) Egg laying can be identified by ants bending their gaster towards their head, staying in this position for several minutes. (C) A worker approaches a gamergate (blue mark) with its body laid down close to the ground in a submissive position (single arrowhead). Compare this with the stance of another worker (double arrowhead) that is not showing a submissive position. (D) Gamergates are usually found on top of the pupae (single arrowhead) and are less active than workers (double arrowhead). (E) Workers can be found either inside (*) or outside (x) the chamber, while gamergates are exclusively found inside the chamber.
Video 1. Antennal dueling during the caste transition from workers (W) to gamergates (G). Note that the dueling occurred between five ants near the center of the chamber, marked with yellow-green, green-green, yellow-yellow, white-green-pink, and pink-green-pink.
Observe egg-laying events in the dueling ants: dueling ants begin laying eggs approximately one week after initiating the transition (Figure 3B). Constant egg-laying during the first three months of caste transition is another indication of mature gamergates. The prospective gamergate also displays dominant behavior: standing tall when confronting a non-reproductive worker, while the submissive worker keeps its head low (Figure 3C). In addition, gamergates are frequently found on the top of the brood pile and located inside the nest (Figure 3D and 3E). In a colony of 30 ants, 3–6 gamergates are expected to be identified.
The worker-to-gamergate transition normally takes three months.
Caste reversion (from gamergates to revertants): this step is to revert mature gamergates obtained from the previous caste transition back to a non-reproductive status (Figures 4 and 5).
Figure 4. Caste reversion. (A) A mature and stable colony is used as a host colony (see diagram in Figure 5). (B) Gamergates are isolated individually in a small nest box prior to policing. (C) Policing is usually observed against isolated gamergates when they are transferred into the host colony. (D) Revertants show worker phenotypes as hunting and stinging crickets. (E) Revertants display an aggressive response when being provoked by featherweight forceps.
Figure 5. Diagram of caste reversion from gamergates (G) to revertants (R). Four gamergates and four workers (W) are collected from the caste transition colony. Four workers are transferred to the host colony. The gamergates are first individually isolated in a small nest box for four weeks, after which they are transferred to the host colony. Light-gray colored ants indicate local gamergates and workers from the host colony, while dark-colored ants represent gamergates and workers transferred for reversion.
From the transition colony, choose four mature gamergates with the most frequent dueling and egg laying events. These gamergates are subject to individual isolation followed by their reversion in another healthy and stable colony (referred now as a host colony in Figure 4A). Choose four workers with the least dueling and no egg-laying events and directly transfer them into the host colony (Figure 5).
Behavioral observations:
Prepare small nest boxes similar to the method of making medium-sized nests, but do not cast a chamber. Add some deionized water to provide humidity to the box. Transfer the gamergates selected in step B3a out of the transition colony and isolate each gamergate in a small nest box (Figure 4B). Add 2–3 small pre-stung crickets and a pinch of wood flour (sawdust), which is used by workers for cleaning and brood rearing tasks. This individual isolation reduces the fertility status due to the lack of social interaction (Penick et al., 2021). During the first week of isolation, gamergates still lay eggs, which helps to confirm their gamergate status. Keep the gamergates isolated for four weeks. The egg-laying events will slowly fade out. Feed them by adding new crickets three times per week.
While step B3b starts, transfer four workers selected in step B3a to the host colony, where these four workers will maintain their worker status.
After four weeks of isolation, transfer each gamergate to the host colony (the same host colony that is used in step B3c). Policing is exerted by workers from the host colony towards the recently transferred gamergates (Liebig et al., 1999; Penick et al., 2021) (Figure 4C). On the contrary, policing events rarely occur towards the workers transferred in step B3c.
After two months, test the behavior of transferred gamergates (now revertants, see below). The revertants stop laying eggs, show foraging behavior (Figure 4D), and spend more time outside the chamber as workers (Figure 3E).
Testing revertants for worker-like behaviors:
Cricket stinging test: place one revertant with one live cricket in an empty nest and test for hunting behavior. When the revertant hunts, it grabs the cricket with its mandibles and stings the cricket with its stinger (Figure 4D). After being stung, the cricket gets paralyzed.
Forceps assay: place a revertant in an empty nest and leave it there for 10 min to acclimate. Then, use featherweight forceps to provoke the revertant by rapidly snapping the forceps in front of their antennae. The revertant aggressively bites or grabs the forceps with its mandible (Figure 4E).
Dissection and staining
Ovary tissue dissection:
Cool 1× PBS by placing it on ice. Place the ant to be dissected in a 1.5 mL tube on ice for a few minutes to let it fall asleep.
Make a cut with scissors in the narrow region (petiole) that connects the gaster with the thorax.
Take the gaster to the black dissection Petri dish. Place it in a drop of ice-cold 1× PBS. For workers, use dissection forceps to pull the stinger out with a single, quick movement. This will take out the gut and other attached tissues, ovaries included. Avoid using this technique for gamergates, as it causes the oocytes to burst. Instead, open the gaster with dissection forceps by pulling apart between the second and third abdominal segments (tergites). Do not insert the tip of the forceps deep into the gaster to avoid damaging the ovaries.
Find the ovaries close to the stinger, ventral to the gut. They look like two structures bound together at the base (Figure 6A and 6B). If it is a gamergate, oocytes are evident. Be careful in this case, as large oocytes can be easily burst (Figure 6B).
Use the dissection forceps to separate the ovaries from the rest of the tissues. Do not break the gut, as the midgut and the rectum will release their contents if burst and will damage surrounding tissues. Fat body cells are usually found attached to the ovaries and should be removed.
Move the isolated ovaries to a separate drop of 1× PBS. The worker ovaries are formed by two groups of four ovarioles and look transparent (Figure 6A). The gamergate ovaries contain egg chambers in different stages of development and can be distinguished by their white yolk and oval shape (Figure 6B).
Figure 6. Ovary dissection and immunofluorescence staining. (A) Brightfield image of worker ovaries, with four transparent ovarioles on each side. Scale bar, 100 μm. (B) Brightfield image of gamergate ovaries showing egg chambers in different developmental stages. The most mature oocyte has an oval shape and contains a dense white yolk. Scale bar, 200 μm. (C) Confocal image of a gamergate ovariole stained with Phalloidin (green) and DAPI (blue). Scale bar, 500 μm. (D) A yellow line is drawn in Fiji (see Data Analysis) around the borders of an egg chamber to measure its area (magnified from C).
Immunofluorescent staining:
Fix the dissected ovaries by adding 1 mL of freshly prepared 4% PFA in 1× PBS in a 1.5 mL tube.
Rotate the fixing tissues for 30 min at room temperature or overnight at 4 °C on a tube rotator.
Remove the fixation solution and then wash the tissue with 1 mL of 0.1% Triton X-100 in 1× PBS (PBSTx). Incubate on a rotator for 15 min.
Repeat step C2c three times.
Remove PBSTx and add 1 mL of PBSTx containing 2 μg/mL DAPI and 1:100 Alexa FluorTM 488 Phalloidin. DAPI and Phalloidin staining detect the nuclei and cell shape, respectively.
Protect the tube from light by covering it with aluminum foil.
Rotate the tube overnight at 4 °C on a rotator.
Remove the solution and replace it with 1 mL of PBSTx.
Rotate the tube for 5 min.
Repeat step C2h and C2i five times.
Cut a pipette tip with scissors to leave a wide-opening tip.
Prepare a glass slide by attaching a 0.5 mm iSpacer on the slide. This prevents the tissue from being smashed after placing the glass cover.
Transfer the tissue to the iSpacer slide from the previous step. Use the wide-opening pipette tip prepared in step C2k to minimize any damage to the tissue.
Remove any excess solution from the iSpacer using a pipette.
Add 20 μL of the Slow-FadeTM mounting solution.
Place the glass cover and seal the edges with clear nail polish.
Air-dry the nail polish for approximately 5 min. Protect it from light.
Keep the slide at 4 °C in the slide tray. Take images with a confocal microscope.
Data analysis
Image analysis
To measure the area of the mature egg chambers, use the Fiji software (Schindelin et al., 2012).
The ovary is a relatively thick tissue. Z-stack confocal images are normally taken to inspect the whole tissue.
Choose the section through the Z-stack that contains the largest surface area of the egg chamber to be measured.
Select Analyze from the menu bar and then Set Scale. A window will pop up, displaying the Scale. Adjust accordingly.
Use the Polygon tool under the menu bar to draw a line that surrounds the egg chamber to be measured as shown in Figure 6C and 6D. An egg chamber is formed by the nurse cells and the oocyte.
Note: For a more accurate drawing, use the Magnifying glass tool under the menu bar to zoom in on the image.
Select Analyze/Measure from the menu bar. The measured surface area appears in a pop-up window.
Recipes
1× PBS pH 7.4
Reagent Final concentration
NaCl 137 mM
KCl 2.7 mM
Na2HPO4 4.3 mM
KH2PO4 1.4 mM
Adjust the pH to 7.4 with HCl
H2O add up to 1,000 mL
Acknowledgments
This protocol was derived from the research published in Yan et al. (2022). This work was supported by HHMI Collaborative Innovation Award (CIA) #2009005 and HCIA #2009005, NIH grants R21GM114457, R01EY13010, R01AG058762, R01DC020203, NIH Ruth L. Kirschstein NRSA postdoctoral fellowship F32AG044971 and NSF I/UCRC CAMTech grant IIP1821914. We thank Jakub Mlejnek for technical support, and Kayli Sieber for help with manuscript preparation.
Competing interests
The authors have no competing interests to declare.
References
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4,771 | https://bio-protocol.org/en/bpdetail?id=4771&type=0 | # Bio-Protocol Content
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Visualizing NBD-lipid Uptake in Mammalian Cells by Confocal Microscopy
JB Julia F. Baum
LB Lasse Bredegaard
SH Sara Abad Herrera
TP Thomas Günther Pomorski
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4771 Views: 683
Reviewed by: Jan HuebingerShalini Low-NamMario Ruiz
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Original Research Article:
The authors used this protocol in Journal of Cell Science May 2022
Abstract
Eukaryotic cells use a series of membrane transporters to control the movement of lipids across their plasma membrane. Several tools and techniques have been developed to analyze the activity of these transporters in the plasma membrane of mammalian cells. Among them, assays based on fluorescence microscopy in combination with fluorescent lipid probes are particularly suitable, allowing visualization of lipid internalization in living cells. Here, we provide a step-by-step protocol for mammalian cell culture, lipid probe preparation, cell labeling, and confocal imaging to monitor lipid internalization by lipid flippases at the plasma membrane based on lipid probes carrying a fluorophore at a short-chain fatty acid. The protocol allows studying a wide range of mammalian cell lines, to test the impact of gene knockouts on lipid internalization at the plasma membrane and changes in lipid uptake during cell differentiation.
Key features
•Visualization and quantification of lipid internalization by lipid flippases at the plasma membrane based on confocal microscopy.
•Assay is performed on living adherent mammalian cells in culture.
•The protocol can be easily modified to a wide variety of mammalian cell lines.
Graphical overview
Analysis of NBD-lipid uptake in adherent mammalian cells by confocal microscopy. Scale bar, 30 μm.
Keywords: Confocal microscopy Lipid transport Mammalian cells NBD-lipid Plasma membrane
Background
Eukaryotic cells use a series of membrane transporters to control the movement of lipids across their plasma membrane. These transporters can be divided into ATP-dependent flippases and floppases—which catalyze the inward movement of phospholipids from the extracellular/luminal leaflet to the cytoplasmic leaflet and the outward movement of lipids, respectively—and ATP-independent scramblases (Holthuis and Levine, 2005; Contreras et al., 2010). Several tools and techniques have been developed to analyze the activity level of these transporters in the plasma membrane of mammalian cells. Among them, assays based on fluorescent lipid probes are particularly suitable, allowing visualization and quantification of lipid internalization by flippases in living cells. Lipid probes bearing a fluorophore such as nitrobenzoxadiazole (NBD) on a short-chain fatty acid at the sn-2 position, together with a long fatty acid residue at the sn-1 position, are most commonly used (Martin and Pagano, 1987; Koval and Pagano, 1991; Hoekstra and Kok, 1992; Rosenwald and Pagano, 1993) (Figure 1). Both BODIPY- and pyrene-labeled lipids have been used to track lipid trafficking in cells (Pagano et al., 1999; Somerharju, 2002), each having advantages and disadvantages (Table 1). Since the lipid polar head group stays unmodified, recognition as a substrate by ATP-dependent flippases and floppases is not affected (Theorin et al., 2019). At the same time, these molecules are less hydrophobic than their naturally occurring counterparts and readily insert into cell membranes when added to the medium. Transport of these short-chain lipid probes is usually monitored by extraction with bovine serum albumin (BSA) of the residual fraction of analogs not transported across the plasma membrane. As BSA extracts all analogs from the exoplasmic monolayer of the plasma membrane, the inaccessible fraction reflects analogs that have been internalized into cells. Alternatively, NBD-labeled lipids on the outer monolayer can be selectively destroyed with the water-soluble quencher dithionite (McIntyre and Sleight, 1991). However, because dithionite can leak through the membrane in some cells, conditions must be carefully adapted to the particular cell type (Pomorski et al., 1994). Furthermore, NBD-lipids are known to be actively metabolized by phospholipase activities. One frequent modification is their hydrolysis into lyso-derivates by phospholipase A2 activities, which results in the removal of the fatty acid attached to the sn-2 position. The liberated labeled C6 fatty acids are released into the medium, hampering the quantitative analysis of NBD-lipid internalization. Thus, the assay is typically performed in the presence of phospholipase inhibitors (Pomorski et al., 1996; Grifell-Junyent et al., 2022; Herrera et al., 2022).
The protocol presented here utilizes fluorescence microscopy to study NBD-lipid internalization in mammalian cells, exemplified on mouse skeletal muscle cell line (C2C12) cells (Grifell-Junyent et al., 2022), and has also been applied by us to fibroblasts (Pomorski et al., 1996). While NBD-lipids offer advantages (Table 1), including the ability to monitor the activity of lipid flippases at the plasma membrane of cells, it is important to note that these probes are less hydrophobic than naturally occurring lipids. This difference in hydrophobicity can affect the way they are transported within cells and may lead to differences in their intracellular trafficking compared to endogenous lipids. The protocol includes cell preparation, preparation of NBD-lipids, labeling of cells with NBD-lipids, fluorescence microscopy, and data analysis. The protocol can be easily adapted to parasites such as Toxoplasma and Leishmania (Weingärtner et al., 2011; Chen et al., 2021). It can also be applied to study: (i) the lipid uptake profile in mammalian cell lines; (ii) the impact of gene deletions and/or single mutations in lipid uptake at the plasma membrane; and (iii) the possible changes in lipid uptake during cell differentiation. For lipid uptake assays optimized for plants or based on flow cytometry, the reader is referred to previously published protocols (Jensen et al., 2016; López-Marqués and Günther Pomorski, 2021; Herrera et al., 2022).
Table 1. Main advantages and disadvantages of common fluorescently tail-labeled lipid probes
Lipid probe Advantages Disadvantages Reference
NBD-lipids
• Commercially available
• Dithionite quenchable
• Low photostability
• High environmental sensitivity
Martin and Pagano, 1987; Kobayashi and Arakawa, 1991; Pomorski et al., 1996)
BODIPY and TopFluor lipids
• High photostability
• Low environmental sensitivity
• Superior brightness
• MαCD-mediated lipid exchange
• Availability restricted
• Poorly extractable by BSA
(Pagano et al., 1999; Kay et al., 2012; Mioka et al., 2018; Segawa et al., 2021)
Pyrene lipids
• Mimic natural lipids
• Unique spectral features
• UV excitation
• Low photostability
• Availability restricted
• Poorly extractable by BSA
• High environmental sensitivity
Tanhuanpää et al., 2000; Somerharju, 2002
Abbreviation: MαCD, methyl-α-cyclodextrin
Figure 1. Chemical structures of fluorescent lipids used for lipid uptake assays. Glycerophospholipids are composed of two hydrophobic fatty acids and a hydrophilic head group, both combined with a glycerol backbone. Sphingolipids are composed of one hydrophobic fatty acid and a hydrophilic head group, both combined via a sphingosine backbone. The structures of the glycerophospholipid head groups corresponding to phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylserine (PS), as well as for sphingomyelin (SM) are depicted. These fluorescently labeled lipids carry a nitrobenzoxadiazole (NBD) group at the sixth carbon in the short-chain fatty acid at the sn-2 position.
Materials and reagents
Mammalian cell culture
In this study, we used mouse myoblast cells (C2C12; cell number: ACC 565, DSMZ Braunschweig, Germany) that were cultured in growth medium (see Recipes). Optimal culture media and conditions may differ for other cell lines.
Dulbecco's Modified Eagle Medium, high glucose, without pyruvate (high-glucose DMEM) (e.g., Sigma-Aldrich, catalog number: D5796), store at 4 °C
Dulbecco's Modified Eagle Medium, low glucose, without pyruvate (low-glucose DMEM) (e.g., Sigma-Aldrich, catalog number: D6046), store at 4 °C
Ethanol absolute ≥ 99.8% (VWR, catalog number: 20821.321)
Fetal bovine serum, heat inactivated before use (FBS) (e.g., Capricorn Scientific, catalog number: FBS-11A), store at -20 °C
Hanks’ balanced salt solution, Ca2+ and Mg2+ free (HBSS) (e.g., Sigma-Aldrich, catalog number: H6648), store at 4 °C
Horse serum (HS) (e.g., Sigma-Aldrich, catalog number: H1138), store at -20 °C
35 mm polymer bottom dishes (e.g., Ibidi, catalog number: 81156)
1.5 mL microcentrifuge tubes (Sarstedt, catalog number: 72.690.001)
Penicillin-streptomycin, 100× solution (e.g., Sigma-Aldrich, catalog number: P4333), store at -20 °C
Pipette controller (e.g., accu-jet pro, Brand, catalog number: 263 00)
Polypropylene tubes of 15 mL capacity (e.g., Falcon tubes, Sarstedt, catalog numbers: 62.554.502 and 62.547.254)
Sterile serological pipettes (e.g., Serological pipettes of 5, 10, and 25 mL; Sarstedt, catalog numbers: 86.1253.001, 86.1254.001, and 86.1685.001)
Sterile culture vessels T-75 flasks (e.g., Sarstedt, catalog number: 83.3911)
Trypsin-EDTA solution (e.g., Sigma-Aldrich, catalog number: T3924), store at -20 °C
Trypan blue solution, 0.4% (Thermo Fischer Scientific, catalog number: 15250061)
Tyrode’s balanced salt solution (TBSS) (see Recipes), store at 4 °C
Preparation of NBD-lipids
Centrifuge glasses DURAN® with conical bottom, 12 mL (Carl Roth, catalog number: K211.1)
Chloroform 99%–99.4% ethanol-stabilized and certified for absence of phosgene and HCl (VWR, catalog number: 22711.290)
Methanol ≥ 99.8% (VWR, catalog number: 20847)
Glass vials (Rotilabo® screw neck ND8 vials, Brown glass, 1.5 mL, Carl Roth, Karlsruhe, Germany, catalog number: KE30.1) with screw caps (without borehole, without septum, PP, black, ND8, Carl Roth, Karlsruhe, Germany, catalog number: KE39.1) for lipid aliquoting
C16:0-C6:0 NBD-lipids purchased in chloroform including NBD-PC (Avanti Polar Lipids, catalog number: 810130), NBD-PE (Avanti Polar Lipids, catalog number: 810153), NBD-PS (Avanti Polar Lipids, catalog number: 810192), and NBD-SM (Avanti Polar Lipids, catalog number: 810218)
NBD-lipid uptake assay
Bovine serum albumin essentially fatty acid free (BSA) (Sigma-Aldrich, catalog number: A6003), store at 4 °C
Calcium chloride (CaCl2) (Grüssing, catalog number: 10043-52-4)
Dimethyl sulfoxide (DMSO) (Carl Roth, catalog number: 4720.4)
Glucose (Duchefa Biochemie, catalog number: G0802.5000)
4-(2-hydroxyethyl)-1-piperazineethanesulphonic acid (HEPES) (Carl Roth, catalog number: 7365-45-9)
Magnesium chloride hexahydrate (MgCl2·6H2O) (Sigma-Aldrich, catalog number: 7791-18-6)
3-(4-octadecyl)benzoylacrylic acid, 4-(4-octadecylphenyl)-4-oxo-2-Butenoic acid (OBAA) (Sigma-Aldrich, catalog number: SML0075), store at -80 °C
Phenylmethylsulfonyl fluoride (PMSF) (Sigma-Aldrich, catalog number: P7626)
Note: OBAA is a potent inhibitor of phospholipase A2 (Eintracht et al., 1998). PMSF has been reported to inhibit phospholipases as well as some esterases (James, 1978; Gadella and Harrison, 2000; Estévez et al., 2012).
Potassium chloride (KCl) (Merck, catalog number: 7447-40-7)
Sodium chloride (NaCl) (Carl Roth, catalog number: 7647-14-5)
Sodium dihydrogen phosphate (NaH2PO4·2H2O) (VWR chemicals, catalog number: 28015.294)
Sodium hydroxide (NaOH) (Merck, catalog number: 30620-1KG-M)
Media and buffers
Growth medium (see Recipes)
Differentiation medium (see Recipes)
TBSS buffer (see Recipes)
NBD-lipid stocks (see Recipes)
PMSF stock of 200 mM (see Recipes)
OBAA stock of 5 mM (see Recipes)
BSA solution in TBSS (see Recipes)
Equipment
Autoclave sterilizer (e.g., Systec VX-65, Systec, Linden, Germany)
Biological safety cabinet certified for handling of biological materials (e.g., Herasafe KSP Class II Biological Safety Cabinets, Thermo Fisher Scientific)
Centrifuge with rotor for 15 mL polypropylene tubes (e.g., Eppendorf 5810 R; Wesseling, Germany)
Computer with monitor (e.g., DELL U2415)
Confocal laser scanning microscope (e.g., Leica TCS SP8)
Eppendorf Research® plus pipettes P2, P20, P200, P1000 (Eppendorf, catalog numbers: 3123000012, 3123000039, 3123000055, 3123000063)
Flow cabinet to work with organic solvents
Freezers -20 °C and -80 °C
Glass desiccator Boro 3.3 with socket in lid, 20 cm, including stopcock (BRAND, catalog number: 65238)
Hamilton 700 Series Syringes 10, 25, 100, and 1,000 μL (Hamilton Company, Nevada, USA)
Incubator with humidity and gas control to maintain 37 °C and 95% humidity in an atmosphere of 5% CO2 in air (e.g., Binder, Tuttlingen, Germany)
Incubator to maintain 20 °C (e.g., incubator with Peltier elements heating up and cooling down seamlessly in one system, Memmert, Schwabach, Germany)
Inverted phase contrast microscope equipped with a 10× objective (HI PLAN I 10×/0.22 PH1; Leica DMi1, Mannheim, Germany)
Neubauer counting chamber (improved dark lines, 0.1 mm) and cover glasses (20 × 26 × 0.4 mm)
Pipette tips 2, 10, 200, and 1,000 μL (Sarstedt, catalog numbers: 70.1130.212, 70.760.002, 70.3030.020, and 70.3050.020)
Refrigerator
Tubing (BRAND, catalog number: 143275)
Vortex mixer (e.g., Vortex Genie 2 Scientific Industries Inc., catalog number: SI-0236)
Vacuum Pump V-100 with Interface I-100 (Buchi, Switzerland, catalog numbers: 11593636 and 11593655D)
Water distillation system
Water bath (e.g., WPE45 Memmert, Schwabach, Germany) for mammalian cells and for NBD-lipid labeling (e.g., Julabo CORIO C-BT5, catalog number: 9011305)
Software
ImageJ (Wayne, Rasband, S., U. S. National Institutes of Health, Bethesda, Maryland, USA, https://imagej.nih.gov/ij/index.html, version v.153q)
Leica Application Suite AF (LAS X 3.5.7.23225, Leitz, Wetzlar, Germany)
Leica’s LAS X Office (LASX Office, ver. 1.4.4 26810, Leica, Wetzlar, Germany)
Procedure
The procedure below outlines four main steps: (A) preparation of mammalian cells, (B) cell differentiation, (C) cell counting, and (D) NBD-lipid uptake assay. This procedure was originally applied to study NBD-lipid internalization in mouse myoblast C2C12 cells (Grifell-Junyent et al., 2022). If this procedure is to be used with other cell types, adjustments might be required regarding the choice of cell culture medium, the requirement for coated surfaces, the cell density for microscopy, and possible detachment during incubation at low temperature (20 °C).
Preparation of mammalian cells
Grow adherent cells in sterile culture vessels (T-75 flask) in growth medium (see Recipes) in a tissue culture incubator (37 °C, 5% CO2, 95% humidity) until they reach ~60%–70% confluency.
Note: C2C12 cells will differentiate if grown too confluent and start to fuse. Differentiation and fusion are accompanied by transcriptional changes, which can lead to different results.
Aspirate and discard growth media with a sterile serological pipette.
Wash cells twice with 5 mL of HBSS (Ca2+ and Mg2+ free, pre-warmed at 37 °C) using a sterile serological pipette.
Add 1.5 mL of trypsin-EDTA solution (pre-warmed at 37 °C) using a sterile serological pipette and incubate the T-75 flasks in a tissue culture incubator (37 °C, 5% CO2, 95% humidity). Tilt the vessel back and forth a few times to make sure the thin layer of trypsin is evenly spread.
After 5 min, check for detachment by gently tilting the vessel and/or observing under an inverted microscope. If all cells have not detached in 5 min, incubate for an additional 1–2 min and check again. Continue to incubate and check as necessary, only until cells are no longer attached to the plate surface.
Note: Avoid prolonged incubation period with trypsin-EDTA solution.
Stop trypsinization by adding 7.5 mL of growth medium (pre-warmed at 37 °C, see Recipes) to the cell suspension.
Transfer the cell suspension into a 15 mL Falcon and set aside 100 μL in a 1.5 mL microcentrifuge tube for cell counting, e.g., using the hemocytometer (see section C).
Centrifuge cells in the 15 mL Falcon tube at 300× g for 5 min at room temperature and discard the supernatant to remove the trypsin-EDTA-containing medium from the cells.
Add 10 mL of fresh growth medium (pre-warmed at 37 °C, see Recipes) to the cell pellet and re-suspend completely by gently pipetting up and down using a serological pipette.
Note: Cells in suspension settle quickly. After counting, we recommend gently re-suspending the cell suspension approximately every 2–3 min when seeding multiple dishes.
After counting (see Section C), seed 1.5 × 104 cells per 35 mm polymer bottom dishes and add growth medium (pre-warmed at 37 °C, see Recipe 1) up to a final volume of 1 mL per dish.
Note: Prepare at least one dish per lipid to be analyzed plus one more for a negative control. We used a low cell number for seeding because C2C12 is a fast-growing cell line (doubling time: ~20 h) and this cell number guarantees that single cells are still present when the assay is performed the next day.
Keep the cells in a tissue culture incubator (37 °C, 5% CO2, 95% humidity) overnight.
Cell differentiation
To test the lipid uptake abilities and changes during differentiation of C2C12 cells, grow cells in 35 mm polymer bottom dishes in growth medium (pre-warmed at 37 °C, see Recipes) in a tissue culture incubator (37 °C, 5% CO2, 95% humidity) until they reach 100% confluency.
Note: To check for changes during differentiation and fusion, we analyze cells before they grow confluent (indicated as day -1), cells when they reach 100% confluency (indicated as day 0), and cells that were grown in differentiation medium for a specific number of days (Grifell-Junyent et al., 2022).
After reaching 100% confluency, aspirate and discard the growth medium.
Wash cells twice with 1 mL of HBSS (Ca2+ and Mg2+ free, pre-warmed at 37 °C).
Add 1 mL of differentiation medium (pre-warmed at 37 °C, see Recipes) per 35 mm polymer bottom dish and keep the cells in a humidified tissue culture incubator (37 °C, 5% CO2, 95% humidity).
Change the differentiation medium (pre-warmed at 37 °C, see Recipes) on a daily basis.
Note: We culture the cells up to seven days in differentiation medium.
Cell counting
The purpose of this step is to quantify the cell concentration to resuspend the cells at the appropriate concentration for the NBD-lipid uptake assay. We routinely use the trypan blue hemocytometer assay. Alternative cell counting methods, such as automatic cell counters, may be used.
Prepare the hemocytometer by cleaning the chambers and coverslip with ethanol. Dry the hemocytometer by using lint-free tissue. Place the glass coverslip over the counting chambers.
Note: The correct placement is indicated by the appearance of the Newton rings.
Add 100 μL of 0.4% trypan blue stock solution to 100 μL of cell suspension (step A6) to obtain a 1:1 dilution using a P200 Eppendorf pipette.
Load the hemocytometer with 10 μL of cell suspension per counting chamber with a P20 Eppendorf pipette and examine immediately under an inverted phase contrast microscope at low magnification (e.g., 5×–10× magnification).
Count the number of viable (seen as bright cells) and non-viable cells (stained blue) in the large outer quadrants.
Calculate the percentage of viable cells: % viable cells = [1.00 - (number of blue cells ÷ number of total cells)] × 100. Cell viability should be at least 95%.
Calculate the cell concentration based on the premise that each square accounts for a volume of 10-4 mL of cell suspension.
To obtain the total number of viable cells per milliliter of aliquot, multiply the total number of viable cells by 2 (the dilution factor for trypan blue) and the correction factor of 104 (volume of each square).
NBD-lipid uptake assay (see Figure 2)
The assay is typically performed at 20 °C to reduce endocytosis. For this, all solutions are pre-warmed at 20 °C before use and all incubation steps are performed in an incubator at 20 °C.
Prior to the start of the assay, prepare BSA solution in TBSS (see Recipes).
Prepare conical bottom glass tubes with NBD-lipid stocks (see Recipes) suspended in 10 μL of DMSO using a P20 Eppendorf pipette for each cell sample.
To prepare the cells for labeling, carefully aspirate and discard the growth medium using a P1000 Eppendorf pipette.
Wash cells twice with 1 mL of TBSS (pre-warmed at 20 °C, see Recipes) using a P1000 Eppendorf pipette.
After washing, add 1 mL of TBSS (pre-warmed at 20 °C, see Recipes) using a P1000 Eppendorf pipette.
To block the conversion of NBD-lipids by cellular phospholipases, add 5 μL of PMSF (see Recipes) and 1 μL of OBAA (see Recipes) to the cells using a P2 and P20 Eppendorf pipette to a final concentration of 1 mM and 5 μM, respectively,
Gently mix by tilting the dish and incubate for 10 min at 20 °C in an incubator.
To start labeling, remove 200 μL of TBSS from the cells in the dish, transfer into the conical bottom glass containing the DMSO NBD-lipid suspension (see step D2), mix well using a P200 Eppendorf pipette, and transfer the NBD-lipid suspension to the cells in the dish.
Gently mix by tilting the Petri dish and incubate for 60 min in a tissue culture incubator at 20 °C.
Note: If different NBD-lipids are to be tested, it is recommended to examine them one after the other rather than in parallel, as the time of microscopy would otherwise result in different timeframes of incubation.
After incubation, mount the dish on the microscope for imaging prior to BSA wash.
Note: At this point, cells can be tested for metabolic conversion of the NBD-lipids using lipid extraction and thin layer chromatography analysis, as previously described (Herrera et al., 2022).
For subsequent BSA back extraction, remove the dish from the microscope.
Remove TBSS and wash cells twice with 0.5 mL of BSA solution in TBSS (pre-warmed at 20 °C, see Recipes) for 1 min using a P1000 Eppendorf pipette.
Note: We routinely use a BSA concentration of 5% (w/v) for NBD-lipid extraction. However, the volume of BSA required for extraction, as well as the incubation time, may vary depending on cell type and lipid analog used in the assay (Fellmann et al., 2000). To determine the optimal conditions, label the cells at 4 °C and measure the cell-associated fluorescence after a different time of contact of the cells with BSA.
Add 1 mL of TBSS (pre-warmed at 20 °C, see Recipes) using a P1000 Eppendorf pipette and image on a confocal microscope.
Figure 2. Schematic illustration of the nitrobenzoxadiazole (NBD)-lipid uptake assay via confocal microscopy. For labeling, cells pre-incubated with phospholipase inhibitors at 20 °C are supplemented with NBD-lipids and then incubated for up to 60 min to allow lipid internalization. After a TBSS wash, cells are analyzed using confocal fluorescence microscopy. Uptake of NBD-lipids results in intracellular labeling, while non-internalized NBD-lipids are found at the plasma membrane. Subsequently, back exchange with BSA is performed to remove non-internalized NBD-lipids at the plasma membrane, which allows visualizing only the internalized portion of the NBD-lipids.
Data analysis
Data acquisition
Images were acquired with a Leica TCS SP8 confocal laser scanning microscope (Leitz, Wetzlar, Germany) equipped with a 63×/1.20 water objective. For detailed imaging settings, see Table 1. All images were acquired at the same resolution, magnification, and orientation. This allows direct comparison of images and saves time when arranging figures.
Turn on the confocal laser scanning microscope and the laser, start the computer, and open the LAS X program.
The white light laser (WLL) intensity should be at 85%. Set the scan speed to 400 Hz. Situate the excitation to 488 nm with an intensity of 17.5%. Select a detector and switch it on. Configure the beam path manually to an emission of 497–600 nm with a gain of 20.3. For detailed imaging settings, see Table 2.
Note: The employed settings can be saved on the instrument to be reused in future experiments by selecting an image of a previous experiment with a right click and selecting Apply Image Settings.
Table 2. Parameters employed for confocal laser scanning microscopy analysis of the mammalian cell line C2C12 labeled with NBD-lipids. Settings employed for each parameter, including scanning, magnification, and information about the used objective, imaging settings, and intensities of the used lasers, excitation, and emission parameters. Note that these values might require adjustment depending on the cell line and/or microscope used.
Name Value
Scan mode XYZ
Logical size X/Y/Z 512/512/1
Physical length X/Y/Z 184.52 μm/184.52 μm/0 μm
Scan direction X Unidirectional
Scan speed 400 Hz
Magnification 63
Objective name HC PL APO CS2 63×/1.20
Immersion Water
Zoom 1
Frame average 1
Line average 6
WLL 85%
Laser line 488 nm excitation 17.5%
Emission 497 - 600 nm Gain 20.3
Transmission channel Gain 300
Detector for fluorescence HyD SMD 2 (HyD type detector)
Detector for brightfield PMT Trans (PMT type detector)
Choose the 63× magnification objective (water objective HC PL APO CS2 63×/1.20) and put a drop of water on the objective.
Note: The quality and resolution of an image is highly dependent on the choice of lens. One of the most important factors determining both sensitivity and resolution is the numerical aperture of the lens. To achieve the best possible image quality, it is strongly recommended to select a lens with a high numerical aperture. Care must be taken when using oil immersion objectives, as the 35 mm polymer bottom plates used in this protocol (see Materials) are only compatible with certain immersion oils.
Place the sample on the stage and search for the cells in brightfield mode.
To record the images, both brightfield and fluorescence signal, press the Live button. With the fluorescence signal on screen, search for representative areas and take several pictures.
Note: Exposure to laser light will cause bleaching. Therefore, it is recommended to adjust the focus in brightfield mode before switching to fluorescent mode and immediately acquire an image.
Save images on the computer as raw data.
Expected results
Depending on the head group and cell type, the lipid analog inserted into the outer plasma membrane leaflet can be internalized by spontaneous flip-flop, by protein-mediated translocation, or endocytosis. In C2C12 cells, for example, disappearance of NBD-phosphatidylserine (PS) from the cell surface is predominantly due to fast protein-mediated translocation across the plasma membrane (and endosomal membranes), resulting in a labeling of various intracellular membranes—including Golgi complex, ER, and endosomes. In contrast, NBD- sphingomyelin (SM) is internalized via endocytic vesicles resulting in the appearance of intracellular fluorescent spots (Figure 3).
Figure 3. Interpretation of nitrobenzoxadiazole (NBD)-lipid labeling patterns.
Cells were incubated with NBD-lipids for 60 min and subjected to back exchange with BSA solution in TBSS. A) In the absence of endocytosis and active transport, NBD-lipids are internalized only when a spontaneous flip-flop occurs. Otherwise, NBD-lipids remain confined to the outer leaflet of the plasma membrane. Therefore, no residual fluorescence is found at the plasma membrane or in the cell interior after the BSA back extraction. B) If endocytosis takes place, fluorescence is mainly observed at the plasma membrane, but a weak signal is also visible inside the cell. After washing with BSA, low fluorescence can be detected inside the cell. C) For active lipid transport systems at the plasma membrane, a strong fluorescent signal is detectable throughout the cell before and after BSA back extraction.
Export the raw image data in a format compatible with ImageJ (e.g., tiff) from the imaging system.
Open the ImageJ software and import the images.
Click on Image in the upper operation row of the interface and change the image type to a 32bit grey scale.
Click again on Image and select the Fire Lookup Table (LUT) in the menu of different options for Lookup Tables.
Note: A LUT is a predefined spectrum of different shades of grey, each corresponding to a designated color. In case of the Fire LUT, a low intensity corresponds to dark grey and thus black and dark purple. A high intensity is converted to a light grey, which in turn corresponds to yellow and white. Therefore, a LUT is a tool to reflect differences in intensity.
Afterwards, select the Color menu under the Image operation and click on Edit LUT.
The LUT Editor will open, and the image colors will change to the corresponding colors of the Fire LUT. Press OK to set the change to the image.
Click on Image in the upper operation row of the interface and change the image type to RGB Color.
Save the images as a .tiff or .jpg file by selecting the File menu in the upper operation row of the interface and press Save As. Representative images are shown in Figure 4.
Note: All images to be compared should be imported and LUT-adjusted simultaneously.
Figure 4. Representative confocal images of proliferating C2C12 wild-type cells. Cells were labeled with the indicated nitrobenzoxadiazole (NBD)-lipids for 60 min and subjected to back exchange with BSA solution in TBSS (pre-warmed at 20 °C, see Recipes). Images were taken by confocal microscopy and color-coded with the Fire Lookup Table to highlight intensity variations. Scale bar, 30 μm.
Quantification of lipid uptake
Import the raw data into Leica’s LAS X Office.
Click on the Quantify tab at the top of the screen and choose Histogram in the left selection area.
Note: To distinguish cells that are in close proximity and to improve the visibility of their outline for more precise gating, the intensity and contrast of the brightfield images can be adjusted.
Using the Polygon drawing tool located above the image, draw gates that encircle the cells closely, as shown in Figure 5.
Gate the cells based on the brightfield image so that gates will appear automatically and simultaneously at the same area on the fluorescence image.
Click on the Statistics bar, located just to the left of the images, to generate a table displaying values for the specific gate in both channels.
Deselect the brightfield channel and export the data table as .csv file format.
Note: The .csv file format is a standard file format that can be imported into other programs like Microsoft Excel or R for further data analysis. The analysis performed here was done on Microsoft Excel.
The exported data will give the pixel size of the drawn gate. Normalize the pixel size of each gate to 10,000 pixels.
Adjust the intensity sum accordingly with the ratio between the original pixel size and a pixel size of 10,000 for each gate.
Calculate the average intensity sum for a given lipid.
Calculate the standard deviation based on the adjusted intensity sum for each gate for a given lipid.
Plot data in a bar diagram (Figure 6).
Note: A negative control without NBD-lipid staining should be used to assess the level of background fluorescence in the sample. This is important because even in the absence of NBD-lipids, there may still be some level of background fluorescence present due to autofluorescence or other sources.
Figure 5. Gating of cells. The microscope images are processed using the Leica Application Suite LAS X software. Cells are selected based on the brightfield image and are then superimposed onto the fluorescence image by the software. The contrast and intensity of the brightfield image has been adjusted to facilitate the identification of the cells.
Figure 6. Average pixel intensity. Images of cells incubated with the indicated NBD-lipids were analyzed by single cell gating and pixel intensity analysis using Leica’s LAS X Office software. All gates are normalized to 10,000 pixels, with the intensity adjusted afterwards. Error bars represent standard deviation. In each case, 35 cells were analyzed from a representative experiment.
Recipes
Buffers were prepared using double-distilled water (ddH2O), which was obtained using an in-house water distillation system. Alternatively, all buffers are prepared using ultrapure water with purification sensitivity of 18 MΩ·cm-1 at 25 °C.
Growth medium
Open a 500 mL flask of high-glucose DMEM medium.
Add 100 mL of FBS.
Optional: Add 5 mL of 100× penicillin-streptomycin solution.
Prepare in sterile cabinet; store at 4 °C.
Differentiation medium
Open a 500 mL flask of low-glucose DMEM medium.
Add 10 mL of HS.
Optional: Add 5 mL of 100× penicillin-streptomycin solution.
Prepare in sterile cabinet; store at 4 °C.
TBSS buffer (1 L)
136 mM NaCl (7.94 g)
2.6 mM KCl (194 mg)
1.8 mM CaCl2 (200 mg)
1 mM MgCl2·6H2O (203 mg)
0.36 mM NaH2PO4·2H2O (56 mg)
5.56 mM glucose (1 g)
5 mM HEPES (1.2 g)
Adjust pH to 7.4 with 1 M NaOH. Complete volume to 1 L. Sterilize by filtering using a 0.22 μm filter. Store at 4 °C up to several months.
NBD-lipid stocks
All steps must be performed in glass tubes in order to prevent nonspecific binding of lipids.
Note: Chloroform and methanol are chemical hazards. Do not breathe gas/fumes/vapor/spray. Wear suitable protective clothing. Work in a fume hood.
Lipids are received suspended in chloroform, packaged in sealed glass ampoules; store at -20 °C until use.
To prepare lipid stocks, transfer a volume corresponding to 100 μg from the Avanti glass ampoule into glass screw neck vials. Evaporate the solvent from the glass vials in a desiccator at 250 mbar for 3 h with an additional 1 h incubation at 30 mbar. Close the vials with screw caps and parafilm and store at -20 °C until further use.
Remove desired lipid stocks from the freezer, place on ice, and dissolve in 1 mL of chloroform:methanol (1:1, vol) to a final lipid concentration of 100 μg/mL.
i. Use a glass syringe to transfer the desired volume of NBD-lipid stock solution into a 12 mm diameter glass tube. Typically, we use 5 nmol of NBD-lipid per dish.
ii. Dry the lipids under a 250 mbar vacuum overnight or under a gentle stream of nitrogen gas for 30 min to 1 h so that a dried lipid film is formed at the bottom of the tube.
iii. Resuspend the NBD-lipids in 10 μL of DMSO shortly before use. Add the DMSO in a circular fashion by placing the pipette tip onto the walls of the glass tube and on top of the dried lipid film. Slowly pipette up and down repeating the circular movement until all lipid film is resuspended.
Note: DMSO lipid suspensions are prone to precipitation, owing to their hygroscopic nature, and should be freshly prepared. DMSO is a chemical hazard. Wear suitable protective clothing.
PMSF stock of 200 mM
Weigh 34.838 mg with protective gear to avoid inhalation or contact with skin.
Dissolve in 1 mL of ethanol.
Store at -20 °C.
Note: PMSF is a chemical hazard, especially in its solid state, with high toxicity if swallowed and resulting in severe skin burns and eye damage upon contact. Wear suitable protective clothing and work in a fume hood when preparing the stock solution. Handle the stock solution carefully.
OBAA stock of 5 mM
Weigh 2.14 mg.
Add 1 mL of ethanol.
Store at -80 °C.
Note: Wear suitable protective clothing.
BSA solution (essentially fatty acid-free, 5% w/v) in TBSS
Weigh 200 mg of BSA in a 15 mL Falcon tube.
Add 4 mL of TBSS.
Incubate at 37 °C in water bath until dissolved.
Store in fridge until next day and use within one week; do not freeze the solution.
Note: This volume of 4 mL is enough to analyze the uptake of four NBD-lipids.
Acknowledgments
This protocol was adapted from our previous work (Grifell-Junyent et al., 2022; Herrera et al., 2022). The work was supported by the Lundbeckfonden (R221-2016-1005 to T.G.P.) and an instrument grant from the Deutsche Forschungsgemeinschaft (INST 213/886-1 FUGG to T.G.P.).
Competing interests
The authors declare that no competing interests exist.
References
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Cuticular Hydrocarbon Profiling by Fractionation and GC-MS in Socially Parasitic Ants
HI Hironori Iwai
NK Nobuaki Kono
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4772 Views: 242
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Original Research Article:
The authors used this protocol in Frontiers in Ecology and Evolution Jun 2022
Abstract
Ants use cuticular hydrocarbon (CHC) as a semiochemical for recognizing their nestmates. For socially parasitic ants, deceiving the CHC is an important survival strategy. Profiling and quantifying CHC is a potent approach to understanding such nestmate discrimination behavior. Thus, a highly efficient, stable, and reproducible extraction method for CHC is essential for this purpose. This paper describes a method for socially parasitic ants to disguise the host species’ CHC profile under laboratory conditions, as well as the extraction and measurement of CHC from ants (from a previous study). First, the artificial isotopic substance is applied to the host worker; then, the socially parasitic ant disguises the host-like CHC profile against the above host worker. Next, the CHC is extracted and fractionated from a socially parasitic ant using hexane and silica gel. After concentrating the fractionated product, this product is then used for measurement by gas chromatographymass spectrometry (GC-MS). The CHC extraction protocol described in this paper may be used for various ant species.
Keywords: Cuticular hydrocarbon (CHC) Social insect Myrmecology Ant Social parasite Chemical disguise Gas chromatographymass spectrometry (GCMS) Semiochemical
Background
Cuticular hydrocarbon (CHC) is the generic term for dozens of hydrocarbons on the surface of insect cuticles. CHC protects insects from desiccation and functions as a semiochemical (Howard and Blomquist, 2005; Sturgis and Gordon, 2012). In particular, the role of CHC in the ant social system has been studied in detail. For example, CHC is used to discriminate between nestmates and non-nestmates (Howard and Blomquist, 2005; Sturgis and Gordon, 2012) and inhibit workers’ oviposition (Howard and Blomquist, 2005; Kocher and Grozinger, 2011). Furthermore, some parasitic species take advantage of this CHC-based nestmate discrimination mechanism to invade the nests of host ants (Dettner and Liepert, 1994; Lenoir et al., 2001; Howard and Blomquist, 2005; Akino, 2008; de la Mora et al., 2020). For example, when the newly mated Polyrhachis lamellidens and Polyergus samurai queens invade a host colony, they perform rubbing behavior against the host worker or kill the host queen. These behaviors help the newly mated queen to disguise her CHC profile and match that of the host ants. By disguising the CHC profile of the host ants, the newly mated queen is recognized as a nestmate, allowing her to achieve the early stages of social parasitism (Tsuneoka and Akino, 2012; Iwai et al., 2022).
Behavioral tests and bioassays using ants, as well as profiling and quantifying CHC, are potent approaches to understanding such nestmate discrimination and chemical disguise behavior. A highly efficient, stable, and reproducible extraction method for CHC is essential for this purpose. In addition, reproducing the chemical disguise behavior of socially parasitic ants in the laboratory is necessary. In this paper, we describe a method for CHC extraction and measurement, as well as the induction of chemical disguise by a socially parasitic ant under laboratory conditions, which has been established as an actual experimental system, using P. lamellidens and its host Camponotus japonicus as examples, referring to the practical techniques used in our previous study (on Polyrhachis and Camponotus species) (Iwai et al., 2022; Kurihara et al., 2022). In addition, we describe the method for the tracing assay using artificial isotopic substances to confirm the transition of the CHC profile from the host species to a socially parasitic ant.
First, the artificial isotopic substance is applied to the host worker; then, the socially parasitic ant performs the chemical disguise against the host worker. The crude extract, containing mainly CHC and polar compounds, is obtained from an anesthetized ant with hexane. Then, CHC is separated to remove other substances using silica gel. The fractionated product is concentrated by volatilizing the solvent with N2 gas. A portion of this concentrate is then used for measurements in the gas chromatographymass spectrometry (GC-MS) system. As a result, several ant-derived CHC peaks are detected. This CHC extraction protocol is mainly for large species such as Polyrhachis and Camponotus, although it should be possible to use for small species by adjusting the number of ants used per sample. In addition, since the main targets of CHC profile disguising in socially parasitic ants are adult host workers and host queens, the use of artificial isotopic substances described in this paper should also be applicable to other species.
Materials and reagents
Autosampler vials for Agilent Technologies (Tokyo Garasu Kikai Co., Ltd., catalog number: LH520867-100)
Shimadzu LabTotal vial for GC/GC-MS (Shimadzu GLC Ltd., catalog number: 227-34002-01)
SIL target Polyspring inserts 100/pk (Shimadzu GLC Ltd., catalog number: GLC4010-S630)
Glass capillary tubes, end-to-end tip 100 mm 100 μL, 100 pieces (AS ONE Co., Ltd., catalog number: 3-5998-13)
Pyrex® 10 mm × 75 mm disposable rimless culture tubes, bulk pack (Corning Inc., catalog number: 99445-10)
9" disposable Pasteur pipettes, borosilicate glass/non-sterile, 229 mm, 144 pieces/box × 5 boxes (Fisher Scientific, catalog number: 13-678-20C)
Quartz wool 2–6 μm (10 G) (AS ONE Co., Ltd., catalog number: 6-570-01)
Hexane for chromatography 500 mL (FUJIFILM Wako Pure Chemical Co., Ltd., catalog number: 086-01166)
Wakogel® C-200 for column chromatography, 500 g (FUJIFILM Wako Pure Chemical Co., Ltd., catalog number: 237-00075)
Balance dishes (Ina-optika Co., Ltd., catalog number: AS-DM)
Kimwipe S-200 (Nippon Paper Crecia Co., Ltd., catalog number: 62011)
Parafilm 2" 250 ft (Bemis Co., catalog number: PM992)
Sterilization Petri dish (shallow type AY) (Sansei Medical Co., Ltd., catalog number: 01-003)
C7–C40 saturated alkanes, standard certified reference material, 1,000 μg/mL each component in hexane (Supelco, catalog number: 49452-U)
n-Docosane (C22H46) (FUJIFILM Wako Pure Chemical Co., Ltd., catalog number: 047-08032)
n-Triacontane-d62 (C30D62) (C/D/N Isotopes, CAS: 93952-07-9)
n-Dotriacontane-d66 (C32D66) (C/D/N Isotopes, CAS: 62369-68-0)
Plastic case (5.0 cm × 4.5 cm × 2.5 cm)
Plaster (Katei Kagaku Kogyo Co., Ltd., catalog number: 4905488024103)
Isotope-labeled triacontane and dotriacontane solutions (16 mg/mL in hexane) (see Recipes)
Internal standard material (10 ng/μL docosane in hexane) (see Recipes)
Standards of saturated alkanes (0.5, 1, 2, 5, and 10 ng/μL) (see Recipes)
Equipment
Digital scale (Shimadzu GLC Ltd., catalog number: TWC623N)
Stand (three legs) stainless steel (Yamanaka, catalog number: 1-9789-02) with clamp
Erlenmeyer flask
eVol® XR kit (SGE Analytical Science Pty., Ltd., catalog number: 2910200)
Pasteur pipette bulb, Color Dropper (Tokyo Garasu Kikai Co., Ltd., catalog number: 161-23-03-32)
Agilent 6890N (Agilent Technologies, catalog number: 6890N)
Agilent 5973 MSD (Agilent Technologies, catalog number: 5973 MSD)
Inlet liner, ultra inert, splitless, single taper, glass wool, 5/pk (Agilent Technologies, catalog number: 5190-3163)
HP-5 MS (length 30 m, diameter 0.250 mm, film thickness 0.25 μm) (Agilent Technologies, catalog number: 19091S-433)
N2 gas cylinder (Shonai Gas Co., Ltd.)
Pressure regulator for N2 gas YR-70-1 (GL Sciences, catalog number: 3-703-0010)
Silicon tube 1.5 × 2.5 1 m (AS ONE Co., Ltd., catalog number: 6-586-04)
Dry block bath test tube concentration MC1020 for 20 holes MC-1020 (AS ONE Co., Ltd., catalog number: 1-5144-01)
Mini disk rotor (BIO CRAFT Co., Ltd., catalog number: BC-710)
Software
MSD ChemStation E.02.02.1431 (Agilent Technologies)
Procedure
Application of artificial isotopic substances for the tracing assay
Note: As this protocol uses organic solvents (e.g., hexane), we do not recommend the use of plastic instruments to avoid contamination by impurities from these instruments.
Dissolve n-Triacontane-d62 and n-Dotriacontane-d66 individually in hexane (16 mg/mL). Add 100 μL of both solutions to a SIL target Polyspring inserts in the autosampler vial for Agilent Technologies. To allow the ants to breathe in the later process, drill a hole in the lid of this vial.
Stir the solution with a glass capillary tube; meanwhile, spray nitrogen gas to evenly apply the above artificial isotopic substances over the entire inner wall of the insert, using a dry block bath test tube (already installed on another stand, with silicon tube, pressure regulator for N2 gas, and N2 gas cylinder) (Figure 1).
Figure 1. Evaporation system of the solvent. A. The appearance of evaporation. B. The process of spraying N2 gas.
Expose a single individual live host worker (e.g., C. japonicus) in a sterilization Petri dish to 4 °C for 2 min and then to -20 °C for 3 min, for anesthetic purposes (Figure 2).
Figure 2. Anesthetizing of an ant. A. Before anesthetizing. B. After anesthetizing.
After the solvent is completely volatilized, carefully lift the sterilization Petri dish containing the anesthetized host worker, taking care not to touch it directly with the hands. Move it from the head side into the entrance of the SIL target Polyspring inserts, where the artificial isotopic substances are applied to the inner wall (Figure 3). Gently shake the insert for 1 h using a mini disk rotor. If the anesthetized host worker is alive, it wakes up in this step.
Figure 3. Application of artificial isotopic substances to an ant. An ant is put into the vial with micro inserts coated with artificial isotopic substances.
Induction of chemical disguise behavior under laboratory conditions
To prevent possible attacks from the host worker (C. japonicus) before the newly mated P. lamellidens queen performs the rubbing behavior, anesthetize the host worker with artificial isotopic substances. Use the same method of anesthesia as in step A3; however, to minimize the risk of the artificial isotopic substances coming off from the host worker, the autosampler vial with SIL target Polyspring inserts containing the host worker with the lid removed should be placed in the Petri dish, allowing the host worker to move around in the dish on its own.
Place the newly mated P. lamellidens queen and host workers in plastic cases containing plaster (Figure 4). After dissolving the plaster in water and placing it in a plastic case, make sure the plaster has hardened in the case before using it in the experiment. The plaster acts as both a scaffold for the ants and a moisturizing factor. If a newly mated P. lamellidens queen is able to perform the rubbing behavior continuously, one new host worker with artificial isotopic substances should be added (all from the same colony) every day until the third day. The anesthetized host worker usually wakes up within approximately 10 min of being introduced.
Figure 4. The rubbing behavior of a newly mated P. lamellidens queen. The newly mated P. lamellidens queen exhibits rubbing behavior against a C. japonicus worker under laboratory conditions. Adapted from Iwai et al. (2022).
Extraction and fractionation
Note: Ants of approximately 1–2 cm in size are used as a model case in this protocol. If the ant size is less than 1 cm, the number of ants should be increased per sample (e.g., three ants per sample).
The conditions for selecting the internal standard are: 1) selecting a substance that has similar properties to the CHC of the ants used in the experiment and 2) that is not present naturally in these species. It is necessary to prepare a suitable internal standard (hydrocarbon with a similar chain length to the CHC of the ants) corresponding to that species.
Expose a single individual live ant (e.g., the newly mated P. lamellidens queen or its host worker) in a sterilization Petri dish to 4 °C for 2 min and then to -20 °C for 3 min for anesthetic purposes (similarly to step A3).
Immediately place the anesthetized ant in a disposable rimless culture tube containing 200 μL of hexane mixed with docosane (10 ng/μL) as an internal standard material (Figure 5). The above solvent is accurately measured by the digital syringe eVol® XR. The whole body of an ant should be immersed in hexane.
Figure 5. An ant is immersed in the solvent (docosane in hexane). Ensure that the entire ant body is immersed.
The chemical materials from the body surfaces of the ant are extracted by dipping the ants in hexane for 5 min. The ant will die during this step.
Add a Pasteur pipette to the stainless-steel stand (three legs) with a clamp and wrap a Kimwipe S-200 around the surface of the Pasteur pipette (Figure 6A). The Kimwipe prevents contamination of the rubber part of the clamp even if a small amount of hexane is accidentally spilled while applying it to the Pasteur pipette.
Figure 6. Extraction and fraction of CHC. A. Joint with the clamp. B. Placement of quartz wool. C. Placement of Wakogel® C-200. D and E. Injection of sample or hexane.
Add quartz wool to the inside of the Pasteur pipette (Figure 7A). The amount of quartz wool used should be sufficient to fill the bottom hole of the Pasteur pipette. At this point, cram the quartz wool into the bottom using another pipette (Figure 6B and Figure 7A).
Figure 7. Preparing a column for fractionation. A. Steps for loading Quartz Wool. B. Steps for loading Wakogel® C-200.
For column chromatography, add 0.5 g of Wakogel® C-200 to the Pasteur pipette with the quartz wool. Measure the weight of the silica gel using balance dishes and a digital scale (Figure 6C and Figure 7B).
Using another Pasteur pipette, add approximately 1 mL of hexane into the above Pasteur pipette with a Pasteur pipette bulb to wash and deaerate the silica gel. Drip the added hexane through the fractionation column into an empty Erlenmeyer flask using a Pasteur pipette bulb (Figure 6D). As 1 mL of hexane will flow down under its own weight within approximately 10 s of being placed on the Pasteur pipette column, it is necessary to prepare an Erlenmeyer flask before performing this step.
Carry out fractionation by adding the ant extract into the above Pasteur pipette. At this time, dispense the ant extract obtained in steps C1 and C2 using another Pasteur pipette with a Pasteur pipette bulb (Figure 6E).
Add 1 mL of hexane to the column after applying the ant extract using another pre-marked 1 mL scale Pasteur pipette equipped with a Pasteur pipet bulb to elute the hydrocarbons. Drip the fractionated product through the fractionation column into an empty autosampler vial using a Pasteur pipette bulb. This process can only fractionate CHC from ant crude extract. As the fractionated product will flow down under its own weight within approximately 10 s of being placed on the Pasteur pipette column, it is necessary to prepare an empty autosampler vial before performing this step. Approximately 1 mL of the fractionated product is yielded in this step. A new column and an empty vial must be prepared for each sample.
Concentration
By setting the autosampler vial containing the fractional product in a dry block bath test (already installed on another stand with silicon tube, pressure regulator for N2 gas, and N2 gas cylinder), the obtained hydrocarbons are concentrated by applying N2 gas until the solvent hexane is completely evaporated.
To elute the concentrated hydrocarbons, add 50 μL of hexane to the autosampler vial using the digital syringe eVol® XR. After adding 50 μL of hexane, elute hydrocarbons adhering to the inner wall by tilting and turning the autosampler vial.
Set the SIL target Polyspring inserts in the Shimadzu LabTotal Vial for GC/GC-MS and transfer 50 μL of eluted hydrocarbons into the insert using the glass capillary tubes.
Analysis using the GC-MS system
To analyze the samples (concentrated hydrocarbons), set up the GC-MS system under the following conditions. The splitless mode is adopted for the sample injection port and the apparatus is maintained at 300 °C. Helium is used as the carrier gas at a constant flow rate setting of 0.9 mL/min. The oven temperature is set as follows: 40 °C for 3 min, 40–260 °C at 30 °C/min, 260–300 °C at 15 °C/min, and 18 min at the final temperature. See Equipment for the liner and column used.
Inject 2 μL of the sample (concentrated hydrocarbons) into the sample injection port of the GC.
Inject also C7–C40 saturated alkanes of known concentrations (0.5, 1, 2, 5, and 10 ng/μL) to use as standard substances.
Data analysis
To detect and calculate each hydrocarbon peak and its areas, use the MSD ChemStation E.02.02.1431. The equivalent chain length (ECL), the value of fragment and molecular ions, and the mass spectrum pattern are used to estimate the type of each CHC (Figure 8).
Figure 8. The estimation of each cuticular hydrocarbon (CHC) of a newly mated P. lamellidens queen (before the rubbing behavior). A. Total ion chromatogram of CHC. B. The fragment pattern of n-Docosane (C25H52, internal standard). C. The fragment pattern of n-Pentacosane (C25H52). D. The fragment pattern of n-Triacontane-d62 (C30H62). A list of other CHCs of a newly mated P. lamellidens queen can be found in Iwai et al. (2022).
Quantify the concentrations of hydrocarbons using standards (the mixtures of C7–C40 saturated alkanes) of known concentrations (0.5, 1, 2, 5, and 10 ng/μL) diluted at five levels; add the internal standard (docosane) to each sample and to the above standards. After normalizing the peak areas of each standard and sample by using internal standards, create a calibration curve to quantify the concentration of hydrocarbons.
The calculated peak area and concentration of each CHC are used in various statistical analyses (e.g., hierarchical cluster analysis, correlation analysis, and quantitative analysis; refer to Iwai et al., 2022).
Recipes
Isotope-labeled triacontane and dotriacontane solutions (16 mg/mL in hexane)
Add 16 mg of n-Triacontane-d62 and n-Dotriacontane-d66 individually to 1 mL of hexane for chromatography. Store at -20 °C to 4 °C.
Internal standard material (10 ng/μL docosane in hexane)
Add 1 mg of Docosane to 100 mL of hexane for chromatography.
This solution should be stored at -20°C to 4 °C, as it is used regularly in experiments.
Standards of saturated alkanes (0.5, 1, 2, 5, and 10 ng/μL)
Add a total of 20 μg of C7–C40 saturated alkanes (1,000 μg/mL of each component in hexane) to 2 mL of hexane for chromatography.
Additionally, prepare other standards of saturated alkanes (0.5, 1, 2, and 5 ng/μL) containing 10 ng/μL docosane as in Table 1.
Table 1. Recipes for standards of saturated alkanes (0.5, 1, 2, and 5 ng/μL)
Standards of saturated alkanes 10 ng/μL of C7–C40 saturated alkanes 10 ng/μL docosane (internal standard) total
0.5 ng/μL 50 μL 950 μL 1 mL
1 ng/μL 100 μL 900 μL 1 mL
2 ng/μL 200 μL 800 μL 1 mL
5 ng/μL 500 μL 500 μL 1 mL
Acknowledgments
This protocol was adapted from our established publication (Iwai et al., 2022; Kurihara et al., 2022). We thank Toshiharu Akino for advising the extraction method of the CHC and the critical suggestions, and Kazuharu Arakawa, Masaru Tomita, and Masataka Wakayama for their continuous support and help. We also thank Mana Masui and Koh Nakagawa for taking pictures, and Kaoru Ogino for his diligent proofreading of this manuscript. This work was supported by research funds from JSPS Fellows (202021677), Taikichiro Mori Memorial Research Grant, Nakatsuji Foresight Foundation Research Grant, KAKENHI Grant-in-Aid for Scientific Research (B) (21H02210), and Yamagata Prefecture, and Tsuruoka City.
Competing interests
The authors declare no competing interests.
Ethical considerations
This protocol does not use human and animal (e.g., mammal) subjects.
References
Akino, T. (2008). Chemical strategies to deal with ants: a review of mimicry, camouflage, propaganda, and phytomimesis by ants (Hymenoptera: Formicidae) and other arthropods. Myrmecol. News 11(11): 173–181.
de la Mora, A., Sankovitz, M. and Purcell, J. (2020). Ants (Hymenoptera: Formicidae) as host and intruder: recent advances and future directions in the study of exploitative strategies.Myrmecol. News 30: 53–71.
Dettner, K. and Liepert, C. (1994). Chemical mimicry and camouflage. Annu. Rev. Entomol. 39: 129–154.
Howard, R. W. and Blomquist, G. J. (2005). Ecological, behavioral, and biochemical aspects of insect hydrocarbons. Annu. Rev. Entomol. 50: 371–393.
Iwai, H., Mori, M., Tomita, M., Kono, N. and Arakawa, K. (2022). Molecular evidence of chemical disguise by the socially parasitic spiny ant Polyrhachis lamellidens (Hymenoptera: Formicidae) when invading a host colony. Front. Ecol. Evol. 10: 915517.
Kocher, S. D. and Grozinger, C. M. (2011). Cooperation, conflict, and the evolution of queen pheromones. J. Chem. Ecol. 37(11): 1263–1275.
Kurihara, Y., Iwai, H., Kono, N., Tomita, M. and Arakawa, K. (2022). Initial parasitic behaviour of the temporary social parasitic ant Polyrhachis lamellidens can be induced by host-like cuticles in laboratory environment. Biol. Open 11(3): bio058956.
Lenoir, A., d’Ettorre, P., Errard, C. and Hefetz, A. (2001). Chemical ecology and social parasitism in ants. Annu. Rev. Entomol. 46: 573–599.
Sturgis, S. J. and Gordon, D. M. (2012). Nestmate recognition in ants (Hymenoptera: Formicidae): a review. Myrmecol. News 16: 101–110.
Tsuneoka, Y. and Akino, T. (2012). Chemical camouflage of the slave-making ant Polyergus samurai queen in the process of the host colony usurpation (Hymenoptera: Formicidae). Chemoecology 22: 89–99.
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© 2023 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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4,773 | https://bio-protocol.org/en/bpdetail?id=4773&type=0 | # Bio-Protocol Content
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Lipidomics Workflow for Analyzing Lipid Profiles Using Multiple Reaction Monitoring (MRM) in Liver Homogenate of Mice with Non-alcoholic Steatohepatitis (NASH)
HW Hai Ning Wee
LL Lye Siang Lee
SH Sharon Hong Yu Han
JZ Jin Zhou
PY Paul Michael Yen
JC Jianhong Ching
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4773 Views: 877
Reviewed by: Durai SellegounderYu Hui KangSayani DasSibapriya Chaudhuri
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Original Research Article:
The authors used this protocol in Nature Communications Sep 2022
Abstract
Non-alcoholic steatohepatitis (NASH) is a condition characterized by inflammation and hepatic injury/fibrosis caused by the accumulation of ectopic fats in the liver. Recent advances in lipidomics have allowed the identification and characterization of lipid species and have revealed signature patterns of various diseases. Here, we describe a lipidomics workflow to assess the lipid profiles of liver homogenates taken from a NASH mouse model. The protocol described below was used to extract and analyze the metabolites from the livers of mice with NASH by liquid chromatography–mass spectrometry (LC-MS); however, it can be applied to other tissue homogenate samples. Using this method, over 1,000 species of lipids from five classes can be analyzed in a single run on the LC-MS. Also, partial elucidation of the identity of neutral lipid (triacylglycerides and diacylglycerides) aliphatic chains can be performed with this simple LC-MS setup.
Key features
• Over 1,000 lipid species (sphingolipids, cholesteryl esters, neutral lipids, phospholipids, fatty acids) are analyzed in one run.
• Analysis of liver lipids in non-alcoholic steatohepatitis (NASH) mouse model.
• Normal-phase chromatography coupled to a triple quadrupole mass spectrometer.
Graphical overview
Schematic procedure for the homogenization and extraction of mouse liver tissue in preparation for LC-MS analysis (Created with BioRender.com)
Keywords: Lipidomics Liver LC-MS Sphingolipids Fatty acids Glycerolipids Phospholipids Cholesteryl esters
Background
Lipids are essential components of energy metabolism that can be disrupted in metabolic diseases. Lipids comprise various classes, each possessing widely different chemical properties, e.g., steroids are much more non-polar than phospholipids. Due to the widely differing properties, it is difficult to have a single method catered to all the different lipids. As a result, different classes of lipids often have dedicated analytical methods, such as various derivatization techniques, columns, and mass spectrometers, which makes it challenging to have an overview of a broad spectrum of lipids in a single analysis (Wu et al., 2020; Zhao et al., 2020). Another issue of lipid analysis pertains to the triglyceride class. With up to three aliphatic chains attached to a glycerol backbone, the task of identifying and quantifying all species is highly challenging and has been extensively discussed, with each technique having its own limitations and advantages (Han and Ye, 2021). Compared to other methods requiring adaptations, such as two-dimensional LC, argentation, or supercritical fluid chromatography, the method described here provides a broad view of various lipid classes using standard methods that can be done in most laboratories with basic training.
In this protocol, we describe one variant of a lipidomics workflow, originally developed by the mass spectrometry company SCIEX, that covers over five classes of lipids (sphingolipids, phospholipids, glycerolipids, cholesteryl esters, and fatty acids) (Figure 1), including a partial identification of a single aliphatic chain within the neutral lipid, triacylglycerol (Ubhi et al., 2016). The workflow uses a variety of lipid standards, including stable isotopes from ceramides, sphingomyelins, cholesteryl esters, phosphatidylcholines, phosphatidylethanolamines, phosphatidylglycerols, phosphatidylinositols, phosphatidylserines, monoacylglycerols, diacylglycerols, and triacylglycerols. The system uses a normal-phased hydrophilic interaction chromatography (HILIC) column, which separates the lipids by classes as opposed to chain length in reverse-phased systems. In total, the multiple reaction monitoring (MRM) method performs a single-point quantitation of over 1,000 lipid species, providing a convenient overview of common and important lipids for studying metabolic diseases. Here, we apply the techniques to studying the lipidome of liver tissue from a mouse model with non-alcoholic steatohepatitis (NASH). These techniques are not limited to studying the liver but can be applied to other tissues, including cell culture.
Figure 1. Key structural features of five broad categories of lipids are included in the lipidomics panel
Part I: Tissue homogenization
Materials and reagents
2 mL cryogenic vial (Corning, catalog number: 430488)
2 mL homogenization tubes (Labcon Co., catalog number: 3661-875-000)
Zirconia beads, 1.0 mm diameter (BioSpec Products, catalog number: 11079110ZX)
0.9% sodium chloride solution (Sigma-Aldrich, catalog number: S8776)
Cylinder of liquid nitrogen
Acetonitrile Optima®, LC-MS grade (Fisher chemical, catalog number: A955-4)
Deionized (MilliQ) water (Arium Pro System, Sartorius AG)
Formic acid, HPLC grade (Merck Supelco, catalog number: 5.43804)
Dry ice
Homogenization buffer (see Recipes)
Recipes
Homogenization buffer
Final concentration Amount
Acetonitrile n/a 498.5 mL
Deionized H2O n/a 498.5 mL
Formic acid 0.3% 3 mL
Equipment
Precellys Evolution tissue homogenizer (Bertin Technologies, France)
Cryolys Evolution cooling unit (Bertin Technologies, France)
Procedure
Pre-chill 0.9% sodium chloride solution on wet ice before animal procedures.
Euthanize the mouse using an approved protocol and remove the liver within 10 min. Wash the liver in cold sodium chloride solution to remove blood on the organ’s surface. Cut the liver into smaller pieces before keeping it in the cryogenic vials, which are snap frozen immediately in liquid nitrogen. The tubes of snap-frozen liver samples can be kept in a -80 °C freezer until ready to be used.
Accurately weigh 50–60 mg of frozen liver tissue samples into 2 mL homogenization tubes and immediately place them on wet ice.
Perform all following procedures on wet ice, unless otherwise stated.
Prepare homogenization buffer (see Recipes).
Add ice-cold homogenization buffer into each homogenization tube to obtain a sample concentration of 50 mg/mL, after assuming a water content of 50 μL in 50 mg of tissue.
Add Zirconia beads with a diameter of 1.0 mm to approximately 25% of the buffer level.
Place the dry ice, the homogenizing tubes containing the liver tissues, the homogenizing buffer, and Zirconia beads in a Precellys Evolution tissue homogenizer equipped with a Cryolys Evolution cooling unit for six cycles of 20 s at 6,500 rpm at 4 °C, with 10 s pause intervals to minimize temperature fluctuations during homogenization.
Aliquot 50 μL of the homogenate and snap freeze in dry ice before storing at -80 °C until further sample processing.
Part II: Lipidomics analysis using a triple quadrupole mass spectrometer
The lipidomics analysis was performed using the protocol provided by SCIEX, as described in the application notes (Ubhi et al., 2016; Ubhi, 2020; Pearson et al., 2023). We selected this protocol because it has been used in many studies and demonstrated good reproducibility (Cao et al., 2019; Loef et al., 2020), accuracy, and precision (Contrepois et al., 2018). The SCIEX LipidyzerTM platform was developed to provide MS-based targeted profiling of over 1,000 lipid species spanning 13 lipid subclasses. The >1,000 lipid analytes included in this lipidomics method fall into five broad categories (Figure 1), based on their basic structural units as described below:
Glycerolipids consist of a glycerol backbone with fatty acids attached to it by ester linkage. They include mono-, di-, and triglycerides (MAG, DAG, TAG).
Sphingolipids consist of a sphingoid long-chain backbone (normally sphingosine or sphinganine) with a fatty acid attached to the C2 position by an amide bond and another moiety (i.e., H, sugar, or phospho-X group) attached at C3. The biologically relevant subclasses include ceramides (CER), dihydroceramides (DCER), lactosylceramides (LCER), hexosylceramides (HCER), and sphingomyelins (SM). SM is also considered a glycerophospholipid.
Glycerophospholipids consist of a glycerol backbone with fatty acids attached to the first two carbon atoms (C1 and C2), while a polar phospho-X headgroup is attached to the C3 position. They include phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylglycerol (PG), phosphatidylinositol (PI), and phosphatidylserine (PS). Lysophospholipids are glycerophospholipids lacking one acyl chain. Hence, their glycerol unit is attached to one fatty acid and one polar phospho-X headgroup. They include lysophosphatidylcholine (LPC), lysophosphatidylethanolamine (LPE), lysophosphatidylglycerol (LPG), lysophosphatidylinositol (LPI), and lysophosphatidylserine (LPS).
Free fatty acids (FA) include acyl chains of various lengths, such as palmitic acid (FA 16:0).
Cholesterol esters (CE) consist of cholesterol with a long-chain fatty acid esterified to its hydroxyl group.
One of the major obstacles that curtail the absolute quantification in MS-based lipidomics method is the matrix effects, whereby the components that co-elute with the analyte of interest may influence its ionization efficiency (Köfeler et al., 2021; Swinnen and Dehairs, 2022). For instance, matrix effects have been known to cause significant ion suppression of phospholipids, e.g., phosphatidylcholine (Guo and Lankmayr, 2011; Khoury et al., 2016) and lysophosphatidylcholine (Morita et al., 2019), in electrospray ionization (ESI)-based methods, making exact quantification difficult. To partially compensate for the influence of matrix effects, we have included an internal standard (IS) corresponding to each of the following lipid classes: SM, CE, CER, DCER, HCER, LCER, TAG, LPC, PC, LPE, PE, and FA. These internal standards also serve to correct for any possible variations during the process of sample preparation and analysis (Wang et al., 2016). Given the large number of lipid analytes (>1,000) in this panel, it is not always possible to find an appropriate internal standard for each lipid class. For lipid classes with a matching internal standard, concentrations of the lipids in samples can be quantified using a single-point quantitation or isotope dilution method. However, for lipid classes where a matching stable isotope internal standard is unavailable, we select one of the internal standards from a different lipid class as its surrogate internal standard and express its concentration as an area-ratio of the analyte to the surrogate internal standard. Therefore, the final data provided by this lipidomics panel should be regarded as relative quantification for lipid species with surrogate internal standards.
Extraction method
The extraction method is based on the protocol developed by SCIEX. Essentially, this method involves the partitioning of lipids in a biphasic mixture of dichloromethane (CH2Cl2) and methanol. We made a slight modification to the volume of extraction solvents used, so that the final volume could fit within a 2 mL microcentrifuge tube. Since the lipid-containing organic phase lies at the bottom of the tube, one must exercise caution when collecting the organic phase using a micropipette (Wong et al., 2019). It is advisable to use micropipette tips of 200 μL or smaller when performing this procedure, so it will be easier to pass through the overlying aqueous phase and non-extractable residue layer to reach the organic phase at the bottom. The detailed protocol for extraction is described below.
Materials and reagents
2 mL microcentrifuge tubes (Corning Axygen®, catalog number: MCT-200-C)
Waters XBridge Amide column 4.6 mm × 150 mm, 3.5 μm (Waters Corp, catalog number: 186004869)
Dichloromethane (CH2Cl2) (Sigma-Aldrich, catalog number: 270997)
Methanol Optima® (MeOH) (Fisher Chemical, catalog number: A456-4)
Ammonium acetate Bioxtra > 98% (Sigma-Aldrich, catalog number: A7330-100g)
Ammonium hydroxide solution (Sigma-Aldrich, catalog number:338818)
Acetonitrile Optima® (Fisher Chemical, catalog number: A955-4)
Internal Standards kit for LipidyzerTM platform (SCIEX, catalog number: 5040156)
Deionized (MilliQ) water (Arium Pro System, Sartorius AG, Germany)
Liquid chromatography solvent A
Liquid chromatography solvent B
Nitrogen cylinder
Internal Standards kit for LipidyzerTM Platform (see Recipes)
Liquid chromatography solvent A (pH 8.2, adjusted using ammonium hydroxide) (see Recipes)
Liquid chromatography solvent B (pH 8.2, adjusted using ammonium hydroxide) (see Recipes)
Recipes
Internal Standards kit for LipidyzerTM Platform
Store standards as per manufacturer’s instructions in a freezer at -25 °C; standards need to be pre-mixed as per the table below.
Internal standard Stock concentration (mg/mL) Volume to mix (μL) Final concentration (μM)
CER(16:0) (d9) 0.02 110 36.57
CE(22:6) (d7) 0.15 110 213.18
TAG(52:1/FA18:0) (d9) 0.14 109 160.95
DCER(16:0) (d9) 0.004 110 7.29
FFA(17:1) 0.05 110 186.23
HCER(16:0) (d9) 0.03 110 42.31
LCER(16:0) (d9) 0.03 110 34.43
LPC(16:0) (d9) 0.1 110 198.14
LPE(18:0) (d5) 0.05 110 102.75
PC(16:0/16:1) (d9) 0.0625 76 84.39
PE(18:0/18:1) (d5) 0.01 109 13.32
SM(16:0) (d7) 0.1 110 140.92
Liquid chromatography solvent A (pH 8.2, adjusted using ammonium hydroxide)
Reagent Final concentration Amount
Ammonium acetate 1 mM n/a
Deionized H2O n/a 50 mL
Acetonitrile n/a 950 mL
Liquid chromatography solvent B (pH 8.2, adjusted using ammonium hydroxide)
Reagent Final concentration Amount
Ammonium acetate 1 mM n/a
Deionized H2O n/a 500 mL
Acetonitrile n/a 500 mL
Equipment
Sorvall Legend Micro 21R microcentrifuge (Thermo Fisher Scientific., catalog number: 75002447)
Vortex-Genie 2 vortex mixer (Scientific Industries, catalog number: SI-0236)
SCIEX Triple Quad 5500 MS System
Agilent 1290 Infinity II LC System
Software
Multi-Quant software (SCIEX)
Procedure
Quality control samples
Ensure that appropriate controls are in place prior to extracting the samples. Required controls include:
Blank solution without internal standards (methanol vehicle only).
Blank solution with internal standards (75 μL of methanol vehicle spiked with 25 μL of internal standard mix).
Pooled quality control samples, made by pooling at least 10% of samples in a large batch study, or an equal volume of each sample into a single tube. These pooled quality control replicates are run regularly over the entire batch, so that any variations can be monitored across the entire run. This is especially important for large study samples that are performed over multiple days.
Extraction procedure
Thaw liver homogenates atop ice; then, pipette 25 μL of each liver homogenate to a 2 mL microcentrifuge tube.
Add 283 μL of deionized H2O to 25 μL of liver homogenate and allow the sample to stand on ice for 10 min.
Pipette 581 μL of MeOH to the mixture.
Pipette 262 μL of CH2Cl2 to the mixture.
Cap the tube and vortex the mixture vigorously for 5 s at maximum speed.
Ensure that the mixture consists of only one single phase. If two distinct phases are observed, add 25 μL of MeOH and vortex, and then check to ensure that a single phase is formed. Otherwise, continue adding 25 μL of MeOH and vortex until a single phase is seen.
Add 25 μL of the premixed lipid internal standard, then vortex and incubate the mixture for 30 min at room temperature.
Pipette 291 μL of deionized H2O to the mixture.
Pipette 262 μL of CH2Cl2 to the mixture.
Gently invert tubes 10 times. DO NOT VORTEX the tube; otherwise, an emulsion will be formed (Video 1).
Video 1. Inversion of tube
Pre-chill centrifuge to 4 °C. Subject the tube to centrifugation at 1,710× g for 10 min.
Transfer the lower organic layer to a fresh microcentrifuge tube (Video 2).
Video 2. Transfer of organic layer
Pipette 581 μL of CH2Cl2 to the remaining extracts in the tube.
Mix by gently inverting 10 times, centrifuge, and then transfer the lower organic layer to the tube from step 12.
Evaporate solvent using a nitrogen blower.
Reconstitute the extracted lipids in 100 μL of MeOH.
LC method and parameters
Chromatographic separation of lipid classes is achieved using a normal-phase column by gradient elution of a 24-min timeframe. The elution order of solutes in normal-phase separation is governed by polarity, with solutes of lower polarity eluting first. Therefore, as a general rule, lipid species within each subclass elute in the order of decreasing chain length (i.e., longer chains elute first) and increasing degree of unsaturation of the fatty acyl groups (i.e., species with more double bonds elute later). The stationary phase (HILIC column) and mobile phases (A and B) are described below:
• Stationary phase: XBridge Amide 3.5 μm, 4.6 mm × 150 mm column.
• Mobile phase A: 1 mM ammonium acetate in 95% acetonitrile.
• Mobile phase B: 1 mM ammonium acetate in 50% acetonitrile (pH to be adjusted by adding ammonium hydroxide).
One should remember to add ammonium hydroxide to adjust the pH of mobile phase B to match the pH of mobile phase A. This is an important step because the retention times on a HILIC column are very sensitive to the pH changes in the mobile phase.
The LC parameters are shown below:
• Gradient
Time (min) Flow rate (mL/min) Solvent A Solvent B
0 0.7 94 6
6 0.7 94 6
10 0.7 75 25
11 0.7 2 98
13 0.7 0 100
13.4 0.7 0 100
13.5 1.5 0 100
18.6 1.5 0 100
18.7 1.5 99.9 0.1
23 1.5 99.9 0.1
23.5 0.7 99.9 0.1
24 Stop
• Oven temperature = 35 °C
MS method and parameters
The method utilizes MRM quantification approach in a triple quadrupole LC-MS instrument to ensure high selectivity of lipid species. Positive ion mode is used to detect SM/CER/DCER/HCER/LCER/TAG/DAG/MAG, while negative ion mode is used to detect PC/PE/PG/PI/PS/FFA.
In MRM, the lipids that have been ionized are scanned in Q1, fragmented to produce product ions in Q2 (also known as the collision cell), and these product ions are scanned in Q3. The first (Q1) and the last (Q3) mass analyzers of the triple quadrupole instrument are used as mass filters to isolate a precursor ion and a corresponding product ion for each lipid species. Generally, the product ions included in this method are either characteristic of the headgroup or the backbone of each lipid class, or they are specific for one of the fatty acids attached to the lipid species. For example, sphingomyelins undergo fragmentation to yield a phosphorylcholine head group as its product ion at m/z 184 (Chen et al., 2010). Ceramides other than sphingomyelins fragment at their sphingoid base backbones to produce either sphingosine at m/z 264 (CER, HCER) or sphinganine at m/z 266 (e.g., DCER, LCER) as their product ions (Chen et al., 2010). Cholesterol esters yield the cholesterol moiety as its product ion at m/z 369.4 (Yu et al., 2014). The phospholipids (PC, PE, PG, PI, PS, LPC, LPE, LPG, LPI, and LPS) fragment to yield one of the fatty acid chains as its product ion (Pi et al., 2016). Fragmentation of glycerolipids (e.g., TAG, DAG, MAG) produces a neutral loss corresponding to one of the fatty acyl chains.
Besides Q1 and Q3 transitions, the mass spectrometry method has to be optimized for collision energy and other parameters. Table A1 in the appendix shows the MRM parameters for each of the lipid metabolites. These parameters will need to be re-optimized in different triple quadrupole instruments.
The MRM scans are scheduled, which means that the instrument only scans for each Q1/Q3 transitions at a specific window, corresponding to the time when the particular lipid class elutes. Therefore, it is important to ensure that the MS peak of interest does not drift out of the scan window as a result of retention time shifts (usually due to changes in mobile-phase composition or column aging). Thus, every time after column or solvent changeovers, it is important to check the retention time to ensure that accurate and appropriate scan windows are set for each lipid of interest. The importance of this is highlighted in other lipidomics protocols as well (Mukhamedova et al., 2020). A typical chromatogram of the representative lipid standards is shown in Figure 2.
Figure 2. Chromatogram of lipid internal standards. 1: CE(22:6) (d7); 2: CER(16:0) (d9); 3: DCER(16:0) (d9); 4: FFA(16:0) (d9); 5: FFA(17:1); 6: TAG(52:1_FA18:0) (d9); 7: HCER(16:0) (d9); 8: PE(18:0_18:1) (d5); 9: PC(16:0_16:1) (d9); 10: SM(16:0) (d7); 11: LCER(16:0) (d9); 12: LPE(18:0) (d5); 13: LPC(16:0) (d9).
Data analysis
Data integration and analysis
Each peak area is integrated using Multi-Quant software. Default settings of peak width, peak height, noise, baseline, and peak splitting on Multi-Quant should generally be able to screen peaks with reasonable quality for integration. Changes to default values should be done with care if optimization is required. A clear and easy reference for using Multi-Quant can be found on https://www.youtube.com/watch?v=yjNvcf9CEL0, an instructional video created by SCIEX. After peak integration, peak areas and the area ratios to internal standards can be exported from the software into Microsoft Excel table format for further processing. To determine the relative concentration of each lipid, the area of each lipid of interest is divided by the area of the relevant internal standard. Normalized fold-change can be computed for each lipid species by dividing the area-ratio of each sample by the mean area-ratio across all the samples. A heatmap can be generated using the normalized fold-change values. It is generally accepted that at least three to five biological replicates should be tested in order to get reasonable data for statistical analysis.
For lipid species with the corresponding internal standards (i.e., SM, CE, CER, DCER, HCER, LCER, TAG, LPC, PC, LPE, PE, and FA), the concentration can be computed by dividing the area of peak by the area of the internal standard and multiplying by the final concentration of internal standard in the sample (Table 1). Detailed application and statistical analysis of the lipidomics data can be found in the statistical analysis section of Zhou et al. (2022), where this protocol was originally described.
Table 1. Lipid classes and their respective internal standard for normalization
Lipid class Internal standard for normalization
SM SM (16:0) (d7)
CE CE(22:6) (d7)
CER CER(16:0) (d9)
DCER DCER(16:0) (d9)
HCER HCER(16:0) (d9)
LCER LCER(16:0) (d9)
TAG TAG(52:1/FA18:0) (d9)
DAG TAG(52:1/FA18:0) (d9)
MAG TAG(52:1/FA18:0) (d9)
LPC LPC(16:0) (d9)
PC PC(16:0/16:1) (d9)
LPE LPE(18:0) (d5)
PE PE(18:0/18:1) (d5)
LPG LPC(16:0) (d9)
PG PC(16:0/16:1) (d9)
LPI LPC(16:0) (d9)
PI PC(16:0/16:1) (d9)
LPS LPC(16:0) (d9)
PS PC(16:0/16:1) (d9)
Myristic acid FFA(17:1)
Palmitic acid FFA(17:1)
Stearic acid FFA(17:1)
Oleic acid FFA(17:1)
Linoleic acid FFA(17:1)
Validation of protocol
As the lipid analysis on amide columns is widely used, several papers have sought to validate lipidomics methods using similar systems (Contrepois et al., 2018; Cao et al., 2019; Loef et al., 2020). For example, Medina et al. (2022) employed an amide column and ammonium acetate buffers in acetonitrile and water for their assay on plasma (Medina et al., 2022). Good linearity (r2 > 0.99) was obtained over broad concentrations, precision was determined by PCA plots of standards, and both intra-day and inter-day repeatability of < 25% were achieved for the majority of the lipids detected in the sample. Another study with similar parameters (Munjoma et al., 2022) obtained reproducibility coefficient of variance (CV) of < 1.0%, intra-day accuracy of ±20% CV of the nominal value, and less than 20% for intra-day precision for most of the lipids (Munjoma et al., 2022).
General notes and troubleshooting
Troubleshooting is a complex process for analytical work using mass spectrometers. Common root causes for issues encountered in LC-MS include:
Clogging of column/in-line filter/transfer tubing;
Leakage in LC instrument;
Stationary phase collapse;
Contamination of ion-source/MS detector by sample molecules;
Signal suppression due to matrix effects.
The recommended corrective actions to address these issues have been outlined on the vendor’s website and other online resources (Steed, 2018; Agilent, 2023; Schug and Watson, 2023). The internal standard mixture or specific lipid standards can be used for troubleshooting purposes for our lipidomics method.
To isolate the underlying problem, it is advisable to follow a step-by-step approach: first, determine whether the issue is caused by LC or MS instrument, and then narrow the issue to specific LC or MS components. Usually, LC-related issues may manifest as alterations in retention time and peak shapes. On the other hand, MS-related issues may affect signal intensity and precision. A good way to distinguish which instrument is faulty is to run either a MS-based method (i.e., direct-injection into the mass spectrometer) or a LC-based method (i.e., using UV-detector) and see whether the issue still persists when one system is bypassed.
If we find that the issue lies with the LC system, it is useful to compare the pressure trace with past records and check whether it is too high or too low. High column backpressure may be indicative of a blockage (i.e., blocked column, plugged inlet-frit, clogged tubing). Low column backpressure is likely due to leakage (i.e., loose fitting, wear and tear of LC components). We can then proceed with the corrective action by starting from the most downstream component and working our way backward. For example, if we suspect there is a blockage, we can:
Clean/replace the column;
Replace the inlet-frit;
Replace tubing;
Clean the autosampler;
Clean the injection loop;
Check for microbial growth in solvent bottles.
Another two key pieces of information are the peak shapes and retention time, which can also be affected by the common root causes that we have listed above. For example, leaks, plugs, and deposits of material in the column may affect flow rate and cause variable shifts in retention time. A partially blocked frit can cause peak splitting or peak tailing because a portion of the sample may take a longer route around the blockage and arrive at the column later than the rest of the sample. Stationary phase collapse often leads to drastic changes in peak shapes and retention time. Readers are encouraged to read up on the following reference to learn how to interpret defective peak shapes (Dolan, 2015).
With regards to the MS system, contaminants constitute a serious problem, because they can potentially reduce signal intensity and increase background noise. For example, detergents such as polyethylene glycol and plasticizers such as phthalate esters are known to interfere with the analytes of interest by causing ion suppression (Rardin, 2018). Therefore, it is important to refrain from using soap or detergent in washing the glassware and minimize the use of plastic containers/vials during LC-MS experimentation. It is also good practice to clean the ion source and front-end of the instrument regularly. Additionally, one should always divert the unretained potion of the gradient at the beginning of each run to the waste.
Another important concern related to MS is matrix effects, whereby the components of the sample matrix may interfere with the ionization of the analytes of interest. One way to ascertain the occurrence of ion suppression is by spiking the lipid standards in pure solvent or sample matrix and comparing their percentage recovery. Matrix effects may be mitigated by diluting the samples, increasing the length of LC separation, or using solid-phase extraction to clean up the samples prior to introducing them into the MS.
Acknowledgments
We thank the editors for recommending the publication of this protocol, which was originally described and validated in the research paper, Zhou et al. (2022).
This work was supported by Ministry of Health, A*STAR and National Medical Research Council Singapore grants MOH-000306 (MOH-CSASI19may-0001) to P.M.Y.; Duke-NUS Medical School and Estate of Tan Sri Khoo Teck Puat Khoo Pilot Award (Collaborative) Duke-NUS-KP (Coll)/2018/0007A to J.Z. Graphic abstract was created with BioRender.com.
We would also like to acknowledge Kee Voon Chua and Qian Ying Ong for assisting in the artwork of the publication.
Competing interests
The authors report no competing interests.
References
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Supplementary information
The following supporting information can be downloaded here:
Table A1
Article Information
Copyright
© 2023 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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4,774 | https://bio-protocol.org/en/bpdetail?id=4774&type=0 | # Bio-Protocol Content
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Peer-reviewed
mRNA Delivery Platform Based on Bacterial Outer Membrane Vesicles for Tumor Vaccine
XG Xiaoyu Gao
YL Yao Li
GN Guangjun Nie
XZ Xiao Zhao
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4774 Views: 1152
Reviewed by: David PaulMiaowei Marwell Mao Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Advanced Materials Mar 2022
Abstract
The rapid display and delivery method for customized tumor mRNA vaccines is limited. Herein, bacteria-derived outer membrane vesicles (OMVs) are employed as an mRNA delivery platform by surface engineering of an RNA-binding protein, L7Ae. OMV-L7Ae can rapidly adsorb boxC/D sequence-labeled mRNA antigens through L7Ae-boxC/D binding and deliver them into HEK-293T and dendritic cells. This platform provides an mRNA delivery technology distinct from lipid nanoparticles (LNPs) for personalized mRNA tumor vaccination and with a Plug-and-Display strategy suitable for rapid preparation of the personalized mRNA tumor vaccine against varied tumor antigens.
Key features
• OMVs are employed as an mRNA delivery platform through L7Ae-boxC/D binding.
Graphical overview
Keywords: RNA binding protein BoxC/D Cancer immunotherapy mRNA vaccines Outer membrane vesicles Rapid display
Background
In recent years, mRNA vaccines have emerged as a promising weapon in cancer immunotherapy (G. Liu et al., 2021). Through precise sequence design, tumor mRNA vaccines encode one or more tumor-specific antigens derived from gene mutations in tumor cells. After vaccine uptake and intracellular protein translation in antigen presenting cells (APCs), tumor-specific antigens form complexes with the major histocompatibility complex I (MHCI) that are presented to T cells to activate a robust antitumor immunity (Miao et al., 2021). However, because of its poor stability, large molecular weight, and high negative charge, mRNA vaccines must rely on efficient delivery carriers to enter APCs (Miao et al., 2019; Barbieri and Kouzarides, 2020). So far, the mainstream mRNA carriers in the clinic are lipid nanoparticles (LNPs) (Liang and Zhao, 2021), which encapsulate mRNA in nanocarriers through a microfluid-based synthesis process (C. Liu et al., 2019; Blass and Ott, 2021; Dolgin, 2021). However, due to the heterogeneity and complexity of tumor antigens, the time-consuming encapsulation process is not suitable to produce personalized tumor vaccines (Wang et al., 2022).
Outer membrane vesicles (OMVs), as biogenic nanocarriers secreted by bacteria, are rich in bacterial components, with the ability to integrate adjuvants and carriers (Cheng et al., 2021; Liang et al., 2022; Yue et al., 2022; Zhao et al., 2022). In addition, OMVs can be obtained by bacterial fermentation in large quantities, which has attracted more and more attention (Cheng et al., 2020; Gao et al., 2022). Here, we describe a Plug-and-Display strategy for mRNA antigen delivery in an OMV-based platform by arming OMVs with an RNA-binding protein, L7Ae (Li et al., 2022). BoxC/D, the matched binding sequence, is added to the 3′-untranslated region (UTR) of the in vitro–transcribed (IVT) mRNA. The archaeal RNA-binding protein L7Ae is fused to the C-terminal of the surface protein ClyA on the OMVs (OMV-L7Ae). The boxC/D sequence adopts the standard k-turn conformation that is specifically recognized by the L7Ae protein, which stabilizes the stem-loop and forms a standard L7Ae-k-turn complex (Moore et al., 2004; Turner et al., 2005; Saito et al., 2010). Through the strong and specific binding between the L7Ae protein and the boxC/D sequence, the boxC/D-labeled mRNA (boxC/D-mRNA) is rapidly adsorbed onto the surface of the OMV-L7Ae, achieving the successful mRNA delivery into cells via endocytosis of OMVs by HEK-293T or bone marrow dendritic cells (BMDCs). This Plug-and-Display strategy is suitable for the rapid preparation of tumor mRNA vaccines against heterogeneous and complex tumor antigens.
Materials and reagents
Biological materials
E. coli [strain BL21 (DE3)] (Tiangeng, catalog number: DHGST-21)
HEK-293T cells (American Type Culture Collection, catalog number: CRL-3216)
C57BL/6 mice (Vital River Laboratory Animal Technology, catalog number: 01059)
Reagents
NaCl (Solarbio, catalog number: S8210)
Yeast extract (Solarbio, catalog number: Y8020)
Tryptone (Solarbio, catalog number: T8490)
Agar (Solarbio, catalog number: A8190)
Chloramphenicol (50 mg/mL) (Solarbio, catalog number: L1311)
Isopropyl-β-d-thiogalactoside (IPTG) (Solarbio, catalog number: BS119)
Note: IPTG causes serious eye irritation. Wear nitrile gloves, safety goggles, and lab coats, and operate carefully in a fume hood.
Fetal bovine serum (FBS) (Wisent, catalog number: 085-150)
Penicillin G and streptomycin (Wisent, catalog number: 450-201-EL)
Phosphate-buffered saline (PBS) (Wisent, catalog number: 311-010-CL)
Dulbecco’s modified Eagle medium (DMEM) (Wisent, catalog number: 319-005-CL)
RPMI 1640 medium (Wisent, catalog number: 350-000-CL)
HiScribe T7 ARCA mRNA kit (with tailing) (New England Biolabs, catalog number: E2060S)
Q5 high-fidelity 2× master mix (New England Biolabs, catalog number: M0492S)
Tris-HCl (1 M, pH 8.0) (Solarbio, catalog number: T1150)
MgCl2 (1 M) (Beyotime, catalog number: ST269)
Diethylpyrocarbonate (DEPC)-treated water (Biosharp, BL510B)
Citric acid sodium citrate buffer (0.1 M, pH = 4.0) (Leagene, R00521)
Ethanol (Macklin, catalog number: E809063)
Ammonium chloride potassium (ACK) lysis buffer (Solarbio, catalog number: R1010)
HEPES (1 M, pH 7.4) (Sigma-Aldrich, catalog number: 83264)
β-Mercaptoethanol (β-ME) (Gibco, catalog number: 21985023)
IL-4 (Sino Biological, catalog number: 51084-MNAE)
Granulocyte-macrophage colony-stimulating factor (GM-CSF) (Sino Biological, catalog number: 51048-M01H)
FITC-anti-mouse CD11c (BioLegend, catalog number: 117306)
APC-anti-mouse CD80 (BioLegend, catalog number: 104713)
PE/Cy7-anti-mouse CD86 (BioLegend, catalog number: 105014)
APC-anti-mouse H-2Kb bound to SIINFEKL (BioLegend, catalog number: 141605)
pST1374-NLS-flag-linker-Cas9 plasmid (Addgene, catalog number: 44758)
Trypsin-EDTA (0.25%) (Meilunbio, catalog number: MB4376)
Solutions
Solid LB medium (see Recipes)
Liquid LB medium (see Recipes)
Reaction buffer for mRNA binding (pH = 5.0) (see Recipes)
Recipes
Solid LB medium
Reagent Final concentration Amount
Yeast extract 5 g/L n/a
Tryptone 10 g/L n/a
NaCl 10 g/L n/a
Agar 15 g/L n/a
H2O n/a 100 mL
Total n/a 100 mL
Prepare the solid LB medium by dissolving 1.5 g of agar, 1 g of NaCl, 1 g of tryptone, and 0.5 g of yeast extract in 100 mL of deionized water. After sterilization for 20 min at 121 °C and 0.1 MPa, add 100 μL of 50 mg/mL chloramphenicol when the temperature reaches approximately 60 °C. Add 10 mL of sterilized LB medium to a dish (10 cm) after fully mixing. After solidification, seal the solid LB medium with sealing film. Store the solid LB medium for one month at 4 °C.
Note: Add antibiotics only after the temperature drops to ~50–60 °C, because their activity is seriously affected by temperature.
Liquid LB medium
Reagent Final concentration Amount
Yeast extract 5 g/L n/a
Tryptone 10 g/L n/a
NaCl 10 g/L n/a
H2O n/a 1 L
Total n/a 1 L
Prepare the liquid LB medium by dissolving 10 g of tryptone, 5 g of yeast extract, and 10 g of NaCl in 1 L of deionized water. After sterilization for 20 min at 121 °C and 0.1 MPa, store the liquid LB medium for one month at 4 °C.
Reaction buffer for mRNA binding (pH = 5.0)
Reagent Final concentration Amount
NaCl 100 × 10-3 M n/a
Tris-HCl (1 M, pH 8.0) 5 × 10-3 M 200 μL
MgCl2 (1 M) 10 × 10-3 M 400 μL
Total n/a 40 mL
Prepare the reaction buffer for mRNA binding by dissolving 233.76 mg of NaCl in 30 mL of DEPC-treated water. Then, add 200 μL of 1 M Tris-HCl (pH = 8.0) and 400 μL of 1 M MgCl2 and fully mix. After adjusting pH to 5.0 with citric acid sodium citrate buffer, set the volume to 40 mL. Seal the reaction buffer for mRNA binding with sealing film and store the reaction buffer for mRNA binding for one month at 4 °C.
Note: To prevent the introduction of RNA enzymes, water must be DEPC treated.
Laboratory supplies
Conical flask (250 mL) (Heqi Glass, catalog number: B-000207)
Conical flask (500 mL) (Heqi Glass, catalog number: B-000209)
Sterile breathable sealing film (Bkmam, catalog number: B-FK14-50E)
Culture plate (10 cm) (Corning, catalog number: 430167)
Centrifuge tube (50 mL) (Corning, catalog number: 430829)
Centrifuge tube (15 mL) (Corning, catalog number: 430791)
Centrifuge tube (1.5 mL) (Kirgen, catalog number: KG2211)
PCR tube (0.2 mL) (Thermo Scientific, catalog number: AB0620)
6-well plate (Corning, catalog number: 3516)
24-well plate (Corning, catalog number: 3524)
Sealing film (Parafilm, catalog number: PM-996)
Injection syringe (Acmec, catalog number: AYA0553)
Filter (0.45 μm) (Merck Millipore, catalog number: SLHPR33RB)
Filter (0.22 μm) (Merck Millipore, catalog number: SLGPR33RB)
Cell strainer (70 μm) (BD Falcon, catalog number: 352350)
Ultrafiltration tube (100 kDa) (Merck Millipore, catalog number: UFC910096)
Ultracentrifugation tube (Beckman, catalog number: 355618)
Quartz cell (Heqi Glass, catalog number: B-037003)
Equipment
Water bath (BluePard, model: HWS-24)
Bacteriological incubator (BluePard, model: THZ-98C)
Clean bench (for bacteria) (Beijing Dinglian Har Instrument Manufacture, model: DL-CJ-1NDII)
Clean bench (for cell) (Thermo Fisher Scientific, catalog number: 1384)
High-speed centrifuge (Thermo Fisher Scientific, model: Sorvall ST 8R)
Ultra-speed centrifuge (Beckman, model: OPTIMA XPN-100)
Flow cytometer (Agilent Technologies, catalog number: 462171219542)
Spectrophotometer (Asone, model: ASV-S3)
Cell incubator (Thermo Fisher Scientific, catalog number: BB150)
pH meter (Mettler Toledo, catalog number: S210FE20K)
Refrigerator (-80 °C) (Haier, model: HYCD-290)
Refrigerator (4 °C and -20 °C) (Haier, model: DE-25W262)
T100 thermal cycler (Bio-Rad, catalog number: 621BR55553)
NanoDrop (Implen, model: N50 Touch)
Automated cell counter (Invitrogen, model: Countess 3)
Autoclave (STIK Instrument Equipment, model: IMJ-54A)
Software
Prism 9 (GraphPad)
Procedure
Plasmid design and construction (timing: 1 day)
Design and construction of pACYC-ClyA-L7AE plasmid:
The following sequences are synthesized, sequentially ligated, and inserted into the pACYCDuet-1 plasmid by NdeI (CATATG) and XhoI (CTCGAG) (Table 1). The product is named pACYC-ClyA-L7AE.
Table 1. Design of pACYC-ClyA-L7AE plasmid
DNAs Sequences
ClyA ATGACTGAAATCGTTGCAGATAAAACGGTAGAAGTAGTTAAAAACGCAATCGAAACCGCAGATGGAGCATTAGATCTTTATAATAAATATCTCGATCAGGTCATCCCCTGGCAGACCTTTGATGAAACCATAAAAGAGTTAAGTCGCTTTAAACAGGAGTATTCACAGGCAGCCTCCGTTTTAGTCGGCGATATTAAAACCTTACTTATGGATAGCCAGGATAAGTATTTTGAAGCAACCCAAACAGTGTATGAATGGTGTGGTGTTGCGACGCAATTGCTCGCAGCGTATATTTTGCTATTTGATGAGTACAATGAGAAGAAAGCATCCGCCCAGAAAGACATTCTCATTAAGGTACTGGATGACGGCATCACGAAGCTGAATGAAGCGCAAAAATCCCTGCTGGTAAGCTCACAAAGTTTCAACACGCTTCCGGGAAACTGCTGGCGTTAGATAGCCAGTTAACCAATGATTTTTCAGAAAAAAGCAGCTATTTCCAGTCACAGGTAGATAAAATCAGGAAGGAAGCGTATGCCGGTGCCGCAGCCGGTGTCGTCGCCGGTCCATTTGGATTAATCATTTCCTATTCTATTGCTGCGGGCGTAGTTGAAGGAAAACTGATTCCAGAATTGAAGAACAAGTTAAAATCTGTGCAGAATTTCTTTACCACCCTGTCTAACACGGTTAAACAAGCGAATAAAGATATCGATGCCGCCAAATTGAAATTAACCACCGAAATAGCCGCCATCGGTGAGATAAAAACGGAAACTGAAACAACCAGATTCTACGTTGATTATGATGATTTAATGCTTTCTTTGCTAAAAGAAGCGGCCAAAAAAATGATTAACACCTGTAATGAGTATCAGAAAAGACACGGTAAAAAGACACTCTTTGAGGTACCTGAAGTC
Linker GGTGGTGGATCA
L7Ae ATGTACGTGAGATTTGAGGTTCCTGAGGACATGCAGAACGAAGCTCTGAGTCTGCTGGAGAAGGTTAGGGAGAGCGGTAAGGTAAAGAAAGGTACCAACGAGACGACAAAGGCTGTGGAGAGGGGACTGGCAAAGCTCGTTTACATCGCAGAGGATGTTGACCCGCCTGAGATCGTTGCTCATCTGCCCCTCCTCTGCGAGGAGAAGAATGTGCCGTACATTTACGTTAAAAGCAAGAACGACCTTGGAAGGGCTGTGGGCATTGAGGTGCCATGCGCTTCGGCAGCGATAATCAACGAGGGAGAGCTGAGAAAGGAGCTTGGAAGCCTTGTGGAGAAGATTAAAGGCCTTCAGAAG
Linker GGTGGCGGATCA
3× HA tag TACCCATACGATGTTCCAGATTACGCTTATCCCTACGACGTGCCTGATTATGCATACCCATACGATGTCCCCGACTATGCC
Termination codon TAA
Notes:
The pACYCDuet-1 plasmid is only a suggested plasmid, which can be replaced by other suitable plasmids.
The principle of plasmid selection: (1) choose a small-molecular-weight plasmid, so that the plasmid is more stable and with a higher copy number; (2) choose a relaxation control plasmid, so that the plasmid can replicate autonomously; (3) choose a plasmid with multiple restriction enzyme cut points; (4) choose a plasmid with easy-to-detect markers, such as resistance markers; and (5) choose a plasmid expressed in E. coli.
Design and construction of pST1374-EGFP-boxC/D plasmid:
The following sequences are synthesized, sequentially ligated, and inserted into pST1374-NLS-flag-linker-Cas9 plasmid by NdeI (CATATG) and ApaI (GGGCCC) (Table 2). The product is named pST1374-EGFP-boxC/D.
Table 2. Design of pST1374-EGFP-boxC/D plasmid
DNAs Sequences
5′-UTR GGGAAATAAGAGAGAAAAGAAGAGTAAGAAGAAATATAAGAGCCACC
EGFP ATGGTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCCATCCTGGTCGAGCTGGACGGCGACGTAAACGGCCACAAGTTCAGCGTGTCCGGCGAGGGCGAGGGCGATGCCACCTACGGCAAGCTGACCCTGAAGTTCATCTGCACCACCGGCAAGCTGCCCGTGCCCTGGCCCACCCTCGTGACCACCCTGACCTACGGCGTGCAGTGCTTCAGCCGCTACCCCGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCCGAAGGCTACGTCCAGGAGCGCACCATCTTCTTCAAGGACGACGGCAACTACAAGACCCGCGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACCGCATCGAGCTGAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACAACAGCCACAACGTCTATATCATGGCCGACAAGCAGAAGAACGGCATCAAGGTGAACTTCAAGATCCGCCACAACATCGAGGACGGCAGCGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCCATCGGCGACGGCCCCGTGCTGCTGCCCGACAACCACTACCTGAGCACCCAGTCCGCCCTGAGCAAAGACCCCAACGAGAAGCGCGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAGTAA
3′-UTR TGATAATAGGCTGGAGCCTCGGTGGCCATGCTTCTTGCCCCTTGGGCCTCCCCCCAGCCCCTCCTCCCCTTCCTGCACCCGTACCCCCGTGGTCTTTGAATAAAGTCTGA
BoxC/D GGGCGTGATGCGAAAGCTGACCCGGGCGTGATGCGAAAGCTGACCCGCTCTGACCGAAAGGCGTGATGAGCGCTCTGACCGAAAGGCGTGATGAGC
Notes:
The pST1374-NLS-flag-linker-Cas9 plasmid is only a suggested plasmid, which can be replaced by other suitable plasmids.
The principle of plasmid selection: (1) choose a small-molecular-weight plasmid, so that the plasmid is more stable and with a higher copy number; (2) choose a relaxation control plasmid, so that the plasmid can replicate autonomously; (3) choose a plasmid with multiple restriction enzyme cut points; (4) choose a plasmid with easy-to-detect markers, such as resistance markers; and (5) choose a plasmid expressed in E. coli.
Design and construction of pST1374-sec-OVA-3HA-MITD- boxC/D plasmid:
The following sequences are synthesized, sequentially ligated, and inserted into pST1374-NLS-flag-linker-Cas9 plasmid by NdeI (CATATG) and ApaI (GGGCCC) (Table 3). The product is named pST1374-sec-OVA-3HA-MITD-boxC/D.
Table 3. Design of pST1374-sec-OVA-3HA-MITD-boxC/D plasmid
DNAs Sequences
5′-UTR GGGAAATAAGAGAGAAAAGAAGAGTAAGAAGAAATATAAGAGCCACC
Sec ATGGTACCGTGCACGCTGCTCCTGCTGTTGGCGGCCGCCCTGGCTCCGACTCAGACCCGCGCG
Linker GGCGGTTCTGGAGGGGGTGGGTCCGGGGGT
OVA CAGCTTGAGAGTATAATCAACTTTGAAAAACTGACT
3 HA TACCCATACGATGTTCCAGATTACGCTTATCCCTACGACGTGCCTGATTATGCATACCCATATGATGTCCCCGACTATGCC
MITD GCGACCGTTGCTGTTCTGGTTGTCCTTGGAGCTGCAATAGTCACTGGAGCTGTGGTGGCTTTTGTGATGAAGATGAGAAGGAGAAACACAGGTGGAAAAGGAGGGGACTATGCTCTGGCTCCAGGCTCCCAGACCTCTGATCTGTCTCTCCCAGATTGTAAAGTGATGGTTCATGACCCTCATTCTCTAGCGTGA
3′-UTR GCTCGCTTTCTTGCTGTCCAATTTCTATTAAAGGTTCCTTTGTTCCCTAAGTCCAACTACTAAACTGGGGGATATTATGAAGGGCCTTGAGCATCTGGATTCTGCCTAATAAAAAACATTTATTTTCATTGCGCTCGCTTTCTTGCTGTCCAATTTCTATTAAAGGTTCCTTTGTTCCCTAAGTCCAACTACTAAACTGGGGGATATTATGAAGGGCCTTGAGCATCTGGATTCTGCCTAATAAAAAACATTTATTTTCATTGC
BoxC/D GGGCGTGATGCGAAAGCTGACCCGGGCGTGATGCGAAAGCTGACCCGCTCTGACCGAAAGGCGTGATGAGCGCTCTGACCGAAAGGCGTGATGAGC
Notes:
The pST1374-NLS-flag-linker-Cas9 plasmid is only a suggested plasmid, which can be replaced by other suitable plasmids.
The principle of plasmid selection: (1) choose a small-molecular-weight plasmid, so that the plasmid is more stable and with a higher copy number; (2) choose a relaxation control plasmid, so that the plasmid can replicate autonomously; (3) choose a plasmid with multiple restriction enzyme cut points; (4) choose a plasmid with easy-to-detect markers, such as resistance markers; and (5) choose a plasmid expressed in E. coli.
The above plasmids are synthesized by GENEWIZ, Suzhou, China (https://www.genewiz.com.cn/).
Plasmid transformation and bacterial culture (timing: 3 days)
Place a centrifuge tube (1.5 mL) containing 100 μL of competent cells [E. coli BL21 (DE3)] in an ice bath until completely thawed.
Notes:
Store the competent cells [E. coli BL21 (DE3)] at -80 °C.
Avoid freezing and thawing to ensure the activity of the competent cells.
After thawing the competent cells, DNA should be quickly added.
Add 1 ng of pACYC-ClyA-L7AE plasmid into the competent cells. Mix gently and let stand in an ice bath for 30 min.
Place the centrifuge tube for 60–90 s in a 42 °C water bath. Transfer the centrifuge tube quickly to an ice bath for 2–3 min to cool the cells.
Note: Do not shake the centrifugal tube during this process.
Add 900 μL of liquid LB medium to the centrifuge tube. Incubate for 45–60 min at 37 °C in a shaker at 150 rpm.
Notes:
The resistance gene has not yet been expressed during this process, so the liquid LB medium must be antibiotic-free.
Growth for 1 h at 37 °C has the best effect on cell recovery and antibiotic resistance expression.
Add 100 μL of transformed cells to a culture plate (10 cm) with solid LB medium containing 50 μg/mL chloramphenicol. Spread the cells gently with a sterile glass stick.
Note: Prepare the culture plate (10 cm) with the solid LB medium containing 50 μg/mL chloramphenicol in advance.
Place the culture plate at room temperature until the liquid is absorbed. Invert the culture plate and culture for 12–16 h at 37 °C. The culture plate should be sealed with sterile sealing film (Troubleshooting 1).
Note: The number of bacteria can be adjusted on the plate, ideally to obtain several dozens of colonies on a plate (10 cm).
Pause point: Store the bacteria on the plate for one month at 4 °C.
Carefully pick one bacterial colony on the plate. Add it into 3 mL of liquid LB medium containing 50 μg/mL chloramphenicol in a centrifuge tube (15 mL). Incubate for 10–12 h at 37 °C with shaking at 180 rpm.
Note: To ensure oxygen supply for bacterial growth, seal the centrifuge tube using the sterile breathable sealing film.
Add 3 mL of bacterial culture medium from step B7 into 300 mL of liquid LB medium containing 50 μg/mL chloramphenicol in a conical flask (500 mL). Incubate at 37 °C with shaking at 180 rpm. A spectrophotometer is used to monitor the optical density at 600 nm (OD600) of the bacterial culture medium. Add 0.1 mM IPTG to the medium when the OD600 reaches 0.6. Incubate for another 16 h at 16 °C with shaking at 180 rpm.
Note: The IPTG-induced expression can be performed for 2 h at 37 °C, but the expression efficiency is not as good as induction at 16 °C.
OMV-L7Ae extraction (timing: 10 h)
Divide the 300 mL bacterial medium into six centrifuge tubes (50 mL). After centrifugation at 7,000× g for 15 min at 4 °C, collect the supernatant.
Notes:
Balance the weight of the centrifugal tubes (50 mL).
Dispose as soon as possible after centrifugation to prevent bacteria from spreading into the supernatant.
Filter the 300 mL supernatant through a filter (0.45 μm). Through centrifugation at 3,000× g for 5–10 min at 4 °C, concentrate the filtering medium to 100 mL using an ultrafiltration tube (100 kDa).
Filter the concentrated solution through a filter (0.22 μm).
Put the filtering medium into two ultracentrifugation tubes.
Note: Clean and dry the ultracentrifugation tubes in advance.
Separate the OMV-L7Ae through ultracentrifugation at 150,000× g for 3 h at 4 °C.
Notes:
Balance the weight of the ultracentrifugation tubes; the weight error of all ultracentrifugation tubes is less than 50 mg.
Tighten the ultracentrifugation tube cover.
Discard the supernatant. To fill up the ultracentrifugation tubes, resuspend the deposited OMV-L7Ae at the bottom of the tubes using DEPC-treated PBS. Repeat the ultracentrifugation at 150,000× g for 3 h at 4 °C.
Discard the supernatant. Resuspend the deposited OMV-L7Ae with the 200 μL of reaction buffer for mRNA binding in each tube (Troubleshooting 2).
Note: After ultracentrifugation, discard the supernatant as soon as possible to prevent the OMV-L7Ae from redissolving in the supernatant.
Pause point: Store the OMV-L7Ae for one week at -80 °C.
BoxC/D-mRNA synthesis (timing: 1 day)
Notes:
The following method is a reference to protocols from NEB’s official website (https://www.neb.com/).
Before the experiment, the table should be wiped with DEPC-water to avoid RNase contamination. Please wear gloves during the entire process. Be sure to use nuclease-free tubes and reagents to avoid RNase contamination.
Preparation of template DNA:
Design of primers (Table 4).
Table 4. Design of primers
DNAs Sequences
Forward primer CTGGCTAACTAGAGAACCCAC
Reverse primer CTAGAAGGCACAGTCGAGGCTG
Put the following PCR reaction master mix into a PCR tube (0.2 mL) (Table 5).
Table 5. PCR reaction master mix
Reagent Amount Final Concentration
Q5 high-fidelity 2× master mix 25 μL 1×
10 μM forward primer 2.5 μL 0.5 μM
10 μM reverse primer 2.5 μL 0.5 μM
Plasmid (pST1374-EGFP-boxC/D)/plasmid (pST1374-sec-OVA-3HA-MITD-boxC/D) variable 0.4 ng/μL
Nuclease-free water to 50 μL /
Notes:
i. The Q5 high-fidelity 2× master mix should be thawed on ice to prevent inactivation.
ii. All the reaction components are operated on ice, and all components should be mixed prior to use.
Collect all liquid to the tube bottom by a quick spin and quickly transfer the reactions to a thermal cycler preheated to the denaturation temperature (98 °C).
Begin PCR thermocycling (Table 6).
Table 6. PCR cycling conditions
Steps Temperature Time Cycles
Initial denaturation 98 °C 30 s 1
Denaturation 98 °C 7 s
Annealing 60 °C 20 s 35 cycles
Extension 72 °C 25 s
Final extension 72 °C 2 min 1
Hold 4 °C Forever
Determine the concentration of PCR product using NanoDrop. Generally, the concentration is greater than 600 ng/μL (Troubleshooting 3).
Pause point: Store the PCR product for one month at -20 or -80 °C.
IVT synthesis of boxC/D-mRNA:
Prepare IVT mix into a PCR tube (0.2 mL) (Table 7).
Table 7. IVT mix
Reagent Amount Final Concentration
Template DNA (PCR product from step D1) variable 0.05 μg/μL
2× ARCA/NTP mix 10 μL 1×
T7 RNA polymerase mix 2 μL /
Nuclease-free water to 20 μL /
Notes:
i. All components should be thawed on ice to prevent inactivation.
ii. All the reaction components are operated on ice, and all components should be mixed prior to use.
Collect all liquid to the tube bottom with a quick spin. Incubate for 30 min at 37 °C.
Note: A longer reaction time would help to produce more IVT product.
Pause point: After the IVT reaction, store the product for one week at -20 °C.
Add 2 μL of DNase I to the IVT product, mix well, and incubate for 15 min at 37 °C to remove template DNA.
Prepare the following poly(A) tailing mix into a PCR tube (0.2 mL) (Table 8).
Table 8. Poly(A) tailing mix
Reagent Amount
IVT product from step D2c 20 μL
10× Poly(A) polymerase reaction buffer 10 μL
Poly(A) polymerase 5 μL
Nuclease-free water to 100 μL
Note: The unpurified IVT product contains enough ATP; no extra ATP is necessary for the poly(A) tailing reaction.
Collect all liquid to the tube bottom with a quick spin. Incubate for 30 min at 37 °C.
BoxC/D-mRNA purification:
To the 50 μL poly(A) tailing product, add 25 μL of LiCl solution and mix well.
Incubate for 30 min at -20 °C.
Centrifuge at 20,000× g for 15 min at 4 °C to pellet the boxC/D-mRNA.
Remove the supernatant carefully. Rinse the pellet by adding 500 μL of cold 70% ethanol.
Note: 70% ethanol should be pre-cooled to -20 °C in advance.
Centrifuge at 20,000× g for 10 min at 4 °C.
Remove the ethanol carefully. Spin the tube briefly to bring down any liquid on the wall. Remove residual liquid carefully using a sharp tip (e.g., loading tip).
Note: The residual liquid should be removed to prevent organic solvents from affecting subsequent experiments.
Air dry the pellet and resuspend the boxC/D-mRNA in 50 μL of DEPC-treated water.
Heat the boxC/D-mRNA for 5 min at 65 °C to completely dissolve the boxC/D-mRNA. Mix well.
Determine the boxC/D-mRNA concentration using NanoDrop. Generally, the boxC/D-mRNA concentration is approximately 500 ng/μL. The mRNA from plasmid pST1374-EGFP-boxC/D is named EGFP-boxC/D mRNA and the mRNA from plasmid pST1374-sec-OVA-3HA-MITD-boxC/D is named OVA-boxC/D mRNA (Troubleshooting 4).
Note: The 260/280 of mRNA is between 1.8 and 2.0; 260/230 ≥ 2 indicates that the prepared mRNA is qualified.
Pause point: Store the boxC/D-mRNA for one month at -20 or -80 °C.
Assembly and delivery of boxC/D-mRNA by OMV-L7Ae (timing: 11 days)
Assembly of boxC/D-mRNA and OMV-L7Ae:
Mix 9 μg of OMV-L7Ae (total protein weight) from step C7 and 1 μg of boxC/D-mRNA (EGFP-boxC/D or OVA-boxC/D) from step D3i and then put the mixture (OMV-L-mRNA) for 5 min at room temperature.
Notes:
OMV-L7Ae and boxC/D-mRNA should be mixed immediately before delivery.
OMV-L7Ae vesicles should be free of bacterial contamination.
Delivery and evaluation of EGFP-boxC/D mRNA by OMV-L7Ae:
Before transfection experiments of 18–24 h, add HEK-293T cells to a 24-well plate at a density of 40,000 cells per well with 500 μL of DMEM medium containing 10% FBS, 100 U/mL penicillin G, and 100 μg/mL streptomycin. The cells are grown at 37 °C in the cell incubator with 5% CO2.
Supplement the OMV-L-mRNA mixture (EGFP-boxC/D mRNA) from step E1 with 500 μL of DMEM medium.
Carefully discard the cell supernatant of the 24-well plate.
Note: The cell density is approximately 80% when transfection experiments are performed.
Add 500 μL of OMV-L-mRNA mixture from step E2b to the HEK-293T cells in the 24-well plate.
Culture cells for 6–8 h at 37 °C in a humidified atmosphere with 5% CO2.
Carefully discard the cell supernatant of 24-well plate. Add 500 μL of DMEM medium containing 10% FBS, 100 U/mL penicillin G, and 100 μg/mL streptomycin to the 24-well plate.
At 24 h after transfection, digest the cells with 100 μL of trypsin-EDTA (0.25%) for 30 s. Add 300 μL of RPMI 1640 medium containing 10% FBS, 100 U/mL penicillin G, and 100 μg/mL streptomycin to stop digestion.
Through centrifugation at 800× g for 5 min at 4 °C, collect the HEK-293T cells.
Discard the supernatant and then resuspend the HEK-293T cells with 200 μL of PBS. Perform the flow cytometry evaluation within 1 h and analyze the cells expressing EGFP (Troubleshooting 5).
Note: HEK-293T cells are fragile and should be handled gently.
Immune stimulation evaluation of OMV-based nanovaccines in vitro through delivery of OVA-boxC/D mRNA by OMV-L7Ae:
Keep the C57BL/6 mice (6–8 weeks old) with a 12 h light/dark cycle and a humidity of 30%–70% at 20–22 °C in a room. Provide food and water ad libitum.
After killing the C57BL/6 mice by cervical dislocation, dissect the mice and obtain the femurs and tibias. Use scissors and tweezers to remove as much muscle tissue as possible around the bones.
Notes:
i. The femur has more bone marrow than the tibia.
ii. Do not destroy the bones to avoid pollution.
Soak the bones in a sterile culture plate containing 70% alcohol for 3 min to disinfect and sterilize and then wash twice with sterile PBS.
Flush repeatedly the bone marrow cells with an injection syringe containing RPMI 1640 medium, 100 U/mL penicillin G, 100 μg/mL streptomycin, and 2% FBS until the bones are completely white.
Note: The femurs and tibias of one mouse are cultured using approximately 40 mL of medium.
Through centrifugation at 800× g for 5 min at 4 °C, collect the bone marrow cells.
Discard the supernatant. To lysis the red blood cells, resuspend the cells with 1 mL of ACK lysis buffer. Incubate for 90 s at room temperature.
Stop the lysis. Add 3 mL of PBS and then filter the cells through a cell strainer (70 μm).
Through centrifugation at 800× g for 5 min at 4 , collect the cells.
Discard the supernatant. Resuspend the precipitated cells with 12 mL of RPMI 1640 medium supplemented with 10% FBS, 100 U/mL penicillin G, 100 μg/mL streptomycin, 1% HEPES, 0.05 mM β-ME, 20 ng/mL IL-4, and 20 ng/mL GM-CSF, and then divide into six wells in a 6-well plate.
Notes:
i. During steps E3a–E3i, make sure all reagents and samples are placed on an ice bath, as this has a positive effect on cell activity.
ii. To maintain a sterile state, perform all steps on a clean bench using sterile containers.
Culture the cells for six days at 37 °C in the cell incubator with 5% CO2.
Notes:
i. Replace half of the medium every 2–3 days.
ii. Remove and add medium gently to avoid interfering with cell growth.
Collect non-adherent cells on day 6 through centrifugation at 800× g for 5 min at 4 , which are known as BMDCs.
Discard the supernatant and then resuspend the precipitated BMDCs with 10 mL of RPMI 1640 medium.
Quantify the cell density using the automated cell counter. Place 100,000 BMDCs into a centrifuge tube (1.5 mL) with 500 μL of RPMI 1640 medium.
Add OMV-L-mRNA mixture (OVA-boxC/D mRNA) from step E1 to the BMDCs in the centrifuge tube (1.5 mL).
Culture cells for 6–8 h at 37 °C in a humidified atmosphere with 5% CO2.
Centrifuge at 800× g for 5 min at 4 °C.
Carefully discard the cell supernatant. Add 500 μL of RPMI 1640 medium containing 10% FBS, 100 U/mL penicillin G, and 100 μg/mL streptomycin to the centrifuge tube (1.5 mL).
At 12 or 24 h after transfection, collect the stimulated BMDCs through centrifugation at 800× g for 5 min at 4 .
Note: Seal the centrifuge tube (1.5 mL) with sealing film but preserve the air hole to ensure oxygen supply.
Discard the supernatant and then resuspend the precipitated BMDCs with 200 μL of RPMI 1640 medium containing 2% FBS.
Add the proper antibodies to analyze the maturation and cross-presentation of BMDCs. For the maturation assay, stain the BMDCs with FITC-anti-mouse CD11c (1:200, 1 μL), APC-anti-mouse CD80 (1:200, 1 μL), or PE/Cy7-anti-mouse CD86 (1:200, 1 μL). For the cross-presentation assay, stain the BMDCs with FITC-anti-mouse CD11c (1:200, 1 μL) and APC-anti-mouse H-2Kb bound to SIINFEKL (MHCI-OVA) (1:80, 2.5 μL).
Stain the BMDCs under dark conditions for 30 min at 4 °C and then add 500 μL of RPMI 1640 medium containing 2% FBS, 100 U/mL penicillin G, and 100 μg/mL streptomycin to stop staining and wash cells.
Through centrifugation at 800× g for 5 min at 4 , collect the stained BMDCs. Discard the supernatant and then resuspend cells with 200 μL of PBS.
Perform the flow cytometry evaluation according to the manufacturer’s protocols within 1 h.
Note: Using the CD80 or CD86 as the maturation marker, the CD80+ or CD86+ cells are gated in the CD11c+ cells. For the cross-presentation assay, the CD11c+MHCI-OVA+ cells are gated in the BMDCs.
Expected outcomes
The flow cytometry results show that the OMV-L-mRNA (EGFP-boxC/D) successfully delivered the EGFP-boxC/D mRNA into HEK-293T cells, resulting in EGFP expression in 29.4% of cells (Figure 1, Supplementary Figure 1).
In the immune stimulation evaluation in BMDCs in step E3, the OMV-L-mRNA (OVA-boxC/D) induced notable maturation and antigen presentation, indicated by the upregulation of the surface expression of CD80, CD86, and MHCI-OVA (Figure 2A–2C, Supplementary Figures 2–3). These data demonstrated that the OMV-based nanocarriers can efficiently deliver mRNA antigens into BMDCs and induce antigen presentation.
Figure 1. Flow cytometry analysis of HEK-293T cells incubated with PBS or outer membrane vesicle (OMV)-L-mRNA (2 μg/mL EGFP-boxC/D mRNA) for 24 h (n = 3). Data are shown as mean ± SD. One-way ANOVA and a Tukey’s multiple comparisons test were used for statistical analysis. ****, p < 0.0001. Figure adapted with permission from Li et al. (2022).
Figure 2. Innate immunity activation and antigen presentation mediated by the outer membrane vesicles (OMV)-based mRNA nanovaccines in vitro. (A, B) Bone marrow dendritic cells (BMDCs) maturation induced by PBS or OMV-L-mRNA (2 μg/mL OVA-boxC/D mRNA) for 12 or 24 h. The expression of CD80 (A) or CD86 (B) in CD11c+ BMDCs was examined using flow cytometry (n = 6). (C) Expression of the MHCI-OVA complex in BMDCs, as measured using flow cytometry (n = 5). The BMDCs were treated with PBS or OMV-L-mRNA (2 μg/mL OVA-boxC/D mRNA) for 24 h. The CD11c+MHCI-OVA+ cells in BMDCs were examined using flow cytometry. Data are shown as mean ± SD. One-way ANOVA and a Tukey’s multiple comparisons test were used for statistical analysis. ****, p < 0.0001. Figure adapted with permission from Li et al. (2022).
Limitations
This study preliminarily demonstrated the feasibility and effectiveness of OMVs as tumor vaccine carriers for mRNA delivery. However, compared with the LNPs-based delivery platform, the efficiency of the OMV-based delivery platform is lower, which needs to be solved urgently in the subsequent clinical transformation process. We can make efforts in the following two aspects: 1) optimize the structure of the archaeal RNA-binding protein L7Ae and the matched binding sequence boxC/D, and 2) more boxC/D sequences can be connected in series to ensure greater binding and delivery efficiency.
In addition, as the bacteria-derived nanobiomaterials, the sterile production of the OMVs requires special attention in future clinical applications. In the procedure for the OMV-based nanocarriers, the bacteria were first centrifuged at 7,000× g for 15 min and almost all the bacteria were precipitated and removed. The OMVs from the supernatant were then filtered twice in 0.45 and 0.22 μm filters to ensure that the final OMVs did not contain bacteria. However, in future clinical applications, radiation sterilization of the final nanocarriers system can be performed to further ensure that the OMV nanocarriers are sterile.
General notes and troubleshooting
Troubleshooting
Problem 1
There are too many or very few bacterial colonies on the plate.
Potential solution
This is one of the most common reasons for failure of the protocol. In many cases, the plasmid resistance genes were not associated with antibiotic, so the plasmid could not be transformed. We need to choose the right antibiotic based on the plasmid resistance genes in the backbone plasmid.
During competent cell resuscitation, the resistance genes were not expressed, and the LB medium with antibiotics was used (step B4), resulting in the failure of plasmid transformation. Therefore, the LB medium must be antibiotic-free in this step.
In the reagent setup of solid LB medium, the appropriate concentration of antibiotics was selected. If the concentration of antibiotics is too high, the target cells will not grow; if the concentration of antibiotics is too low, the growth of other bacteria will not be inhibited, resulting in the failure of plasmid transformation. In step B5, the volume of cells can be adjusted. When the volume of cells is too high, the cells will be too dense on the plate; when the volume of cells is too low, the target cells will not grow, resulting in the failure of plasmid transformation.
Problem 2
The amount of extracted OMV-L7Ae is very low.
Potential solution
Pick the single bacterial colony on the antibiotic plate as the culture source, not from the cryopreserved bacteria in glycerin.
After ultracentrifugation, the OMV-L7Ae was not treated as quickly as possible, resulting in redissolution of the OMV-L7Ae in the supernatant.
Problem 3
The template DNA preparation was not successful.
Potential solution
Q5 high-fidelity 2× master mix is highly susceptible to inactivation by repeated freezing and thawing. On the first use, the Q5 high-fidelity 2× master mix was partitioned. Alternatively, replace the Q5 high-fidelity 2× master mix.
Problem 4
The production of boxC/D-mRNA is low in the transcription reaction.
Potential solution
Determine the correct concentration of PCR product from step D1. In the preparation, nuclease-free water was used to reduce the introduction of RNase.
Problem 5
The transfection of HEK-293T cells was unsuccessful.
Potential solution
HEK-293T cells are fragile and should be handled gently in transfection experiments. The density of HEK-293T cells reached 80% when transfected.
OMV-L7Ae and boxC/D-mRNA (EGFP-boxC/D) may be placed for too long, resulting in mRNA degradation. OMV-L7Ae and boxC/D-mRNA (EGFP-boxC/D) can be extracted again to ensure that boxC/D-mRNA (EGFP-boxC/D) is not degraded.
The solution and equipment used in the experiment need to be fully rinsed with DEPC-treated water; ensure that there is no bacterial contamination.
Acknowledgments
This work was supported by grants from the National Key R&D Program of China (2022YFB3808100 and 2021YFA0909900, X.Z.), the CAS Project for Young Scientists in Basic Research (YSBR-010, X.Z.), the Beijing Natural Science Foundation (Z200020, X.Z.), the Beijing Nova Program (Z201100006820031, X.Z.) and the National Natural Science Foundation of China (32222045 and 32171384, X.Z.). This protocol was adapted from a previous study (Li et al., 2022).
Competing interests
G.N., X.Z., and Y.L. are inventors on a filed provisional application China patent (A general mRNA vaccine vector based on bacterial outer membrane vesicles) submitted by the National Center for Nanoscience and Technology that covers the potential diagnostic and therapeutic uses of the vaccine for cancer immunotherapy. The authors declare no other competing financial interests.
Ethical considerations
The animal study should comply with the relevant ethical regulations for animal testing and research. The Institutional Animal Care and Use Committee of the National Center for Nanoscience and Technology approved our animal study.
References
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Cheng, K., Zhao, R., Li, Y., Qi, Y., Wang, Y., Zhang, Y., Qin, H., Qin, Y., Chen, L., Li, C., et al. (2021). Bioengineered bacteria-derived outer membrane vesicles as a versatile antigen display platform for tumor vaccination via Plug-and-Display technology. Nat. Commun. 12(1): e1038/s41467-021-22308-8.
Dolgin, E. (2021). How COVID unlocked the power of RNA vaccines. Nature 589(7841): 189–191.
Gao, X., Feng, Q., Wang, J. and Zhao, X. (2022). Bacterial outer membrane vesicle-based cancer nanovaccines. Cancer Biol. Med. 19(9): 1290–1300.
Li, Y., Ma, X., Yue, Y., Zhang, K., Cheng, K., Feng, Q., Ma, N., Liang, J., Zhang, T., Zhang, L., et al. (2022). Rapid Surface Display of mRNA Antigens by Bacteria‐Derived Outer Membrane Vesicles for a Personalized Tumor Vaccine. Adv. Mater. 34(20): 2109984.
Liang, J., Cheng, K., Li, Y., Xu, J., Chen, Y., Ma, N., Feng, Q., Zhu, F., Ma, X., Zhang, T., et al. (2022). Personalized cancer vaccines from bacteria-derived outer membrane vesicles with antibody-mediated persistent uptake by dendritic cells. Fundam. Res. 2(1): 23–36.
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Liu, C., Feng, Q. and Sun, J. (2019). Lipid Nanovesicles by Microfluidics: Manipulation, Synthesis, and Drug Delivery. Adv. Mater. 31(45): 1804788.
Liu, G., Zhu, M., Zhao, X. and Nie, G. (2021). Nanotechnology-empowered vaccine delivery for enhancing CD8+ T cells-mediated cellular immunity. Adv. Drug Delivery Rev. 176: 113889.
Miao, L., Li, L., Huang, Y., Delcassian, D., Chahal, J., Han, J., Shi, Y., Sadtler, K., Gao, W., Lin, J., et al. (2019). Delivery of mRNA vaccines with heterocyclic lipids increases anti-tumor efficacy by STING-mediated immune cell activation. Nat. Biotechnol. 37(10): 1174–1185.
Miao, L., Zhang, Y. and Huang, L. (2021). mRNA vaccine for cancer immunotherapy. Mol. Cancer 20(1): e1186/s12943-021-01335-5.
Moore, T., Zhang, Y., Fenley, M. O. and Li, H. (2004). Molecular Basis of Box C/D RNA-Protein Interactions. Structure 12(5): 807–818.
Saito, H., Kobayashi, T., Hara, T., Fujita, Y., Hayashi, K., Furushima, R. and Inoue, T. (2010). Synthetic translational regulation by an L7Ae–kink-turn RNP switch. Nat. Chem. Biol. 6(1): 71–78.
Turner, B., Melcher, S. E., Wilson, T. J., Norman, D. G. and Lilley, D. M. (2005). Induced fit of RNA on binding the L7Ae protein to the kink-turn motif. RNA 11(8): 1192–1200.
Wang, S., Cheng, K., Chen, K., Xu, C., Ma, P., Dang, G., Yang, Y., Lei, Q., Huang, H., Yu, Y., et al. (2022). Nanoparticle-based medicines in clinical cancer therapy. Nano Today 45: 101512.
Yue, Y., Xu, J., Li, Y., Cheng, K., Feng, Q., Ma, X., Ma, N., Zhang, T., Wang, X., Zhao, X., et al. (2022). Antigen-bearing outer membrane vesicles as tumour vaccines produced in situ by ingested genetically engineered bacteria. Nat. Biomed. Eng. 6(7): 898–909.
Zhao, X., Zhao, R. and Nie, G. (2022). Nanocarriers based on bacterial membrane materials for cancer vaccine delivery. Nat. Protoc. 17(10): 2240–2274.
Supplementary information
The following supporting information can be downloaded here:
Figures S1: Gating strategy and representative scatter plots for analysis of transfection in HEK-293T cells (cf. Figure 1)
Figures S2: Gating strategy and representative scatter plots for analysis of maturation in BMDCs (cf. Figure 2a-2b)
Figure S3: Gating strategy and representative scatter plots for antigen presentation in BMDCs (cf. Figure 2c)
Article Information
Copyright
© 2023 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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Cancer Biology > Tumor immunology > Cancer therapy
Molecular Biology > RNA > mRNA translation
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4,775 | https://bio-protocol.org/en/bpdetail?id=4775&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
In vitro Selection and in vivo Testing of Riboswitch-inspired Aptamers
MM Michael G. Mohsen
RB Ronald R. Breaker
Published: Vol 13, Iss 13, Jul 5, 2023
DOI: 10.21769/BioProtoc.4775 Views: 793
Reviewed by: Gal HaimovichRakesh Chatrikhi Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nucleic Acids Research Jan 2023
Abstract
Engineered aptamers for new compounds are typically produced by using in vitro selection methods. However, aptamers that are developed in vitro might not function as expected when introduced into complex cellular environments. One approach that addresses this concern is the design of initial RNA pools for selection that contain structural scaffolds from naturally occurring riboswitch aptamers. Here, we provide guidance on design and experimental principles for developing riboswitch-inspired aptamers for new ligands. The in vitro selection protocol (based on Capture-SELEX) is generalizable to diverse RNA scaffold types and amenable to multiplexing of ligand candidates. We discuss strategies to avoid propagation of selfish sequences that can easily dominate the selection. We also detail the identification of aptamer candidates using next-generation sequencing and bioinformatics, and subsequent biochemical validation of aptamer candidates. Finally, we describe functional testing of aptamer candidates in bacterial cell culture.
Key features
• Develop riboswitch-inspired aptamers for new ligands using in vitro selection.
• Ligand candidates can be multiplexed to conserve time and resources.
• Test aptamer candidates in bacterial cells by grafting the aptamer back onto its expression platform.
Graphical overview
Keywords: Aptamer Directed evolution In vitro selection Riboswitch Synthetic biology
Background
Aptamers are ligand-binding oligonucleotides that are becoming increasingly useful for broad applications in diagnostics, therapeutics, and synthetic biology (Keefe et al., 2010; Topp and Gallivan, 2010; Wang et al., 2019). Aptamers occur naturally in the context of riboswitches, where they monitor the concentration of a target ligand and manipulate the folding of an adjoining expression platform to control the expression of their associated genes (Sherwood and Henkin, 2016; Kavita and Breaker, 2023). Engineered aptamers that bind different ligands can be developed using a technique called in vitro selection (Ellington and Szostak, 1990; Tuerk and Gold, 1990). This process involves generating a large combinatorial pool of oligonucleotides, selecting for those that bind a target ligand, and amplifying the selected oligonucleotides. This process is repeated iteratively until aptamers for the target ligand are identified.
Ideally, engineered aptamers could be useful for intracellular applications. However, the physiochemical conditions inside of a cell differ substantially from that of a test tube. Thus, aptamers developed in vitro might fail to perform inside of a cell, likely due to intrinsic factors such as the failure to reliably fold into the structure required to form the ligand binding pocket (Filonov et al., 2014). To address this, researchers have exploited the architectures of natural riboswitch aptamers to provide scaffolds for combinatorial RNA pools (Porter et al., 2017; Dey et al., 2022; Mohsen et al., 2023). We recently reported the Graftamer approach (Mohsen et al., 2023), in which engineered aptamers that contain a natural riboswitch scaffold are grafted back onto the natural expression platform (Figure 1). Using this approach, we developed aptamers for quinine and caffeine that retain the Guanine-I riboswitch (Mandal et al., 2003) scaffold from the initial combinatorial pool. These aptamers were each grafted back onto the expression platform of a Bacillus subtilis xpt-pbuX Guanine-I riboswitch. The resulting engineered quinine and caffeine riboswitches each display ligand-mediated gene regulation in B. subtilis cultures, indicating that the quinine and caffeine aptamers are functional in cells.
Figure 1. Overview of the Graftamer approach. First, a combinatorial RNA pool is designed by inserting regions of random RNA sequence (Nx, Ny, and Nz) in between structural features of a natural riboswitch aptamer. The depicted riboswitch contains an aptamer with paired elements P1, P2, and P3. Second, in vitro selection is performed to develop riboswitch-inspired aptamers that bind ligands different than that of the natural riboswitch. Third, validated aptamers that maintain the structural features of the natural riboswitch are grafted back onto the expression platform of the natural riboswitch to construct engineered riboswitches. Fourth, plasmids containing engineered riboswitches positioned upstream of a reporter gene are transformed into a suitable model organism. In the depicted example, the riboswitch functions as an OFF switch. Increased ligand concentration reduces the expression of a lacZ reporter gene, resulting in a corresponding decrease in blue color in the presence of X-gal indicator.
Here, we provide a protocol for this approach, beginning from the design and generation of the initial RNA pool. The selection process described herein is based on Capture-SELEX (Nutiu and Li, 2005; Stoltenburg et al., 2012; Yang et al., 2016; Lauridsen et al., 2018; Boussebayle et al., 2019), wherein the RNA pool is hybridized to a 3′-biotinylated DNA capture oligonucleotide, which itself is immobilized on a streptavidin-agarose column (Figure 2). In principle, RNA molecules that undergo a structural change upon binding a ligand are able to release from the capture oligonucleotide (Nutiu and Li, 2005). However, selfish molecules that slowly release from the capture oligonucleotide in a ligand-independent manner can jeopardize the selection (Mohsen et al., 2023). To counter the proliferation of selfish molecules, we recommend stringent washing and relatively short incubation times. Contamination between parallel lines of in vitro selection poses a threat as well. Thus, we suggest designing a different set of primers for each selection line performed in the same laboratory space. We anticipate that this protocol might have reduced utility for researchers pursuing aptamers of compounds that occur naturally in the target organism or that are unable to accumulate to appreciable intracellular concentrations (e.g., due to rapid efflux or metabolism, or an inability to permeate the cell wall or membrane). Nevertheless, we expect that this protocol will be broadly useful for improving best practices for in vitro selection and for increasing the likelihood of success for other researchers working to develop novel aptamers.
Figure 2. Overview of the in vitro selection scheme. First, the template oligodeoxynucleotide and forward primer are used to generate the initial DNA pool by primer extension. Second, in vitro transcription with T7 RNA polymerase is performed to produce the initial RNA pool. Third, the RNA pool is hybridized with a 3′-biotinylated DNA capture oligonucleotide. Fourth, RNA molecules that display affinity for a ligand candidate are isolated via a selection step. Fifth, the RNA molecules that survive the selection step are amplified by reverse transcription-polymerase chain reaction (RT-PCR). Steps 2–5 are repeated iteratively until the RNA pool is sufficiently enriched with functional aptamers. Finally, aptamer sequences are identified by sequencing and testing.
Materials and reagents
Biological materials
Bacillus subtilis strain 1A1 (American Type Culture Collection)
Escherichia coli strain BW25113 (Coli Genetic Stock Center at Yale University)
Reagents
Agarose (IBI Scientific, catalog number: IB70071)
Boric acid (Sigma, catalog number: B0394)
Bromophenol blue (Sigma, catalog number: B5525)
Dimethylformamide (Sigma, catalog number: 319937)
Dithiothreitol (DTT) (American Bioanalytical, catalog number: AB00490)
Ethanol, 200 proof HPLC grade (Sigma, catalog number: 459828)
Ethylenediaminetetraacetic acid (EDTA) (Sigma, catalog number: E5134)
Glacial acetic acid (J.T. Baker, catalog number: 9508-33)
HEPES [(4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid] (American Bioanalytical, catalog number: AB00892)
Magnesium chloride hexahydrate (MgCl2·6H2O) (J.T. Baker, catalog number: 2444-01)
Potassium chloride (KCl) (Sigma, catalog number: 746436)
Purple 100 base pair DNA ladder (New England Biolabs, catalog number: N0551S)
Purple gel loading dye (6×) (New England Biolabs, catalog number: B7024S)
Sodium acetate (NaOAc) (Sigma, catalog number: 241245)
Sodium hydroxide (NaOH) pellets (J.T. Baker, catalog number: 3722-01)
Spermidine (Sigma, catalog number: S2501)
Sucrose (Sigma, catalog number: S0389)
SuperScript III Reverse Transcriptase (Invitrogen, catalog number: 18080093)
T7 RNA Polymerase (purified in-house; equivalent to New England Biolabs, catalog number: M0251L)
Taq DNA polymerase (New England Biolabs, catalog number: M0273L)
Trizma (Tris) base (Sigma, catalog number: T6066)
TURBO DNase (2 U/μL) (Invitrogen, catalog number: AM2238)
Urea (Sigma, catalog number: U5378)
UreaGel 19:1 concentrate (National Diagnostics, catalog number: EC-830)
UreaGel buffer (National Diagnostics, catalog number: EC-835)
UreaGel system diluent (National Diagnostics, catalog number: EC-840)
X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) (Cayman, catalog number: 16495)
Xylene cyanol FF (Sigma, catalog number: X-4126)
Solutions
2 M KCl (see Recipes)
5 M NaCl (see Recipes)
3 M NaOAc (see Recipes)
1 M MgCl2 (see Recipes)
1 M spermidine (see Recipes)
1 M Tris (pH 7.5 at ~20 °C) (see Recipes)
1 M dithiothreitol (DTT) (see Recipes)
X-gal (100 μg/mL) in dimethylformamide (see Recipes)
0.5 M EDTA (pH 8.0 at ~20 °C) (see Recipes)
Tris-borate-EDTA (TBE) (10×) (see Recipes)
Tris-acetate-EDTA (TAE) (50×) (see Recipes)
Transcription buffer (10×) (see Recipes)
Selection buffer (10×) (see Recipes)
Loading buffer (2×) (see Recipes)
Crush-soak buffer (see Recipes)
Recipes
2 M KCl
Add 14.9 g of KCl to a 100 mL flask. Add deionized H2O to 100 mL. Stir to mix. Filter-sterilize.
5 M NaCl
Add 29.2 g of NaCl to a 100 mL flask. Add deionized H2O to 100 mL. Stir to mix. Filter-sterilize.
3 M NaOAc
Add 24.6 g of NaOAc to a 100 mL flask. Add deionized H2O to 100 mL. Stir to mix. Filter-sterilize.
1 M MgCl2
Add 9.5 g of MgCl2 to a 100 mL flask. Add deionized H2O to 100 mL. Stir to mix. Filter-sterilize.
1 M spermidine
Add 145 mg of spermidine to a 1.5 mL microcentrifuge tube. Add 1 mL of dH2O. Vortex to mix.
Add 1 mL of deionized, sterile H2O (dH2O).
1 M Tris (pH 7.5 at ~20 °C)
Add 121.1 g of Tris to a 1 L flask. Add deionized H2O to ~900 mL. While stirring, adjust pH with hydrochloric acid to pH 7.5. Add deionized H2O to 1 L. Autoclave.
1 M dithiothreitol (DTT)
Add 154 mg of DTT to a 1.5 mL microcentrifuge tube. Add 1 mL of dH2O. Vortex to mix.
X-gal (100 μg/mL) in dimethylformamide
Add 100 mg of X-gal to a 1.5 mL microcentrifuge tube. Add 1 mL of dimethylformamide. Vortex to mix.
0.5 M EDTA (pH 8.0 at ~20 °C)
Add 186.1 g of EDTA to a 1 L flask. Add deionized H2O to ~800 mL. Stir to mix. Add 15 g of NaOH pellets. Stir to mix. While stirring, adjust pH with 10 N sodium hydroxide to pH 8.0. Add deionized H2O to 1 L. Autoclave.
TBE (10× concentrated, 4 L)
To a 4 L flask, add 432 g of Tris base, 220 g of boric acid, 14.9 g of EDTA, and deionized H2O to 4 L. Stir to mix. Filter particulates. Sterilize by autoclaving. The final solution is a 10× concentration buffer containing 0.9 M Tris, 0.9 M borate, and 10 mM EDTA pH 8.0 at ~20 °C.
TAE (50× concentrated, 1 L)
To a 1 L flask, add 242 g of Tris base, deionized H2O to ~800 mL, 57.1 mL of glacial acetic acid, and 100 mL of 0.5 M EDTA pH 8.0 at ~20 °C. Add deionized H2O to 1 L. Filter particulates. Sterilize by autoclaving. The final solution is a 50× concentrated buffer containing 2 M Tris, 1 M acetate, and 50 mM EDTA pH 8.0 at ~20 °C.
Transcription buffer (10× concentrated, 1 mL)
Mix 150 μL of 1 M MgCl2, 20 μL of 1 M spermidine, 500 μL of 1 M Tris (pH 7.5 at ~20 °C), 50 μL of 1 M DTT, and 280 μL of dH2O in a 1.5 mL microcentrifuge tube. 1× mixture: 150 mM MgCl2, 20 mM spermidine, 500 mM Tris (pH 7.5 at ~20 °C), and 50 mM DTT. Store at -20 °C
Selection buffer (10× concentrated, 100 mL)
Mix 4.77 g of HEPES, 50 mL of 2 M KCl, 1 mL of 1 M MgCl2, and 35 mL of deionized H2O together in a flask containing a magnetic stir bar. Adjust the pH of the solution to 7.4 by adding pre-weighed NaOH pellets while stirring. Once a pH of 7.4 is attained, remove any remaining NaOH pellets with a clean spatula. Dry and weigh the pellets. After calculating the amount of NaOH required to adjust the pH (typically ~70 mM), add a volume (typically ~0.6 mL) of 5 M NaCl to the solution to bring the total [Na+] to 100 mM. Add deionized H2O to adjust the final volume of the solution to 100 mL. Filter-sterilize the resulting solution. The final solution is a 10× concentrated selection buffer containing 200 mM HEPES, 1 M KCl, 30 mM NaCl (total [Na+]: 100 mM), and 10 mM MgCl2, pH 7.4 at ~20 °C [1×: 20 mM HEPES, 100 mM KCl, 3 mM NaCl (total [Na+]: 10 mM), and 1 mM MgCl2]. HEPES is light-sensitive, so store this buffer in a darkened compartment (cabinet, refrigerator, etc.).
Loading buffer (2× concentrated, 40 mL)
Add 44 g of urea to a flask and add 28 mL of dH2O. Stir with gentle heat until urea is dissolved. Add 8 g of sucrose, 20 mg of bromophenol blue, 20 mg of xylene cyanole FF, 0.4 mL of 10% SDS, and 4 mL of 10× TBE. Stir with gentle heat until sucrose is dissolved. Store at 4 °C.
Crush-soak buffer
To a 1 L flask, add 40 mL of 5 M NaCl, 10 mL of 1 M Tris (pH 7.5 at ~20 °C), and 2 mL of 0.5 M EDTA (pH 8.0 at ~20 °C). Add deionized H2O to 1 L. Autoclave.
Laboratory supplies
0.5 mL low adhesion microcentrifuge tubes (USA Scientific, catalog number: 1405-2600)
1.5 mL low adhesion microcentrifuge tubes (USA Scientific, catalog number: 1415-2600)
8-strip 0.2 mL PCR tubes (Dot Scientific, catalog number: 401)
Gel-loading pipette tips (VWR, catalog number: 76321-828)
Individual PCR tubes 8-tube strip, clear (Bio-Rad, catalog number: TLS0801)
iTaq Universal SYBR Green Supermix (Bio-Rad, catalog number: 1725121)
Luer-Lok 3 mL syringe (BD, catalog number: 309657)
Micro Bio-SpinTM chromatography columns (Bio-Rad, catalog number: 7326204)
Optical flat 8-cap strips for 0.2 mL tube strips (Bio-Rad, catalog number: TC20803)
PierceTM Streptavidin agarose (Thermo Scientific, catalog number: 20353)
PrecisionGlide needle 27 G × 1/2 (BD, catalog number: 305109)
QIAquick PCR Purification kit (Qiagen, catalog number: 28104)
Sterile pipette tips with filters (filters help to avoid contamination)
Vivaspin 500 molecular weight cutoff 10 kDa (Cytiva, catalog number: 28932225)
Equipment
Analytical balance (Mettler Toledo, catalog number: AG285)
Cary 60 UV-Vis spectrophotometer (originally Varian, now Agilent)
CFX Opus 96 Real-Time PCR System (Bio-Rad, catalog number: 12011319)
Geiger counter (Ludlum)
Gel Doc Go Gel imaging system (Bio-Rad, catalog number: 12009077)
Handheld shortwave (254 nm) ultraviolet lamp (UVP, catalog number: UVG-65)
Innova 42R shaking incubator (Eppendorf, catalog number: M1335-0004)
Mastercycler Nexus GX2 (Eppendorf, catalog number: 6336000015)
Milli-Q Advantage A10 (Millipore-Sigma, catalog number: Z00Q0V0WW)
Mini centrifuge (Thermo Fisher, catalog number: 75004061)
NanoDrop 8000 spectrophotometer (Thermo Fisher, catalog number: ND-8000)
Orion Star A211 Benchtop pH meter (Thermo Fisher, catalog number: STARA2110)
Phosphorimager Typhoon FLA 9500 (originally GE Healthcare, now Cytiva)
PowerPac HV power supply (Bio-Rad, catalog number: 1645056)
Slab Gel Dryer Model 583 (Bio-Rad, catalog number: 165-1745)
Sorvall Legend Micro 21R refrigerated centrifuge (Fisher Scientific, catalog number: 75002446)
Speed Vac (originally Thermo-Savant, now Thermo Fisher)
Standard set of pipettes that can transfer volumes in the range of 1–1,000 μL
Synergy Neo2 Multimode Reader (originally BioTek, catalog number: 1351000, now Agilent)
Vortex-Genie 2 (Scientific Industries, catalog number: SI-0236)
Software
R2R version 1.0.6, 2018-12-10 (https://sourceforge.net/projects/weinberg-r2r/)
CMfinder version 0.4.1.18, 2019-04-22 (https://sourceforge.net/projects/weinberg-cmfinder/)
Scripts toTally.py and selfishCluster.py, 2023-01-09 (Mohsen et al., 2023)
Procedure
Pool design and generation
Choose a riboswitch to exploit as a basis for initial RNA pool design. In the absence of other options, the Guanine-I riboswitch class (Mandal et al., 2003) contains a malleable scaffold that can be exploited for this protocol. Key considerations:
The riboswitch aptamer has conserved tertiary contacts.
A single stem (called P1, Figure 1) encloses the entire aptamer domain.
Crystal structure data are available.
The riboswitch is present in a model organism (e.g., E. coli or B. subtilis) that the researcher can culture with relative ease.
Determine which nucleotides to randomize.
Based on crystallographic data, determine which nucleotides interact with the natural ligand.
Randomize nucleotides that interact with the natural ligand, as well as nucleotides that are not required to form conserved tertiary structural interactions. A greater number of randomized nucleotides provides a larger sequence space, though we have performed successful selections with as few as 23 randomized nucleotides.
Design a 3′-biotinylated DNA capture oligonucleotide.
The capture oligo should be 12–18 nucleotides in length and should contain 10–15 nucleotides of complementarity with a constant region in the RNA pool.
The capture oligo can be designed to compete with the P1 stem of the RNA pool.
The capture oligo should contain a 3′-biotin or 3′-biotin-triethylene glycol (TEG) modification. The TEG spacer provides additional space between the RNA/capture oligo hybrid and the streptavidin/biotin complex. We have performed successful selections using capture oligos with and without the TEG spacer.
Design primer-binding regions and primers for polymerase chain reaction (PCR).
Design forward and reverse primer-binding regions into the RNA pool that have roughly the same melting temperature (Tm). Avoid natural sequences from common laboratory model organisms because this can lead to amplification of nucleic acid contaminants.
Design a forward primer (sense sequence) that starts with a T7 RNA polymerase (RNAP) promoter sequence at the 5′ end. The T7 RNAP promoter sequence with two additional G nucleotides at the 3′ terminus (for increased transcription efficiency) is as follows: TAATACGACTCACTATAGG. RNA transcribed from this template will start with two G nucleotides. Add the sense DNA sequence corresponding to the forward primer-binding region from the RNA pool after these two G nucleotides.
Design a primer (reverse primer) that is the reverse complement DNA sequence of the 3′ primer-binding region from the RNA pool.
Design an oligodeoxynucleotide template pool.
Design a single-stranded reverse complement DNA that contains ~15 base pairs of overlap with the forward primer (not including the T7 RNAP promoter sequence) (Figure 2, Generate Pool Generation). This strand contains randomized nucleotides (N). Hand-mixed phosphoramidites provide an even distribution between all four nucleotides, but machine mixing is typically more cost effective and is sufficient for this selection protocol.
Order and purify custom oligodeoxynucleotides.
Longer strands such as the template pool should be ordered at a synthesis scale (≥ 200 nmol) that ensures a sufficient quantity of full-length material. Shorter strands such as forward primer, reverse primer, and capture oligonucleotide can be ordered at the smallest synthesis scale.
We typically order custom oligodeoxynucleotides with standard desalting and purify in house by denaturing (8 M urea) 10% polyacrylamide gel electrophoresis (PAGE). Alternatively, oligodeoxynucleotides could be ordered with PAGE purification.
Synthesize the generation zero (G0) double-stranded DNA template by primer extension.
To a 0.2 mL tube, add 100 pmol of reverse complement template DNA, 150 pmol of forward primer, and dH2O to 30 μL.
Incubate at 90 °C for 1 min. Cool at room temperature for 3 min.
Add 5 μL of 10 mM dNTPs, 10 μL of 5× first strand buffer (provided by manufacturer), 2.5 μL of 0.1 M DTT, and 2.5 μL of SuperScript III reverse transcriptase (RT).
Incubate at 55 °C for 1 h.
Heat-inactivate RT by incubating at 75 °C for 15 min.
Synthesize the generation zero (G0) RNA pool via in vitro transcription.
Set up the following in vitro transcription reaction in a 0.5 mL tube:
Component
10× transcription buffer
10 mM NTPs
dsDNA G0 template
dH2O
T7 RNA polymerase Quantity
10 μL
20 μL
12.5 μL (~25 pmol)
55 μL
2.5 μL
Optional: Scale up in vitro transcription by setting up multiple reactions to sample a larger sequence space.
Incubate at 37 °C for 2 h or until a white precipitate (Mg2P2O7) is visible at the bottom of the tube (sometimes overnight incubation is required).
During this long incubation, prepare a denaturing (8 M urea) 10% polyacrylamide gel following the protocol provided with National Diagnostics UreaGel reagents. If possible, use a comb with wells that can each accommodate a ~200 μL sample but that are not so large that the band would be diffuse.
Add 1 μL of TURBO DNase. Incubate at 37 °C for 10 min.
Add 101 μL of 2× loading buffer. Load the entire sample in one well. If a ~200 μL sample cannot be accommodated by a single well, the RNA can alternatively be concentrated using an RNA clean-up kit or by precipitation with ethanol. In this case, add a corresponding quantity of 2× loading buffer to the concentrated RNA solution.
Run the gel until the xylene cyanol marker has migrated ~5–10 cm from the well, to adequately separate the RNA band from DNA template and from truncated in vitro transcription products. Remove the gel from the glass plates and cover both sides with plastic wrap.
Visualize the RNA band by ultraviolet (UV) shadowing. In a dark room, place the gel on top of an uncut thin layer chromatography (TLC) silica gel sheet (with fluorescence indicator 254 nm). Shine a handheld UV lamp (short wavelength) on the gel. The RNA band appears as a dark band distinct from the dye band. However, if insufficiently separated, the RNA band might overlap with the xylene cyanol band. Using a marker, draw a box on the plastic wrap to indicate the location of the band. Work quickly to minimize UV light exposure to the RNA. Caution: Avoid personal exposure to UV light.
Excise the marked band with a razor blade. Exclude the bottom and top 20% of the band to avoid propagating unwanted sequences with different lengths than the original pool design.
Transfer the excised gel piece to a pre-weighed 1.5 mL tube. Determine the weight of the excised gel piece. Crush the gel using a small pestle.
Add 2× v/w crush-soak buffer relative to the weight of the excised gel piece. For example, if the gel piece weighs 0.2 g, add 0.4 mL of crush-soak buffer. Vortex briefly and then incubate at room temperature with rotation for at least 30 min (can also be incubated at 4 °C with rotation).
Transfer the solution to a Spin-X filtration column. A benchtop centrifuge operated at maximum speed for ~1 min can be used to filter gel particles from the solution.
Approximate the buffer volume remaining after filtration using a pipette. Add 0.1 volumes of 3 M NaOAc and 2.5 volumes of cold (-20 °C) ethanol (100%). Optionally incubate at -20 °C for at least 20 min or overnight.
Centrifuge at 17,000× g for 15 min at 4 °C.
Aspirate the supernatant using a pipette, taking care not to disturb the pellet. If the quantity of RNA is sufficient, the pellet might be observed at the bottom of the tube, positioned in the direction of centrifugal force.
Dry the pellet by centrifugal evaporation (speed-vac) on medium heat for 5 min or until dry. Alternatively, the pellet can be air dried.
Resuspend the pellet in 50 μL of dH2O. Quantitate the concentration of the resulting solution using a NanoDrop spectrophotometer.
Figure 3. Graphical overview of the selection protocol. A. Laboratory materials required to perform the selection. B. Graphical summary of the selection procedure described in step B2.
In vitro selection
Hybridize RNA pool to capture oligonucleotide.
Prepare the following mixture in a 0.5 mL or 1.5 mL tube:
Component
10× selection buffer
RNA pool
Capture oligonucleotide (10 μM)
dH2O Quantity
10 μL
x μL*
y μL**
90 - x - y μL
*For round one, input 100–1,000 pmol RNA. For subsequent rounds, add 1–10 pmol RNA.
**Add a 10× molar excess of capture oligonucleotide relative to RNA pool input.
Incubate the tube at 90 °C for 1 min and then allow to cool at room temperature for at least 5 min.
Selection:
While the RNA-capture oligonucleotide solution is cooling, prepare a column for selection. Add 100 μL of streptavidin-agarose to a Micro Bio-Spin column using a P1000 pipette (the wider tip openings transfer the bead solution more accurately). Place the column in a 1.5 mL tube.
Prepare an air pressure control device (Figure 3A). Attach a 27 G × 1/2 needle to a 3 mL syringe. Then, poke the needle through the center of a Micro Bio-Spin cap. Pull the syringe plunger until it is fully extended.
Holding the column in one hand and the air pressure control device in the other, place the cap of the device on top of the column with sufficient pressure to create a seal, without shutting the tube. Apply pressure by pressing down on the syringe plunger. This should drain the storage buffer into the collection tube, while the streptavidin-agarose beads remain in the column.
Wash the column with 100 μL of 1× selection buffer six times to equilibrate the column in selection buffer (Figure 3B). Each wash is executed by gently pipetting 100 μL of buffer onto the column resin and subsequently using the air pressure control device to drain the buffer. Two consecutive washes can be collected in a single 1.5 mL tube, after which the column should be transferred to a new collection tube. After six washes, transfer the column to a new 1.5 mL collection tube.
Briefly centrifuge the RNA-capture oligonucleotide solution (after cooling for at least 5 min) and apply the entire 100 μL solution to the column. Use the air pressure control device to push the solution through the resin to the collection tube. To maximize the quantity of biotinylated capture oligonucleotide bound to the streptavidin column, re-apply the eluate to the column two additional times.
Wash the column with 100 μL of 1× selection buffer 10 times to remove RNA molecules retained by nonspecific interactions. In this case, each wash is performed by gently pipetting 100 μL of 1× selection buffer on top of the column resin and then using the air pressure control device to push the solution through, such that it saturates the resin but does not go through into the collection tube. After incubating for 30 s, use the air pressure control device to drain the solution into the collection tube.
Incubate three times with a solution of the chosen ligand candidate(s) in 1× selection buffer for 30 s. It is critical that the incubations with ligand solution are performed identically to the washes described in the preceding step, with the only difference being the presence of the ligand candidates.
Combine the ligand pool eluates and transfer to a Vivaspin 10 kDa molecular weight cutoff column. The mass of RNA pool molecules is expected to be > 10 kDa and should be retained by the column.
Centrifuge at 12,000× g for 15 min at room temperature (~20 °C).
Discard flowthrough. Add 300 μL of dH2O to the column and centrifuge again at 12,000× g for 15 min at room temperature.
Recover the concentrated RNA (typically < 15 μL) by aspirating with a gel-loading tip.
Synthesize complementary DNA (cDNA) by reverse transcription.
During the first round of selection, prepare a generation zero (G0) marker. In parallel with the steps described below, perform reverse transcription using 2 pmol of the initial RNA pool.
To a 0.5 mL tube, add up to 12 μL of the concentrated RNA, 1 μL of 10 mM dNTPs, and 1 μL of 2 μM reverse primer (2 pmol). If the total volume of the concentrated RNA is < 12 μL, add dH2O to a total volume of 14 μL. Store any remaining RNA in excess of 12 μL at -20 °C for archiving.
Incubate the mixture at 65 °C for 5 min and then immediately incubate on ice for at least 1 min.
Briefly centrifuge and then add 4 μL of 5× first strand buffer (provided by manufacturer), 0.1 M DTT (provided by manufacturer), and 1 μL of SuperScript III RT.
Incubate at 55 °C for 60 min.
Incubate at 70 °C for 15 min to heat-inactivate RT.
Quality control and scouting with quantitative polymerase chain reaction (qPCR)
Set up the following two reactions in 0.2 mL tubes:
Component
iTaq master mix
Forward + reverse primers (mixed, 4 μM each)
cDNA (from RT reaction)
dH2O cDNA
10 μL
2 μL
2 μL
6 μL Control
10 μL
2 μL
-
8 μL
Pipette-mix and briefly centrifuge the tubes ensuring that there are no bubbles, which can reflect light in unexpected ways and interfere with the analysis.
Set the qPCR program as follows:
i. 95 °C, 1 min 45 s (initial denaturation)
ii. 95 °C, 15 s (denaturation)
iii. x °C, 30 s (annealing, x is annealing temperature)
iv. 68 °C, 30 s (extension)
v. Fluorescence measurement
vi. Repeat steps ii–v 40×
Note: Annealing temperature can be determined using the NEB Tm Calculator (accessible at tmcalculator.neb.com)
Inspect the resulting fluorescence curves. The control sample often displays an increase in fluorescence intensity after 30–35 cycles, which might indicate amplification of a contaminant. If there is no difference between the cDNA and control curves, or if the curves begin to increase within only a few cycles of each other, the selection might be compromised by contamination.
Determine the number of PCR cycles to perform. This can be done manually by visually inspecting the fluorescence curves. The ideal number of cycles is just after the fluorescence curve for the cDNA sample has plateaued, indicating that amplification is near completion without over-amplification, which can result in undesired PCR products. Ideally, there should be no detectable fluorescence from the control sample at this chosen cycle number. Do not choose a cycle number greater than 30, as this is likely to lead to unwanted amplification of artifacts or contaminants.
Amplify cDNA by PCR.
Note: During the first selection round, continue the preparation of a G0 marker in parallel with the steps described below. Use 5 μL of the G0 reverse transcription reaction instead of the G1 reverse transcription reaction.
Prepare a PCR reaction in a thin-walled 0.2 mL tube as illustrated below. One to three 50 μL PCR reactions can be prepared and performed simultaneously.
Component
dH2O
Standard Taq buffer
10 mM dNTPs
4 μM forward + reverse primers
cDNA
Taq DNA polymerase Quantity
34.5 μL
5 μL
1 μL
5 μL
5 μL
0.5 μL
Using the chosen number of cycles, set up and run a PCR program as follows:
i. 95 °C, 1 min 45 s (initial denaturation)
ii. 95 °C, 15 s (denaturation)
iii. x °C, 30 s (annealing, x is annealing temperature)
iv. 68 °C, 30 s (extension)
v. Repeat steps ii–v y times (y is the chosen number of cycles)
vi. 68 °C, 2 min (final extension)
vii. Hold at 10 °C
Take a 5 μL aliquot from the completed PCR reaction and mix it with 1 μL of 6× purple loading dye in a new tube.
To prepare the G0 marker, add 10 μL of 6× purple loading dye to the PCR tube and optionally transfer to a 1.5 mL tube for easier storage at 4 °C or -20 °C.
Prepare a 1.5% agarose gel by dissolving 0.6 g of agarose in 40 mL of TAE in a 250 mL Erlenmeyer flask. Microwave the solution in bursts with frequent stirring until the agarose is completely dissolved. Just before casting the gel, add 2 μL of ethidium bromide and mix by swirling.
Load 5 μL each of 100 base pair ladder, G0 marker, and the PCR solution from the current generation in consecutive lanes.
Run the gel at 115 V for 20 min. Visualize the resulting gel using a Bio-Rad Gel Doc Go Gel imaging system or an equivalent instrument. The bands resulting from the G0 marker and from the current generation should appear to have migrated the same distance, indicating that the amplicons originated from the pool and not from a contaminant.
Purify the PCR product using a QIAquick PCR purification kit or equivalent.
Quantitate the concentration of DNA using a NanoDrop spectrophotometer. Convert the concentration from ng/μL to μM by dividing by the approximate molecular weight of the DNA construct.
Synthesize the RNA pool for the subsequent generation by in vitro transcription.
Set up the following in vitro transcription reaction in a 0.5 mL tube:
Component
10× transcription buffer
10 mM NTPs
dsDNA from PCR
dH2O
T7 RNA polymerase Quantity
10 μL
20 μL
x μL (10 pmol)
67.5 - x μL
2.5 μL
Follow the protocol for in vitro transcription from section A.
Iterative rounds of selection:
Perform iterative rounds of selection until the pool is sufficiently enriched. After the first few rounds, the researcher can optionally decide to decrease the concentration of ligand candidates to apply additional selection pressure. Another option that can be applied concurrently is to decrease the quantity of input RNA in later selection rounds.
Typically, the RNA pool will be sufficiently enriched after 8–12 rounds of selection. Enrichment can be assessed by an elution profile, which is performed similarly to the selection process described above.
Elution profile
Note: This step involves handling radioactive materials (32P). If the researcher’s laboratory is not equipped to handle radioactive materials, the entire process could be performed with unlabeled RNA. In this case, qRT-PCR could alternatively be used to determine relative quantities of RNA eluted with each ligand candidate.
Prepare the following mixture in a 0.5 or 1.5 mL tube:
Component
10× selection buffer
5′ 32P-labeled RNA pool
Capture oligonucleotide (10 μM)
dH2O Quantity
10 μL
x μL (~50,000 counts per minute)
1 μL
89 - x μL
Incubate the tube at 90 °C for 1 min and then allow to cool at room temperature for at least 5 min.
While the RNA-capture oligonucleotide solution is cooling, prepare a column for the elution profile. Add 100 μL streptavidin-agarose to a Micro Bio-Spin column using a P1000 pipette. Place the column in a 1.5 mL tube.
Prepare an air pressure control device: attach a needle to a 3 mL syringe. Then, poke the needle through the center of a Micro Bio-Spin cap. Pull the syringe plunger until it is fully extended.
Holding the column in one hand and the air pressure control device in the other, place the cap of the device on top of the column with sufficient pressure to create a seal, without shutting the tube. Apply pressure by pressing down on the syringe plunger. This should drain the storage buffer into the collection tube, while the streptavidin-agarose beads remain in the column.
Wash the column with 100 μL of 1× selection buffer six times to equilibrate the column in selection buffer. Each wash is executed by gently pipetting 100 μL of buffer onto the column resin and subsequently using the air pressure control device to drain the buffer. Two consecutive washes can be collected in a single 1.5 mL tube, after which the column should be transferred to a new collection tube. After six washes, transfer the column to a new 1.5 mL collection tube.
Briefly centrifuge the RNA-capture oligonucleotide solution (after cooling for at least 5 min) and apply the entire 100 μL solution to the column. Use the air pressure control device to push the solution through the resin to the collection tube. To maximize the quantity of biotinylated capture oligonucleotide bound to the streptavidin column, re-apply the eluate to the column two additional times. After applying and eluting the solution three times, label the tube containing the final eluate as Unbound RNA.
Incubate the column with 100 μL of 1× selection buffer for 30 s to remove RNA molecules retained by nonspecific interactions. Each incubation is performed by gently pipetting 100 μL of 1× selection buffer on top of the column resin and then using the air pressure control device to push the solution through, such that it saturates the resin but does not go through into the collection tube. After incubating for 30 s, use the air pressure control device to force the solution through into the collection chamber. Repeat this step until eluates reach background radiation, as assessed by a Geiger counter (typically 6–8 incubations). Collect each eluate in a separate tube.
Incubate the column with a 100 μL solution of one of the chosen ligand candidates in 1× selection buffer. It is critical that the incubations are performed identically to the negative selection steps above, with the only difference being the presence of the ligand candidate. Repeat this step two times, collecting each eluate in a separate tube. If you performed selection with only one ligand candidate, the elution profile, skip to step C12.
Incubate the column with 100 μL of 1× selection buffer for 30 s to remove the previous candidate ligand as well as RNA molecules retained by the column. Repeat this step two additional times before applying the solution with the next candidate.
Incubate the column with a 100 μL solution of the next ligand candidate in 1× selection buffer three times. Repeat this step for all ligand candidates, washing three times with 1× selection buffer in between candidates.
After all compounds have been assayed, use a pen to mark a grid on a piece of filter paper. The grid should be able to accommodate all samples. Pipette 1 μL of each sample in the center of each square marked by the gridlines. Allow the filter paper to air dry before covering it in plastic wrap.
Expose the filter paper to a phosphor screen overnight and image the following day using a Typhoon phosphorimager. Note whether an increase in signal is observed in eluates with any of the compounds relative to the eluates without compound (see Figure 4).
If selection was performed with multiple ligand candidates, repeat the elution profile, this time reversing the order of the ligand candidates. Signals usually fade over the course of the elution profile as less radiolabeled RNA remains on the column. This can sometimes provide a false positive signal for ligand candidates that are assayed earlier in the experiment.
If the RNA pool appears to respond to one or more of the ligand candidates, continue on to Section D. Otherwise, more selection is required to further enrich the RNA pool.
Figure 4. Illustration of a hypothetical elution profile readout that would be performed after a selection with three ligand candidates: A, B, and C. The eluate containing unbound RNA (UR) is followed by six washes with selection buffer (W1–W6), after which radiation levels are close to background radiation. Eluates containing compound A (A1–A3), compound B (B1–B3), and compound C (C1–C3) are each followed by three washes with selection buffer. Based on this result, the aptamer or RNA pool would appear to respond to compound B.
Identification of candidate sequences using next generation sequencing and bioinformatics
Preparation of DNA pools for next-generation sequencing is better covered in other protocols. Briefly, prepare an aliquot of the desired DNA library (at least 50 ng). We typically submit samples to a sequencing core (Yale Center for Genomic Analysis) where an Illumina NovaSeq is used to perform next generation sequencing with a read depth of ~40 million reads. Paired-end reads are sequenced with a read length of 150 base pairs. Ensure that the read length is greater than the length of the DNA pool including the T7 RNAP promoter sequence.
After obtaining the sequencing data files (.fastq.gz format), use toTally.py to count the number of reads for each unique sequence and rank them according to abundance in tab-separated values (.tsv) format.
$ ./toTally.py -i <path/to/file1.fastq.gz> -j <path/to/file2.fastq.gz>
-5 <fwdPrimerSequence> -3 <revPrimerSequence>
Generate putative classes for the top-ranked sequences using selfishCluster.py. This program outputs several fasta (.fa) files for each candidate. The 5′ and 3′ input sequences here are the primer binding regions of the RNA library (sense sequence). With default parameters, the output will be five fasta files. There will be fewer output files if two or more of the top five ranked sequences belong to the same class.
$ ./selfishCluster.py -i <file.tsv> -5 <5' constant region> -3 <3' constant region>
Use CMfinder to generate a list of conserved secondary structure motifs for each class. This step will generate many Stockholm (.sto) files. Many of these motifs represent a small portion of the full-length RNA.
$ ./cmfinder-0.4.1.18/bin/cmfinder04.pl -skipClustalw -combine <file.fa>
Use R2R to draw the conserved motifs outputted by CMfinder. If it is not possible to identify which .sto file contains the full-length RNA, it might be necessary to use R2R to draw all of them and inspect each output (.pdf) manually.
$ ./R2R-1.0.6/src/r2r -GSC-weighted-consensus <fileName.sto> <fileName_cons.sto>
3 0.97 0.9 0.75 4 0.97 0.9 0.75 0.5 0.1
$ ./R2R-1.0.6/src/r2r -disable-usage-warning <fileName_cons.sto> <filename_cons.pdf>
Identify conserved nucleotides that can be exploited to design disruptive mutations.
Biochemical validation of aptamer candidates
Perform an elution profile as described in Section C using a single aptamer candidate instead of an RNA pool. For a valid aptamer candidate, we typically expect to observe an increase in signal for only one of the compounds.
There are various biochemical methods that can be used to further validate aptamer-ligand binding, including in-line probing (Soukup and Breaker, 1999), selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE) (Merino et al., 2005), isothermal titration calorimetry (Slavkovic and Johnson, 2023), and surface plasmon resonance (Arney and Weeks, 2022). Our lab prefers in-line probing, for which 5′ 32P-labeled RNAs already prepared for the elution profile can be utilized. Refer to previously reported protocols for in-line probing (Regulski and Breaker, 2008). Techniques such as in-line probing and SHAPE that provide structural information are especially useful because they can confirm whether the engineered aptamers retain the structural features of the riboswitch aptamer from which they were derived. With the user’s preferred method for biochemical validation, test candidate aptamers for binding with the target ligand and confirm that disruptive mutant(s) display reduced binding.
Grafting aptamer candidates onto their expression platforms and functional testing in cells
Starting from the sequence of the natural riboswitch that was chosen in part A, remove the entire aptamer domain as well as 2–3 base pairs in the P1 stem located adjacent to the aptamer domain (Figure 1).
Remove the primer-binding regions from the sequence of a biochemically validated aptamer, as well as the sequence of the P1 stem, except for the 2–3 base pairs immediately adjacent to the aptameric core. The number of base pairs retained from the aptamer sequence should be equal to the number that is removed from the expression platform sequence.
Graft the engineered aptamer sequence onto the natural expression platform.
Place a common promoter sequence (e.g., thiC for E. coli, lysC for B. subtilis) upstream of this engineered riboswitch.
Using molecular cloning techniques, install this sequence upstream of a lacZ reporter gene within an appropriate plasmid. Another reporter gene, such as green fluorescent protein or luciferase can optionally be used.
Transform this plasmid into a model organism that naturally contains the original riboswitch.
Test the function of the engineered riboswitch by culturing the transformed cells in media supplemented with X-gal (100 μg/mL). If the target ligand does not occur naturally in the cell (e.g., a drug compound), supplement different cultures with and without ligand. Differential blue color between cultures with or without the target ligand indicates that the riboswitch is functional.
Confirm that the directionality of the chosen riboswitch (ON or OFF switch) is reflected in the observed result. Additionally, confirm that the switching effect diminishes between cultures that contain a disruptive mutant construct.
Data analysis
Analysis of next-generation sequencing data as described in this protocol requires some familiarity with bash commands. Other software, such as FASTAptameR 2.0 (Kramer et al., 2022), can be accessed via the web to facilitate this analysis.
Validation of protocol
To validate the function of engineered quinine and caffeine riboswitches, we quantitated specific β-galactosidase activity using a Miller assay in the presence and absence of the ligand (Mohsen et al., 2023). Three technical replicates were performed. Statistical analysis was performed with a t-test (two-tailed distribution, two sample equal variance).
Acknowledgments
RNA research in the Breaker laboratory is supported by the Howard Hughes Medical Institute (HHMI). M.G.M. is an HHMI Awardee of the Life Sciences Research Foundation. This protocol was originally employed in a recently published article (Mohsen et al., 2023).
Competing interests
The authors declare no competing interests.
References
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LiverQuant: An Improved Method for Quantitative Analysis of Liver Pathology
DH Dominick J. Hellen
SK Saul Joseph Karpen
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4776 Views: 1272
Reviewed by: Masahiro MoritaAmr Galal Abdelraheem IbrahimNimesha Tadepalle
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Abstract
Current means to quantify cells, gene expression, and fibrosis of liver histological slides are not standardized in the research community and typically rely upon data acquired from a selection of random regions identified in each slide. As such, analyses are subject to selection bias as well as limited subsets of available data elements throughout the slide. A whole-slide analysis of cells and fibrosis would provide for a more accurate and complete quantitative analysis, along with minimization of intra- and inter-experimental variables. Herein, we present LiverQuant, a method for quantifying whole-slide scans of digitized histologic images to render a more comprehensive analysis of presented data elements. After loading images and preparing the project in the QuPath program, researchers are provided with one to two scripts per analysis that generate an average intensity threshold for their staining, automated tissue annotation, and downstream detection of their anticipated cellular matrices. When compared with two standard methodologies for histological quantification, LiverQuant had two significant advantages: increased speed and a 50-fold greater tissue area coverage. Using publicly available open-source code (GitHub), LiverQuant improves the reliability and reproducibility of experimental results while reducing the time scientists require to perform bulk analysis of liver histology. This analytical process is readily adaptable by most laboratories, requires minimal optimization, and its principles and code can be optimized for use in other organs.
Graphical overview
Keywords: Cholangiocyte Macrophage Fibrosis Whole slide quantification Immunohistochemical Immunofluorescence Bile duct Batch
Background
Histology is the use of immunodetection-based reagents to visualize cellular and extracellular compartments of tissue, usually via thin-slice light or fluorescent microscopic analysis. Histologic analysis is routinely applied daily in clinical and research settings and is considered essential for visually assessing the structure and pathology of the tissue under study. Advances in cell-based immunohistochemical and immunofluorescent detection protocols have led to multi-faceted progress in most biological fields (Kamimoto et al., 2020; Ramachandran et al., 2020; Tan et al., 2020; Taylor, 2020; Guilliams et al., 2022). Historically, light microscopy has been utilized by pathologists to determine disease etiology as well as determining the extent or degree of damage. More recently, there has been a need, more so in research settings, to provide quantitation of cellular infiltrates, proliferation, and cell-specific expression of selected protein markers, to define the cellular response to damage as well as to identify pathways of repair (Ben-Moshe and Itzkovitz, 2019; Saviano et al., 2020; Hu et al., 2022). The typical method of quantifying cells or fibrosis has been to randomly select regions of interest (ROI) and manually count cells or notable puncta (Abercrombie, 1946; Endo et al., 2002; Baratta et al., 2009; Vogel et al., 2015; Baghdasaryan et al., 2016; Husain et al., 2018). There are no standards for this approach but, by definition, there are inadvertent opportunities for significant regional cherry-picking, and as it does not take into account the entire tissue area of the slide, this method may not provide accurate quantitation (Baker, 2016; Murphy and Aguinis, 2022). Moreover, staining variation between and within slides, small sample sizes (< 1,000 detections/sample), extensive use of laboratory time, and no agreed-upon method all limit the utility and applicability of these methods between experiments and laboratories (Gurcan et al., 2009; Alturkistani et al., 2016; Bogen, 2019).
To address these issues, we focus on this over-arching problem using a specific model organ, the liver. The liver serves as the human body’s biofilter and can be highly reactive in response to disease, genetic predisposition, or external challenges (Nagy et al., 2020). The main cell types in the liver include parenchymal hepatocytes, liver sinusoidal endothelial cells, Kupffer cells/macrophages, cholangiocytes, and endothelium (Heymann and Tacke, 2016). Biomarkers of liver-related injury predominately present through the recruitment of F4/80+ macrophages, an increase in the extracellular production of collagen, and the rapid expansion of cholangiocytes (Fabris et al., 2019).
In this report, we present our experience with an automated whole-slide approach to quantify cells and fibrosis in standard liver slide histology termed LiverQuant. LiverQuant is a new customizable method that uses scripts written in QuPath to automate the detection of macrophages, fibrosis, and cholangiocytes in whole-slide scanned images of liver tissue. QuPath is a free, open-source program for analyzing pathology, and can be used by anyone with an efficient computer (Bankhead et al., 2017). Recently, LiverQuant methodology was used to quantitate various cell types and fibrosis in a novel genetic-based mouse model of biliary atresia, the liver conditional knockout of Pkd1l1 (Hellen et al., 2023).
While this protocol is intended for the detection of specific cell and extracellular markers within the liver, its principles and code can be optimized for use in other organs, within any species, and can be accomplished using either Mac or Windows-based computers.
Materials and reagents
CK19, 1:100 (DSHB, catalog number: TROMA-III)
Alpha-actin (αSMA), 1:250 (Sigma, catalog number: A2547)
F4/80, 1:300 (Cell Signaling, catalog number: CS70076)
Picosirius Red/Fast Green, KIT (Chondrex, catalog number: 9046)
2° Antibodies (immunofluorescence), 1–100 (Alexa Fluor® Invitrogen)
HRP, mouse (immunohistochemistry), 1–100 (Cell Signaling, catalog number: 8125)
SignalStain® DAB (immunohistochemistry), KIT (Cell Signaling, catalog number: 8059)
DAPI, 1:1,000 (Invitrogen, catalog number: D1306)
Equipment
Vectra Polaris Scanner
Note: Any microscope or scanner that can scan whole sections is sufficient.
Computer: tested and optimized with Precision 5820 (2021) with Intel 4.7 GHz processor, Nvidia Quadro RTX4000, 64 GB of RAM, 4 TB SSD, two 8 TB hard drives on RAID 0, and a 30" monitor. Additionally validated with MacBook Pro (2020), M1 processor, and 16 GB of RAM
Note: This protocol is achievable with computers/laptops that have less computing power.
Software
QuPath 0.4.2 open-source software (https://qupath.github.io/)
QuPath scripts are available at https://github.com/DominickHellen/LiverQuant
GraphPad Prism (https://www.graphpad.com)
Procedure
Immunohistochemistry and scanning
The method utilized for isolation, preparation, and scanning of liver sections will not be covered within this protocol. All mouse liver sections that were used for this method were fixed in a standard approach with paraformaldehyde, embedded in paraffin, sectioned at 5 μm thickness, and stained using the appropriate immunohistochemical techniques. Within this study, we will be using mouse sections that were stained using primary antibodies against cytokeratin 19 (cholangiocytes) with α-smooth muscle actin as an endothelial counterstain, F4/80 (macrophages), and picosirius red (collagen) with a Fast Green counterstain. For additional details see Hellen et al. (2023). All sections were imported into QuPath in NDPI file format taken using a Vectra Polaris scanner at 40× magnification. QuPath supports a bank of file extensions for importing, spanning greater than 150 different file formats. Prior to importing files for downstream analysis using QuPath, it is suggested to transfer the desired files onto the computer that will be used for quantification. This will aid in speed and processing time, as internal file access speed is typically the most rate-limiting factor in any of these analyses. It is additionally recommended that an entire cross-section (whole slide) is scanned into QuPath, rather than individual smaller ROIs.
Downloading GitHub files (https://github.com/DominickHellen/LiverQuant)
Navigate to the GitHub repository link.
Download the ZIP file of contents by clicking <> Code → Download ZIP (Figure 1A).
Figure 1. Retrieving LiverQuant files from GitHub. (A) All files necessary for the protocol are available for download online. The red arrow indicates exactly where to download the necessary files. B) All .GROOVY and .JSON files should be available for use after download and unzipping.
Unpack ZIP files by right-clicking → Open.
All files for LiverQuant Bio-Protocol should be within the code folder (Figure 1B).
Preparing the project
Preparing a project in QuPath starts with creating a folder in a file manager of choice (e.g., Mac Finder, Windows File Explorer). To have all files of interest in the right place, it is better to create a folder on the computer and place all files within that folder.
Within the designated folder that contains all the files of interest, create another folder. This will be the hub that QuPath will use for all related paths and metadata (Figure 2A).
Note: Keep this folder empty and entitle it New Folder.
Figure 2. Preparing LiverQuant. A) Representative image of the New Folder where the images and scripts needed for analysis will be stored. B) Example of images ready to be imported into QuPath. C) Within New Folder, generate the scripts folder, where D) all downloaded .GROOVY scripts should be placed. E) Within the classifiers folder, generate pixel_classifiers folder, and F) place all downloaded .JSON files.
Download the most recent edition of QuPath and start the program.
Note: These scripts have been designed and tested using QuPath v0.4.3.
Create a project within QuPath using the Create Project button.
Navigate to the empty New Folder that has been created in step 1 and click Select folder.
The Add images button within QuPath should now be accessible. Click Add images and proceed to add the desired histological image files.
After selecting chosen images, click Open.
Verify that all of the images are added to the Import Images to project pop-up and click Import (Figure 2B).
Leave the Image provider as Default (let QuPath decide).
Leave the image type blank.
Note: Image type will vary depending on the analysis (Brightfield H-DAB – Macrophages, Brightfield Other – Fibrosis, or Fluorescence – Cholangiocytes) and is automatically selected within the detection scripts.
Do not choose to rotate the image, as it can occasionally slow down QuPath’s ability to read the file.
Leave Auto-generate pyramids checked.
Note: Most whole-slide scanned images are pyramidal, which carries data at multiple resolutions and can be sub-sampled depending on the analysis.
The Import objects option can remain unchecked for a new project.
Finally, navigate back to the New Folder in the file manager and add a folder entitled scripts (Figure 2C).
Place Fibrosis detection script, Macrophage detection script, Cholangiocyte detection script, Fibrosis intensity script, and Macrophage intensity script (.GROOVY files), taken from the GitHub link, into the scripts folder (Figure 2D).
Note: These scripts should show up in the toolbar under Automate > Project scripts…
Within New Folder, open classifiers, and create a new folder entitled pixel_classifiers (Figure 2E).
Add the Fibrosis and Macrophage .JSON files into the pixel_classifiers folder (Figure 2F).
QuPath immunohistochemical and immunofluorescent analysis
Whole-slide analysis of immune cell types, fibrosis, and biliary tract composition (cholangiocytes) can provide researchers with an efficient understanding of liver health. This protocol will outline the automated quantification of macrophages, fibrosis, and cholangiocytes within whole-slide sections of liver tissue, respectively. This section will be divided by the respective arc of liver histology chosen for analysis. Each protocol relies heavily upon customizable scripts written in the Groovy language within QuPath. The scripts are open-source and contain detailed annotations of each function and command used for their respective analysis. It is highly recommended to analyze the scripts prior to performing analyses in order to avoid potential pitfalls, optimize, and understand possible errors.
Automated quantification of macrophages within QuPath
Note: For this type of analysis, the 3,3′-Diaminobenzidine (DAB) chromogenic dye was used to detect the secondary antibody attached to the primary anti-F4/80 antibody.
Open one image by double-clicking the image icon.
Set the image type to Brightfield H-DAB (Figure 3A).
(Automatic Thresholding) Annotate three ROIs (using the brush tool) within the image that are good samples of how intense the macrophage stain is (Figure 3B).
Tip: The larger the ROIs, the more accurate the downstream analysis will be.
Note: Self-thresholding is outlined in section H.
When finished annotating, run the macrophage intensity analysis by going to Automate → Project scripts… and clicking on Macrophage intensity script.
For this script to run effectively, researchers need to type in their directory pathname on lines 24 and 29.
Note: This protocol segment provides the Macrophage detection script with an average intensity threshold for quantification specific to the selected histological samples.
Once the Macrophage intensity script is completed, open the Macrophage detection script in Automate → Project scripts… → Macrophage detection script.
Within the Macrophage detection script, right-click and click Run… > Run for project.
The script will automate whole tissue annotation (Figure 3C) and quantify macrophages for each image (Figure 3D).
Notes:
i. The running time for each image is approximately 3 min.
ii. Self-annotation for each image is covered in section I.
Figure 3. Macrophage detection using LiverQuant. A) Upon opening the chosen DAB-stained image, set the image type to Brightfield H-DAB. B) Example of three annotations used for the Macrophage intensity script to gather an average intensity of DAB staining. C) The Macrophage detection script will automate tissue annotation and D) generate positive macrophage detections (red). B and C, scale bars: 2 mm. D, scale bars: 25 μm.
If satisfied with the analysis, proceed to section J to export measurements.
Automated quantification of fibrosis within QuPath
Note: For this analysis, a picosirius red stain with a Fast Green counterstain was used. Of course, a different counterstain can be used and still achieve accurate results.
Open one image by double-clicking the image icon.
Set the image type to Brightfield Other (Figure 4A).
(Automatic Thresholding) Annotate three ROIs (using the brush tool) within the image that are good samples of how intense the picosirius red stain is (Figure 4B).
Tip: The larger the ROIs, the more accurate the downstream analysis will be.
Note: Self-thresholding is outlined in section H.
When finished annotating, run the fibrosis intensity analysis by going to Automate → Project scripts… and clicking on Fibrosis intensity script.
For this script to run effectively, researchers need to type in their directory pathname on lines 24 and 29.
Note: This protocol segment provides the Fibrosis detection script with an average intensity threshold for quantification specific to the samples.
Once the Fibrosis intensity script is completed, open the Fibrosis detection script in Automate → Project scripts… → Fibrosis detection script.
Within the Fibrosis detection script, right-click and click Run… > Run for project.
The script will automate whole-tissue annotation (Figure 4C) and quantify fibrosis for each image (Figure 4D).
Notes:
i. The running time for one image is approximately 2 min.
ii. Self-annotation is outlined in section I.
Figure 4. Fibrosis detection using LiverQuant. A) After opening the chosen picosirius red–stained image, set the image type to Brightfield Other. B) Example of three annotations used for the Fibrosis intensity script to gather an average intensity of picosirius red staining. C) The Fibrosis detection script will automate tissue annotation and D) differentiate between positive (fibrotic septa-red) and negative (parenchymal-blue) staining. B and C, scale bars: 2 mm. D, scale bars: 50 μm.
If satisfied with the analysis, proceed to section J to export measurements.
Automated detection of cholangiocytes within QuPath
Note: The script that was written for cholangiocyte detection utilizes the positive cell detection plugin found within QuPath. It may be useful to further optimize the Positive cell detection plugin to a ROI within the slide and apply its values to the .GROOVY script for batch analysis. The most important parameter to adjust, as mentioned in the .GROOVY script is threshold.
Open one image by double-clicking the image icon.
Set the image type to Fluorescence (Figure 5A).
Figure 5. Cholangiocyte detection using LiverQuant. A) Upon opening the chosen cytokeratin 19 (CK19) stained image, set the image type to Fluorescence. B) The Cholangiocyte detection script will automate tissue annotation and C) generate positive cholangiocyte detections. B, scale bar: 2 mm. C, scale bars: 50 μm. αSMA: Smooth muscle alpha-actin.
Begin the analysis by going to Automate → Project scripts… and clicking on Cholangiocyte detection script.
Within the Cholangiocyte detection script, right-click and click Run… > Run for project.
The script will automate whole tissue annotation (Figure 5B) and quantify cholangiocytes for each image (Figure 5C).
Notes:
i. The running time for one image is approximately 5 min.
ii. Self-annotation is outlined in section I.
Proceed to section J for exporting measurements if satisfied with the analysis.
Self-thresholding for fibrosis and macrophages detection within QuPath
Open one image by double-clicking the image icon.
Set the image type to Brightfield (H-DAB) (Figure 3A) for macrophages or Brightfield (other) (Figure 4A) for fibrosis.
Annotate an area of interest (using the brush tool).
When finished annotating, zoom in on any area of interest.
This area will be used as a probe to determine the appropriate filters to quantify the macrophages or fibrosis within the scanned image.
Open the Pixel classification tool within the Classify menu bar and select Pixel classification > Create thresholder.
Within the Create thresholder pop-up, adjust the parameters to accurately quantify macrophages (Figure 6A) or Fibrosis (Figure 6B).
Note: To visualize the positive and negative selections, make sure the Show pixel classification button has been clicked (the button with the letter C on it).
Figure 6. Self-thresholding for downstream LiverQuant analysis. A) Representative photo of Create thresholder tool in QuPath for macrophage, and B) fibrosis detection, respectively. A, scale bars: 50 μm. B, scale bars: 100 μm.
For macrophage detection:
Set Above threshold to Positive and Below threshold to Negative.
For fibrosis detection:
Set Above threshold to Negative and Below threshold to Positive.
Set Channel to Dab for macrophages and Residual for fibrosis.
Set Resolution to High.
Note: The higher the resolution, the slower QuPath will function in the analysis of the section of interest.
Smoothing Sigma can remain at 0 and can be changed to 1 if background staining is strong to help eliminate possible false positive detections.
Starting threshold should be 0.2 for macrophages and 0.0 for fibrosis and optimized from there to isolate true-positive dab or picosirius staining, respectively. Macrophages will show up as red and the background as blue (Figure 6A). Fibrotic septa will show up as red and the background as blue (Figure 6B).
Set Region to Any annotations.
Enter the Classifier name as LiverQuantM to utilize the Macrophage detection script and LiverQuantF to utilize the Fibrosis detection script.
After optimizing the threshold for LiverQuantM or LiverQuantF, proceed to step E6 or F6 for batch detection of macrophages or fibrosis, respectively.
Annotating slides by hand within QuPath
Note: Self-annotation must be followed by the specific steps outlined below.
Annotate the desired areas within each image that require quantification using QuPath’s brush tool (Figure 7A).
Tip: Avoid liver borders, and longitudinally cut large central veins or portal triads, as some will have false-positive staining.
Tip: To subtract annotation, hold Alt for Windows or Command for Mac while annotating.
Figure 7. Self-annotation for downstream LiverQuant analysis. A) Location of the brush annotation tool in the QuPath toolbar is indicated with a black arrow. B) Representative photo of where two slashes // should be placed for self-annotation within any of the detection scripts. Red arrow indicates exact location to place //.
Open the detection script for the specified analysis (e.g., Macrophage detection script, Fibrosis detection script, and Cholangiocyte detection script).
Place two // before lines 17 and 21 of the script (Figure 7B).
Tip: An alternative is to delete both of these lines.
Note: If this is not done, each annotation that is already created will be deleted and replaced by an automated annotation.
Proceed to step E6, F6, or G4 for batch detection of macrophages, fibrosis, or cholangiocytes, respectively.
Exporting measurements
To export measurements, click on Measure > Export measurements, move all the samples analyzed to the Selected side, choose the appropriate Output file destination, and make sure the Export type is set to Annotations and the Separator is set to Comma (.csv). Finally, click Export.
The exported .csv file should contain all the data needed for analysis.
For macrophages, the measurement for comparative analysis is Detections (Figure 8A).
Note: Divide Detections by Area μm2 and multiply by 1,000,000 to normalize to area.
For fibrosis, the measurement for comparative analysis is LiverQuantF: Positive% (Figure 8B).
For cholangiocytes, the measurement for comparative analysis is Num Positive per mm2 (Figure 8C).
Figure 8. Measurements generated by LiverQuant for comparative analyses. Representative photos from the .csv files generated by exporting LiverQuant measurements for A) macrophages, B) fibrosis, and C) cholangiocytes. Red box indicates the important metric for data analysis.
Data analysis
The detection of macrophages, fibrosis, and cholangiocytes using the described protocol has been recently published (Hellen et al., 2023). In summary, the number of macrophages, fibrosis, and cholangiocytes were compared in a genetic model of biliary atresia (the Pkd1l1 liver conditional knockout mouse) between control and Pkd1l1-deficient livers. Macrophages, fibrosis, and cholangiocytes were all significantly increased within the knockout group, and those levels were further exacerbated in response to additional liver injury (bile duct ligation). All comparative analyses using the normalized exported measurements (Detections for macrophages, LiverQuantF: Positive % for fibrosis, and Num Positive per mm2 for cholangiocytes) (Figure 8) can be undertaken in PRISM, R, or Microsoft Excel. In most comparative workflows, a Student’s t-test or one-way ANOVA is sufficient. It is recommended that these comparisons be done using an n ≥ 5 for each group.
Validation of protocol
Fibrosis (Picosirius Red/Fast Green), cholangiocyte (CK19), and macrophage (F4/80) counts were analyzed from the same mouse liver slides using three methodologies: manual count, FIJI, and LiverQuant. Analysis in FIJI was done using a standard cell-counting approach whereby a Python-based script was used to threshold → convert to binary → watershed → analyze particles (Collins, 2007; Crowe and Yue, 2019; Szafranska et al., 2021). Standard metrics such as accuracy, rate of detection, and slide areas covered by maximum ROI were used to assess and establish the validity of LiverQuant.
Accuracy for macrophage and cholangiocyte cell detections was determined by comparing the number of positive cell classifications within the same ROI in LiverQuant and FIJI against the amount quantified by manual cell counting. Photoconversion to binary of a fibrosis-stained ROI resulted in the detection of all septa (manual count) and was compared to the amount recognized using LiverQuant and FIJI. The average rate for each methodology was calculated through the division of positive detections by the amount of time taken (in seconds). The largest ROI area that an individual could accurately quantify (manual count), and that FIJI could similarly work with to accurately detect cells, served as their respective maximum ROIs.
The package of scripts in LiverQuant, when compared to two conventional methodologies of histological quantification (manual and FIJI), demonstrated comparable accuracy of detection and quantification of fibrosis and cell counts (Figure 9A). Importantly, LiverQuant had two significant advantages over conventional quantitative methodologies: increased speed (Figure 9B) and a 50-fold greater tissue area coverage (Figure 9C).
Figure 9. Evaluation of the LiverQuant methodology. A) Fibrosis, macrophage, and cholangiocyte quantification accuracy was compared in LiverQuant, FIJI, and manual counting (True Value; 100%). B) Detections per second for cell counts and fibrosis was faster in LiverQuant vs. FIJI and manual counting. C) The maximum ROI that LiverQuant can utilize for accurate profiling of histological matrices is greater than that of FIJI and manual.
Acknowledgments
Dominick J. Hellen received funding from the NIH (5T32GM008490-30). Saul J. Karpen received funding from the Mason Trust and the Meredith Brown Foundation.
Competing interests
D.H.: None
S.J.K.: Consultant for Albireo, Hemoshear, Intercept, Mirum
Ethical considerations
All mouse experiments were approved by the Emory University Institutional Care and Use Committee (IACUC).
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Article Information
Copyright
© 2023 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
Immunology > Immune cell imaging > Epifluorescence Microscopy
Cell Biology > Cell imaging > Fixed-tissue imaging
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4,777 | https://bio-protocol.org/en/bpdetail?id=4777&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Isoform-specific, Semi-quantitative Determination of Highly Homologous Protein Levels via CRISPR-Cas9-mediated HiBiT Tagging
KS Kristina Seiler
SR Sreoshee Rafiq
MT Mario P. Tschan
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4777 Views: 897
Reviewed by: David PaulManoj B. MenonTalita Diniz Melo Hanchuk
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Original Research Article:
The authors used this protocol in Cell Death & Disease May 2022
Abstract
Many protein families consist of multiple highly homologous proteins, whether they are encoded by different genes or originating from the same genomic location. Predominance of certain isoforms has been linked to various pathological conditions, such as cancer. Detection and relative quantification of protein isoforms in research are commonly done via immunoblotting, immunohistochemistry, or immunofluorescence, where antibodies against an isoform-specific epitope of particular family members are used. However, isoform-specific antibodies are not always available, making it impossible to decipher isoform-specific protein expression patterns. Here, we describe the insertion of the versatile 11 amino acid HiBiT tag into the genomic location of the protein of interest. This tag was developed and is distributed by Promega (Fitchburg, WI, USA). This protocol describes precise and specific protein expression analysis of highly homologous proteins through expression of the HiBiT tag, enabling protein expression quantification when specific antibodies are missing. Protein expression can be analyzed through traditional methods such as western blotting or immunofluorescence, and also in a luciferase binary reporter system, allowing for reliable and fast relative expression quantification using a plate reader.
Graphical overview
Keywords: Protein isoform Antibody-free detection Endogenous tagging Protein expression HiBiT Split-luciferase
Background
There is hardly a protein to be found within the human body that does not share extensive sequence homology to a family member or isoform, as the human cell requires redundancies to ensure continuous work. Protein isoforms are sometimes encoded by different genes and, often, specific genes have multiple transcripts, returning slightly different proteins depending on how they are spliced. Many diseases are characterized by the predominant expression of specific enzyme isoforms, and isoform expression is often quantified at RNA levels by RT-qPCR, sequencing, or microarrays. Quantifying levels of highly homologous proteins, on the other hand, is difficult and limited by the availability of isoform-specific antibodies. Additionally, batch-to-batch variations in antibody performance as well as recognition of unspecific binding partners sometimes hamper precise protein identification through this method (Voskuil, 2017). Furthermore, when studying newly identified isoforms or lesser studied protein families, there are often no monoclonal, isoform-specific antibodies available. We encountered this issue in our investigation of hexokinase 2 (HK2) and HK3, glycolytic enzymes of the hexokinase family. While sequence homology between HK2 and HK3 is only roughly 54%, we struggled to find appropriate antibodies detecting specifically one isoform or the other when 293T cells were transfected with either HK1, HK2, or HK3 and subsequently stained with a monoclonal antibody for HK2 as well as three antibodies for HK3 (two polyclonal, one monoclonal). The use of polyclonal antibodies should be assessed carefully, as larger immunogens increase the likelihood of producing signals from various family members. If the proteins do not vary in size, their distinction is impossible. HK2 and HK3 are both roughly 100 kDa in size; using antibodies against HK3, we detected signals on western blot in samples that were confirmed HK3-negative through mass spectrometry. This prompted us to generate a protocol for isoform-specific detection of endogenous protein expression without cross-reaction bias.
We hereby suggest the use of an endogenous tag that can be inserted into precise genomic locations and allows relative quantification and identification of protein isoforms through various methods. The system was developed by Promega (Fitchburg, WI, USA) (Schwinn et al., 2018 and 2020). The HiBiT® peptide tag used is 11 amino acids small and binds tightly to an adaptor protein (LgBiT®) when supplied. Through this interaction, the bright luminescent protein NanoBiT® is reconstituted (Dixon et al., 2016). Luminescent intensity from reconstituted NanoBiT is directly proportional to the amount of HiBiT present. This allows for relative protein quantification using a common plate reader. Cell lysate can also be analyzed via SDS gel and subsequent incubation of the membrane with LgBiT®. Furthermore, Promega recently launched an anti-HiBiT monoclonal antibody, which allows visualizing protein localization and expression within a cell using fluorescent microscopy. The anti-HiBiT antibody can also be used in traditional western blotting.
In order to insert the tag into the genomic locus of interest, CRISPR/Cas9-mediated genome editing is used. For successful editing, a sgRNA that binds very close to the desired insertion of the HiBiT tag is complexed with Cas9, and single-strand donor DNA templates (ssODN) for homology-directed repair are supplemented. A few days after electroporation of the cells with Cas9/sgRNA ribonucleoprotein complex (RNP) and ssODN, cells can be assessed for insertion of the tag using a plate reader. While luminescent signal for bulk edited cells in culture was stable during cell propagation, we recommend growing monoclonal populations with 100% of cells expressing the tag for reliable protein quantification.
With the recently launched anti-HiBiT monoclonal antibody, further downstream experiments such as fluorescent microscopy are possible but will require prior optimization.
This example follows the tagging of two hexokinase proteins (HK2 and HK3), two highly homologous proteins encoded on different chromosomes (Figure 1). Here, we show the results of adding the HiBiT tag to the C-terminal end, but inserting the tag at the N-terminus resulted in the generation of a luminescent signal as well. While this strategy is therefore applicable to splice isoforms that differ in either the first or last exon, we do not currently know whether insertion of the tag within the gene body will allow for binding of the adaptor protein and whether this technique can be applied for splice isoforms that share the first and last exon. Insertion of the tag within the gene body could potentially alter protein folding and/or function, presenting a potential limitation for this technique.
Figure 1. Schematic illustration of tagging options for homologous proteins hexokinase 2 (HK2) and HK3. HiBiT tag can be inserted at either the N- or C-terminal end. Data shown in this protocol are generated with HK2 and HK3 tagged at their C-terminal end.
Materials and reagents
Gene-specific Alt-R® CRISPR-Cas9 sgRNA [Integrated DNA Technologies (IDT)], store at -80 °C
ssODN: gene-specific Alt-R® HDR Donor Oligos (IDT)
eSpCas9-GFP protein (Sigma-Aldrich, catalog number: ECAS9GFPPR-50UG), store at -20 °C
Note: Using a GFP-tagged Cas9 helps visualize electroporation efficiency, but is not needed
Alt-R® HDR enhancer 100 μL (IDT, catalog number: 1081072), store at -20 °C
NeonTM Transfection System 10 μL kit, containing buffers R, T, and E, as well as NeonTM tips and NeonTM electroporation tubes (Thermo Fisher Scientific, catalog number: MPK1025), store buffers at 4 °C after opening; store the remaining components at room temperature (RT)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S3014)
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9541)
Sodium phosphate dibasic (Na2HPO4) (Sigma-Aldrich, catalog number: S9763)
Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P5655)
Growth medium suitable for the cell type used [here, we used RPMI 1640 (Sigma Aldrich, catalog number: R8758) supplemented with 10% fetal bovine serum]
Nano-Glo® HiBiT lytic detection system (Promega, catalog number: N3030), store at -20 °C
cOmpleteTM proteinase inhibitor cocktail (Roche, catalog number: 11697498001), store at -20 °C
Nano-Glo® HiBiT blotting system (Promega, catalog number: N2410), store at -20 °C
White microplates suitable for luminescence
4%–20% Mini-PROTEAN® TGC Stain-FreeTM protein gel (Bio-Rad, catalog number: 4568093)
Molecular weight marker, such as Precision Plus ProteinTM KaleidoscopeTM Standard (Bio-Rad, catalog number: 1610375EDU)
Trans-Blot Turbo RTA Mini 0.2 μm Nitrocellulose Transfer kit (Bio-Rad, catalog number: 1704270)
Tris-HCl (Sigma-Aldrich, catalog number: 1083190100)
Triton® X-100 (Sigma-Aldrich, catalog number: X100)
Sodium deoxycholate (Sigma-Aldrich, catalog number: D6750)
Tris base (Trizma® base, Sigma-Aldrich, catalog number: 93352)
Tween® 20 (Sigma-Aldrich, catalog number: P1379)
PBS (see Recipes)
Lysis buffer for HiBiT blotting (see Recipes)
TBS-T (see Recipes)
Equipment
NeonTM transfection system (Thermo Fisher Scientific, catalog number: MPK5000)
Luminescence plate reader (Tecan Group, Tecan Infinite® 200 PRO)
2-D protein electrophoresis equipment (Mini-PROTEAN Tetra Vertical Electrophoresis Cell and PowerPac Power supply, Bio-Rad)
Trans-Blot Turbo transfer system
ChemiDoc XRS+ imaging system (Bio-Rad)
Software
gRNA design tools:
https://chopchop.cbu.uib.no/, https://eu.idtdna.com/site/order/designtool/index/CRISPR_SEQUENCE
http://crispor.tefor.net/
ImageLab: acquisition of chemiluminescent images using ChemiDoc imaging system
ImageJ: relative quantification of chemiluminescent signal
Procedure
Design of gRNA and ssODN
Identify desired locus of HiBiT tag insertion (for example at the C- or N-terminal end, directly after the start codon or right before the stop codon).
Identify appropriate gRNA sequences within ± 30 nucleotides of locus of insertion; for improved result, compare gRNA efficiency over various publicly available algorithms (we have not tested gRNA sequences more than 30 nt away from the site of insertion).
Design ssODN with at least 80 nucleotides homology arms upstream and downstream of Cas9 cut site (total length of ssODN is 180–200 nt).
To prevent Cas9 from binding to and cutting the ssODN, insert 2–3 silent mutations into the gRNA sequence within the ssODN, preferentially the PAM sequence if possible.
Table 1. Sequences for homology-directed repair–mediated insertion of HiBiT-tag into C-terminus of Hexokinase isoforms
Protein sgRNA sequence PAM ssODN
HK2 ATAGAACCCCTGAAATCGGA AGG CTGCAGTCAGAGGATGGCAGCGGGAAGGGGGCGGCGCTCATCACTGCTGTGGCCTGCCGCATCCGTGAGGCTGGACAGCGAGTGAGCGGCTGGCGGCTGTTCAAGAAGATTAGCTAGATCCCCTGAAATCGAAAGCGACTTCCTCTTTCTCTCCTTCTTCCCTGTTTTAAATTATAAGATGTCATCCCCTTGTGT
HK3 TCGTGTCTGAGGAAACCTCC CGG GATGGGTCCGGCAAAGGTGCGGCCCTGGTCACCGCTGTTGCCTGCCGCCTTGCGCAGTTGACTCGTGTCGTGAGCGGCTGGCGGCTGTTCAAGAAGATTAGCTGAGGAAACCTCCACGCTGAGGAGGTCTCCGCCGCAGCCTTGCTGGAGCCGGGTCGGGGTCTGCCTGTTTCCCAGCCAGGCCC
blue: HiBiT tag; orange: sgRNA sequence; red: Stop codon; green: PAM sequence; purple: silent mutations inserted to prevent Cas9 from binding to ssODN
Electroporation
Prepare stock solutions:
Cas9: 20 μM
sgRNA: 50 μM
ssODN: 100 μM
In a total volume of 7 μL, add 15 pmol of Cas9 and 75 pmol of sgRNA to buffer R or T (depending on the cell type, see NeonTM instructions).
Note: The ratio between sgRNA and Cas9 can be empirically titrated for best efficiency. We recommend a ratio of 5:1–9:1.
Example, per reaction:
1.5 μL of sgRNA (75 pmol)
0.75 μL of Cas9 (15 pmol)
4.75 μL of buffer R or T
Incubate for 15 min at RT. RNP complexes are stable for 1 h at RT.
Note: RNPs can be complexed prior to the experiment and stored for two weeks at 4 °C or up to at least 10 weeks at -80 °C without loss in performance.
Prepare a 96-well plate with 100 μL of growth medium (suitable for your cell type) containing 30 μM of HDR enhancer to collect cells after electroporation. Prewarm to 37 °C.
Add 75 pmol of ssODN (0.75 μL of 100 μM stock) to the complexed RNPs.
Wash in PBS.
Prepare the electroporator station: fill electroporation tube with 3 mL of buffer E. Settings vary depending on the cell line, and optimized settings and cell numbers for many cell lines can be found at Thermo Fisher (https://www.thermofisher.com/us/en/home/life-science/cell-culture/transfection/neon-transfection-system/neon-transfection-system-cell-line-data.html).
Count cells, take the appropriate cell number, and resuspend in 5 μL of buffer R or T per electroporation, including appropriate controls (electroporation only, untargeted sgRNA).
Add 5 μL of cell suspension to 7 μL of RNP complex. Keep cell exposure to buffers R or T at a minimum (≤ 15 min).
Using a 10 μL NeonTM electroporation tip, electroporate cells and transfer into prewarmed media (step B4).
Transfer cells to a humidified incubator at 37 °C with 5% CO2 immediately.
Do not disturb cells for 24 h.
After 24 h, electroporation efficiency can be assessed visually by examining the presence of eCas9-GFP in live cells using a fluorescent microscope. Handle plate with care and avoid extended exposure to ambient temperature.
Check for HiBiT insertion
Let cells grow until you can comfortably remove 5 × 104 cells.
Prepare Nano-Glo® HiBiT lytic detection reagent by allowing buffer to thaw to RT.
Note: This assay can be performed in various well formats. We prefer using the 384-well format and adding 20 μL of detection reagent to 20 μL of cell suspension. For statistical reasons, we recommend performing the assay in no less than triplicates.
Dilute assay reagents into assay buffer according to the manufacturer’s instructions.
Count cells and add equal numbers of cells per well.
Note: We use 15,000 cells per 384-well plate.
Add detection reagent and ensure homogeneous lysis by pipetting up and down. Be sure to include a no-HiBiT-expressing negative control.
Equilibrate to RT for 10 min.
Measure luminescence.
Monoclonal population
In case editing efficiency or endogenous protein expression is very low, single-cell monoclonal populations can be obtained through limiting dilution assay.
To improve efficiency in growing edited monoclonal populations, we recommend preselecting HiBiT-expressing populations grown from 10 cells and performing limiting dilution assay after preselection.
Of edited bulk population, plate 10 cells/well in a 96-well format.
After enough cells have grown, screen populations for highest HiBiT expression using Nano-Glo® HiBiT lytic detection.
From the highest expressing populations, perform limiting dilution assay.
Grow and screen monoclonal populations.
HiBiT blotting (optional)
To ensure the HiBiT insertion has occurred at the correct protein, the size of the fusion protein can be tested via blotting.
Isolate protein using the lysis buffer for HiBiT blotting (see Recipes) and quantify protein content (e.g., Bradford assay).
Separate the appropriate volume of protein for the expected protein expression on a gel (e.g., 4%–20% Mini-PROTEAN® TGC Stain-FreeTM protein gel) and include a molecular weight marker.
If using stain-free gel technology, activate gel according to the manufacturer’s protocol.
Transfer protein onto a nitrocellulose membrane (e.g., via semi-dry Turbo-Blot transfer).
After transfer, wash membrane in TBS-T (see Recipes). Do not allow membrane to dry.
Acquire total protein image to allow for relative quantification.
Prepare Nano-Glo® HiBiT blotting solution according to the manufacturer’s instructions: dilute LgBiT protein 200-fold with 1× Nano-Glo® blotting buffer, mix by inversion, remove TBS-T, and cover the membrane in blotting solution.
Incubate for 1 h at RT while gently rocking.
Note: For increased signal, incubation can be prolonged up to overnight incubation at 4 °C. If incubating at 4 °C, allow equilibration to RT before adding the substrate after incubation.
Dilute the Nano-Glo® Luciferase Assay Substrate 500-fold into the solution covering the membrane.
Incubate for 5 min at RT.
Acquire chemiluminescent image using an appropriate imager (e.g., ChemiDoc XRS+).
Note: As an alternative to the HiBiT blotting system, a conventional immunoblot can be performed using Promega’s mouse anti-HiBiT mAB (catalog number: N7200).
Perform your experiment
Quantify relative endogenous protein expression at desired time points using NanoGlo® HiBiT lytic detection (Figure 2), Nano-Glo® HiBiT blotting system (Figure 3), or conventional immunoblot.
Figure 2. Relative quantification of hexokinase 2 (HK2) and HK3 protein levels upon all-trans retinoic acid (ATRA, 1 μM) treatment of HiBiT-tagged HL60 AML cell lines. Endogenous tagging allows to specifically detect upregulation of isoform 3, while levels of isoform 2 remain constant.
Figure 3. Nano-Glo® HiBiT blotting system. Steady-state hexokinase 2 (HK2) and HK3 levels in HL60 AML cells detected via HiBiT tag. Of note, HK3 is very lowly expressed at steady-state level in AML cells.
Data analysis
Nano-Glo® HiBiT lytic detection: quantify relative protein expression using luminescence readings, subtracting readings of negative control.
Nano-Glo® HiBiT blotting system: quantify relative protein expression via luminescence signal on images, using software like ImageJ. Normalize to the volume of total protein or to a housekeeping reference protein (β-tubulin, GAPDH).
Notes
To verify insertion of HiBiT tag, it is also possible to perform a PCR reaction using a primer pair that is specific for both the tag as well as the locus of insertion.
Insertion locus of the tag: the optimal tag-insertion locus for each protein of interest should be determined individually. Tag insertion at either the C- or N-terminal end of HK2 and HK3 resulted in the generation of a luminescent signal. We do not currently know if insertion of the tag at genomic regions other than the termini allows for binding of the LgBiT adaptor protein.
Splice variants: this protocol is applicable for highly homologous proteins encoded on different chromosomes as well as splicing variants that differ in either their first or last exon. For other splice isoforms, it would need to be tested whether integration of the HiBiT tag internally allows for binding of the LgBiT adaptor protein while preserving protein function.
Recipes
PBS
137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8mM KH2PO4
Lysis buffer for HiBiT blotting
50 mM Tris-HCl, 150 mM NaCl, 1% Triton® X-100, 0.1% sodium deoxycholate, pH 7.5
TBS-T
20 mM Tris base, 150 mM NaCl, 0.1% Tween® 20
Acknowledgments
We thank Promega for technical support during optimization of this protocol. This study was supported by grants from the Swiss National Science Foundation and Krebsforschung Schweiz (31003A_173219 to MPT and MD-PhD 03/17 Scholarship to KS).
Competing interests
The authors declare no conflict of interests.
References
Dixon, A. S., Schwinn, M. K., Hall, M. P., Zimmerman, K., Otto, P., Lubben, T. H., Butler, B. L., Binkowski, B. F., Machleidt, T., Kirkland, T. A., et al. (2016). NanoLuc Complementation Reporter Optimized for Accurate Measurement of Protein Interactions in Cells. ACS Chem Biol 11(2): 400-408.
Schwinn, M. K., Machleidt, T., Zimmerman, K., Eggers, C. T., Dixon, A. S., Hurst, R., Hall, M. P., Encell, L. P., Binkowski, B. F. and Wood, K. V. (2018). CRISPR-Mediated Tagging of Endogenous Proteins with a Luminescent Peptide. ACS Chem Biol 13(2): 467-474.
Schwinn, M. K., Steffen, L. S., Zimmerman, K., Wood, K. V. and Machleidt, T. (2020). A Simple and Scalable Strategy for Analysis of Endogenous Protein Dynamics. Sci Rep 10(1): 8953.
Voskuil, J. L. A. (2017). The challenges with the validation of research antibodies. F1000Res 6: 161.
Article Information
Copyright
© 2023 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
Cancer Biology > General technique > Biochemical assays
Molecular Biology > Protein > Detection
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4,778 | https://bio-protocol.org/en/bpdetail?id=4778&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Relative Membrane Potential Measurements Using DISBAC2(3) Fluorescence in Arabidopsis thaliana Primary Roots
SD Shiv Mani Dubey
MF Matyáš Fendrych
NS Nelson B.C. Serre
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4778 Views: 557
Reviewed by: Samik BhattacharyaIgnacio Lescano Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Plants Sep 2021
Abstract
In vivo microscopy of plants with high-frequency imaging allows observation and characterization of the dynamic responses of plants to stimuli. It provides access to responses that could not be observed by imaging at a given time point. Such methods are particularly suitable for the observation of fast cellular events such as membrane potential changes. Classical measurement of membrane potential by probe impaling gives quantitative and precise measurements. However, it is invasive, requires specialized equipment, and only allows measurement of one cell at a time. To circumvent some of these limitations, we developed a method to relatively quantify membrane potential variations in Arabidopsis thaliana roots using the fluorescence of the voltage reporter DISBAC2(3). In this protocol, we describe how to prepare experiments for agar media and microfluidics, and we detail the image analysis. We take an example of the rapid plasma membrane depolarization induced by the phytohormone auxin to illustrate the method. Relative membrane potential measurements using DISBAC2(3) fluorescence increase the spatio-temporal resolution of the measurements and are non-invasive and suitable for live imaging of growing roots. Studying membrane potential with a more flexible method allows to efficiently combine mature electrophysiology literature and new molecular knowledge to achieve a better understanding of plant behaviors.
Key features
• Non-invasive method to relatively quantify membrane potential in plant roots.
• Method suitable for imaging seedlings root in agar or liquid medium.
• Straightforward quantification.
Keywords: Auxin In vivo Microfluidics Microscopy Non invasive
Background
Establishment of a membrane potential is essential to support cell function in all living organisms. In plants, membrane potential reflects the electrical difference between the cytoplasm and the apoplast (in root epidermis between -120 and -160 mV, Sze et al., 1999; reviewed in Serre et al., 2021). This phenomenon is mostly driven by the H+-ATPases proton pumps, which extrude positively charged protons from the cytoplasm to the apoplast (Sze, 1985), resulting in negative membrane potentials (Sze et al., 1999). These negative membrane potentials are essential for ion transport across the plasma membranes, and thus play an important role in plant nutrition, growth, and development (Tyerman and Schachtman, 1992; Ragel et al., 2019). Reflecting the spectacular ability of plants to adapt to changing local environments, membrane potential can be quickly shifted from the resting potential value to a hyperpolarized (more negative) or a depolarized (more positive) state. Responses vary according to the stimuli and can potentially lead to stress-induced electrical signaling (Mudrilov et al., 2021). For example, salt stress (NaCl) causes a quick and massive entry of positively charged sodium ions into the root tissues, triggering a leak of potassium ions outside the cells to counterbalance the membrane depolarization (for review, see Cuin et al., 2008). Equally, the hormone auxin triggers almost instantaneous membrane depolarization in roots, leading to root growth inhibition by mechanism(s) still not fully understood (Tretyn et al., 1991; Dindas et al., 2018 and 2020, Paponov et al., 2019; Li et al., 2021; Serre et al., 2021; Qi et al., 2022; Roosjen et al., 2022).
Membrane potential is traditionally measured by impaling electrodes into living tissues (Shabala et al.,2005). While this method gives extremely accurate and quantitative measurements, it is invasive, requires dedicated equipment, and is not fully adapted to the study of growing tissues for a long period of time. Furthermore, the reported potential only concerns one cell and thus limits high resolution spatio-temporal studies.
To circumvent these challenges, we reviewed the literature and available voltage reporter dyes. We selected DISBAC2(3) for its uniform staining of roots and lack of obvious toxicity (Serre et al., 2021). This staining method allowed us to relatively observe membrane potential over time in different tissues. However, the method does not give actual potential values (in mV), lacks sensitivity in strongly hyperpolarized states, and so should always be used in comparison to roots observed under control conditions.
Here, we explain how to analyze relative changes in membrane potential using confocal fluorescence microscopy, taking as an example the imaging of rapid auxin-induced cellular depolarization in Arabidopsis thaliana primary root. We describe how to prepare experiments for imaging of the membrane potential in microfluidics to observe the first instants of the response, as well as imaging in agar media for observation of established responses. Microfluidics is adapted to this type of observation, as each plant can be observed first in control and then in treatment condition, granting internal control fluorescence intensities for every seedling.
The protocols detailed here can be adapted for other dyes to study dynamic events in A. thaliana roots in response to various (a)biotic stimuli.
Materials and reagents
General
Arabidopsis thaliana wildtype Columbia-0 (Col-0) sterile seeds
Porous tape 1.25 cm large (Duchefa, Leucopore, L3302)
Autoclaved wooden toothpicks (any supermarket)
Square Petri dishes (e.g., 15 mm × 15 mm or 10 mm × 10 mm, any brand)
1.5 mL tubes (any brand)
15 mL tubes (any brand)
0.5 mL tubes (any brand)
96% ethanol (Sigma, catalog number: 1.59010), store at room temperature in a flame-protective cabinet
Auxin 3-indoleacetic acid (IAA) (Sigma-Aldrich, catalog number: I2886-5G), store at -20 °C
Murashige and Skoog (MS) medium (Duchefa, catalog number: M0221.0050), store at room temperature
Plant agar (Duchefa, catalog number: P1001), store at room temperature
Sucrose (Duchefa, catalog number: S0809), store at room temperature
MES (Duchefa, catalog number: M1503.0100), store at room temperature
1 M KOH (Sigma-Aldrich, catalog number: 06103), store at room temperature
DISBAC2(3) (Invitrogen, Thermo Fisher, catalog number: B413), store at -20 °C in dark
DMSO (Sigma-Aldrich, catalog number: 94563), store at room temperature
Fluorescein Dextran 10 K MW (Thermo Fisher, Invitrogen, catalog number: D1821)
DISBAC2(3) master stock (see Recipes)
DISBAC2(3) working aliquots (see Recipes)
70% ethanol (approximately) prepared from 96% ethanol (see Recipes)
500 mL of buffered liquid 1/2 MS (see Recipes)
500 mL of buffered solid 1/2 MS (see Recipes)
IAA 10 mM stock (see Recipes)
IAA 100 μM solution (see Recipes)
Fluorescent tracer (to follow treatment arrival in microfluidics) (see Recipes)
Specific to agar media experiments
Round 9 cm diameter Petri dishes (e.g., SterilinTM, Thermo Fisher, catalog number: 101R20)
NuncTM Lab-TekTM II microscopy chambers (VWR, catalog number: 734-2055) or 3D printed (see https://cellgrowth-lab.weebly.com/3d-prints.html)
Specific to microfluidic experiments
Microfluidic chips suited for A. thaliana root imaging (e.g., Grossmann et al., 2011; Fendrych et al., 2018; Serre et al., 2021; for review, see Yanagisawa et al., 2021)
Filtering system (Nalgene, P-Lab, N325045)
0.45 μm pore size 47 mm diameter round filtering membrane (Macherey-Nagel, P-Lab, M680447)
Equipment
Precision scale (any brand)
Vortex bench mixer (any brand)
Sterile laminar flow hood (any brand)
Lab scale (any brand)
Thin tweezers (Dumont No.5, Youlab, F003)
Autoclave (any brand)
pH meter (any brand)
Stirring magnetic rod and magnetic mixer (e.g., 2MAG, Dutscher, catalog number: 699010)
Autoclavable 0.5 L borosilicate glass bottles (e.g., SLS, Dutscher, catalog number: 257065)
500 mL graduated cylinder (any brand, plastic or glass)
1,000 mL beaker (any brand, plastic or glass)
Thin spatula (e.g., Heathrow, Dutscher, catalog number: 037874)
Growth room set in long days [e.g., 23 °C by day (16 h), 18 °C by night (8 h), 60% humidity, and light intensity of 120 μmol photons m-2·s-1]
Confocal microscope setup (see below)
Observation of the membrane potential with DISBAC2(3) staining requires a confocal laser or spinning disk microscope (with a 515 nm source of excitation). Here, we present an example based on our microscopy setup. Spinning disk confocal microscopy is particularly adapted to the imaging of fast responses in high spatio-temporal resolution, as the images are acquired under short exposure times. Moreover, the use of vertical inverted stage microscopy allows to keep the seedlings in their natural orientation (towards gravity) and thus avoid bias based on putting the seedlings on the horizontal plane (von Wangenheim et al., 2017).
Microscope body: Axio Observer 7 inverted (Zeiss); the microscope was set up on its back to obtain a vertical stage (details in von Wangenheim et al., 2017; Zeiss also provides custom mounting on request).
Spinning disk: CSU-W1-T2 spinning disk unit with 50 μm pinholes (Yokogawa)
Excitation source: VS-HOM1000 excitation light homogenizer (Visitron Systems)
Objective: Plan-Apochromat × 20/0.8 (Zeiss)
Detection: PRIME-95B Back-Illuminated sCMOS camera 1,200 × 1,200 pixels (Photometrics)
For microfluidic only: microfluidic pressure driven flow setup
We also describe a protocol to prepare media to observe membrane potential using microfluidics. There are a multitude of microfluidic (or perfusion) setups possible based on manufacturer, budget, and need. The same for the production of microfluidic chips. A detailed protocol describing the establishment of a microfluidic setup and the microfluidic chip production is thus out of scope of this protocol. For an example of microfluidic setup using manually closable and reusable chips, see https://www.elveflow.com/microfluidic-applications/microfluidic-cell-culture/microfluidic-microscopy-imaging/. Note that any microfluidic chip adapted to A. thaliana would work, either conventional glass coverslip–sealed microfluidic chips (e.g., Grossmann et al., 2011; Fendrych et al., 2018; for review, see Yanagisawa et al., 2021) or manually closed microfluidics chips (Serre et al., 2021).
Software
Microscopy imaging software (here, we used Visiview, Visitron, https://www.visitron.de)
ImageJ (v1.53t) Fiji (v2.9.0) (Schindelin et al., 2012)
LibreOffice Calc (https://www.libreoffice.org/) or Microsoft Office Excel (https://www.microsoft.com/fr-fr/microsoft-365/microsoft-office)
R (R Core Team, 2022, https://www.R-project.org/) with the nparcomp (Konietschke et al., 2015) package installed
R studio IDE (optional, https://posit.co/)
Python in the Anaconda distribution platform [tested with Python v3.8, https://www.anaconda.com/products/distribution)] with Spyder IDE (tested with v5.2.2, command: conda install spyder) and installed Seaborn package (tested with v0.11.2, command: conda install seaborn)
Optional: Inkscape
Procedure
Procedure to grow the seedlings for agar experiments and manually closed microfluidic chips
Sterilize A. thaliana Col-0 (or line of interest) seeds with the method of your choice (e.g., chlorine gas; Lindsey et al., 2017).
If the buffered solid 1/2 MS (see Recipes) is solidified, dissolve it completely in the microwave with the lid half unscrewed. If the medium has been kept at 60 °C after sterilization, proceed to the next step.
Critical: The following steps have to be carried out under a sterile laminar flow hood.
Pour 50 mL of buffered solid 1/2 MS into a square Petri dish and allow to solidify (approximately 15 min)
Place approximately 10–20 sterilized Col-0 seeds (or line of interest) in a row (approximately 3 cm from the top) using a sterilized toothpick (or method of your choice).
Close and seal the Petri dish with micropore tape.
Transfer the plate to 4 °C for two days for stratification.
Transfer the plate to the growth room for 4/5 days (primary root length between 2 and 4 cm). Note that seedlings need to fit the microscopy chamber.
Note: Given the short-term aspect of the observations carried out, the experiments are performed in a non-sterile environment.
Experiments with agar medium for the observation of stable membrane polarization states
A video demonstrating these steps is available in Video 1.
Video 1. Preparation of solid media in microscopy chambers for agar experiments
Prepare an IAA 100 μM solution (see Recipes).
Note: Steps are graphically represented in Figure 1 and demonstrated in Video 1.
Figure 1. Preparation of solid media in microscopy chambers for agar experiments
Pour 40 mL of buffered 1/2 MS pH 5.8 agar medium (see Recipes) into a 50 mL tube.
Wait a few minutes for the medium to cool down (until not burning in the hand).
Add 60 μL of DISBAC2(3) from a 2 mM aliquot to make the final concentration 3 μM.
Gently mix by inversion until the pink staining is uniform.
Pour 20 mL of this solution into a Petri dish.
Add 20 μL of IAA 100 μM into the leftover media in the tube to obtain a 100 nM final concentration (avoid agar temperature above 60 °C).
Gently mix by inversion.
Pour the IAA supplemented medium into another Petri dish.
Let the medium solidify (approximately 15 min).
In the small Petri dish, cut the agar to delimit a patch that would fit the microscopy chamber (using a thin spatula).
Transfer your seedlings to the control and treatment patches using a toothpick or tweezers to gently lift the seedlings by their cotyledons.
Transfer the patches with seedlings to the microscopy chamber (so that the seedlings are sandwiched between the agar and the chamber cover glass).
Caution: Avoid contact between both media in the chamber to prevent diffusion of IAA into the control medium.
Install the chamber on the microscope stage and let the plants recover from the transfer stress for at least 20 min (this minimum as been determined by the stage at which root elongation and fluorescence reach a stable plateau after the mechanical stress induced by the transfer).
Note: Waiting longer than 20 min does not harm the assay. During this time, you can set your microscope stage positions on your microscope software.
Once recovered, start imaging with a 515 nm excitation.
Note: Imaging settings will depend on the microscope setup and will need to be adapted by the user pre-experiment; do not change them between experiments/replicates to be able to compare control and treated plants. We routinely image plants every 10 min for 40 min to quantify root elongation as well.
Microfluidic experiments to observe fast depolarization/hyperpolarization events
A video demonstrating these steps is available in Video 2.
Video 2. Preparation of liquid media for microfluidics experiments
Prepare an IAA 100 μM solution (see Recipes).
Note: Steps are graphically represented in Figure 2 and demonstrated in Video 2.
Figure 2. Preparation of liquid media for microfluidics experiments
Filter liquid buffered 1/2 MS media pH 5.8 (see Recipes) using, for example, a 0.45 μm membrane in a siphon filter.
Add 0.1 g of sucrose to an empty 15 mL tube.
Add 10 mL of the filtered media (from step C2) to the same tube.
Vortex and/or shake to fully dissolve the sucrose into the medium.
Add 10 μL of DISBAC2(3) (from a 2 mM aliquot in DMSO) to obtain a final concentration of 2 μM.
Vortex and/or shake to dissolve.
Transfer 5 mL to another 15 mL tube.
Critical: Steps C3–C8 ensure that the dye and sucrose are at the exact same concentrations in both tubes. DISBAC2(3) is used for relative measurements and thus every step counts to increase reproducibility.
Add 5 μL of buffered 1/2 MS media pH 5.8 in one tube (control tube) and 5 μL of IAA 100 μM in a second tube (treatment tube).
Continue with the procedure adapted to your microfluidic setup.
If you are using a manually closable microfluidic chip in which you directly transfer 4–5-day-old seedlings, let the seedlings recover for at least 20 min with control media flow after closing the device.
Note: During this time, you can set your microscope stage positions on your microscope software.
Start imaging with a 515 nm excitation.
After at least 5 min, switch the flow in the chip from control media to treatment media.
Note: Imaging settings depend on the microscope setup and must be adjusted by the user prior to the experiment. They must not change between experiments/replicates, to allow comparison between control and treated plants. We routinely image the plants every 30 s to observe rapid responses. For example: 5 min control condition/12 min treatment condition.
Data analysis
In this protocol, we use the imaging of the rapidly induced IAA root depolarization to showcase DISBAC2(3) as a relative membrane potential reporter. We noticed that the depolarization was the strongest above the root cap, in the root transition zone (Serre et al., 2021), a zone involved in many root responses to stresses (Baluška et al., 2010). We then carried out measuring the DISBAC2(3) fluorescence only in the root transition zone. Moreover, we focus on the interface between the cortex and the epidermis, as the dead lateral root cap cells were strongly fluorescent (open membranes for the dye to react to). The data presented here are re-analyzed images from Serre et al. (2021). Raw data can be found on Zenodo (https://zenodo.org/record/4922659). All the scripts used in this protocol can be found on the public repository https://sourceforge.net/projects/disbac2-3-data-analysis/.
Here, we describe a method to:
Quantify DISBAC2(3) fluorescence in the transition zone at a given point (agar experiment) or over time (microfluidics) using the ImageJ/Fiji software.
Quantify root elongation either as an average growth (agar experiment) or over time (microfluidics).
Normalize the microfluidics measured data in Calc or Excel.
Represent the data either with R or Python and the Seaborn plugin.
Carry non-parametric statistical analysis using the nparcomp package in R.
Note: This process would be hard to automate and, therefore, can be subjective. To avoid unconscious measuring bias, we recommend measuring the fluorescence blindly (not knowing which conditions are measured). For example, ask a colleague to change the name of your files, keeping the original names associated with the blind identifiers.
Analysis of experiments carried out in agar media
Note: Membrane potential is only measured on the first timeframe of the stack, as we are looking for a well-established response after 25 min of treatment. Root elongation is calculated as an average of 40 min of growth.
Image analysis
Fluorescence measurements:
Open one microscopy image stack in ImageJ/Fiji.
Check if scale settings are correct (Menu: Analyze > Set Scale).
Set measurements to at least Mean gray value (Menu: Analyze > Set Measurements).
Select the segmented line tool (right-click on the default straight line).
Increase line width of the segmented line tool to 20 pixels (double-click on the segmented line tool).
Display the first timeframe of the image stack.
Adjust brightness to comfortably see the root transition zone cells (shortcut Control + Shift + C or in the menu: Image > Adjust > Brightness/Contrast, Figure 3a).
Figure 3. Measurements and plotting in solid media experiments. (a) Example of images with the gray values represented as a LUT. (b) Selection of the transition zone at the intersection between cortex and epidermis. (c) Measurements of root elongation. (d) Measured membrane potential fluorescence and (e) root elongation, values of Col-0 treatment with 0 or 100 nM IAA. Scale bar: 50 μm. For (d) and (e), the * represents a p-value < 0.05, obtained by a non-parametric Student test (npar.t.test function in the R nparcomp package).
Optional: Set a LUT to display intensities as color gradients (Main bar: Lut, e.g., Green Fire blue or Fire)
Draw a segmented line on the transition zone cells on the right side of the root, centered on the membrane between root cortex and epidermis (Figure 3b, left-click to add segment, right-click to end segmented line selection).
Note: For a definition of transition zone location, see Baluška et al. (2010).
Make sure that the result table window is not open or is empty.
Measure (shortcut m or menu: Analyze > Measure).
Repeat the process from e to j for the left side of the root to measure the fluorescence (Figure 3b).
Measure (shortcut m or menu: Analyze > Measure).
Repeat process for all the roots in one condition.
Copy the measured Mean (mean fluorescence of the selection) from the result table window to an Excel spreadsheet or similar software.
Clear result table (either in result window: Results > Clear Results or by closing the window).
Root elongation measurements:
Be sure to be on the first timeframe of the image stack and that the segmented line tool is selected.
Select the extreme tip of the root (left-click).
Go to last timeframe (scroll down or image slide bar).
Select the extreme tip of the root (right-click to end selection) (Figure 3c).
Measure (shortcut m or menu: Analyze > Measure).
Repeat process for all the roots in one condition.
Copy the measured Length from the result table window to an Excel spreadsheet or similar software.
Clear result table (either in result window: Results > Clear Results or by closing the window).
Data analysis
Organize the measured data to fit Table 1 (can be in the same file, different sheets).
Table 1. Characteristics of table with measured membrane potential fluorescence and root elongation for plotting and statistics
Filename Condition Fluorescence TZ right Fluorescence TZ left Average fluorescence Root length increment (here 2 h) Root elongation in μm/min (increment/120)
0nM_root1.tif 0nM_IAA 381.715 345.623 363.669 164.622 1.37185
0nM_root2.tif 0nM_IAA 447.295 433.388 440.3415 405.015 3.375125
0nM_root3.tif 0nM_IAA 343.961 436.011 389.986 435.681 3.630675
0nM_root4.tif 0nM_IAA 392.971 481.415 437.193 456.521 3.804341667
0nM_root5.tif 0nM_IAA 488.14 423.733 455.9365 426.512 3.554266667
0nM_root6.tif 0nM_IAA 371.998 367.106 369.552 345.512 2.879266667
0nM_root7.tif 0nM_IAA 546.02 392.141 469.0805 294.079 2.450658333
100nM_root1.tif 100nM_IAA 490.262 627.423 558.8425 122.673 1.022275
100nM_root2.tif 100nM_IAA 585.199 444.535 514.867 84.957 0.707975
100nM_root3.tif 100nM_IAA 538.38 497.549 517.9645 91.505 0.762541667
100nM_root4.tif 100nM_IAA 414.893 454.586 434.7395 101.832 0.8486
100nM_root5.tif 100nM_IAA 501.4 498.181 499.7905 148.582 1.238183333
100nM_root6.tif 100nM_IAA 552.743 614.447 583.595 85.581 0.713175
100nM_root7.tif 100nM_IAA 517.162 718.587 617.8745 113.049 0.942075
Open R software (or R Studio IDE).
Open the R script supplemented. For comparisons and plotting of two conditions, use “R_script-agar_2conditionsgenotypes.R.” For more than two conditions, use: “R_script-agar_3andplusconditionsgenotypes.R.”
Follow the script comments to modify the script to your conditions.
Execute Script.
Save boxplot as svg (for possible aesthetical modifications in Inkscape, Figure 3d–3e).
Save output of the statistics by copy and pasting into a notepad.
Analysis of experiments carried out in microfluidics
In microfluidics experiments, membrane potential is measured over time to have access to the dynamic response after treatment. We also propose a way to measure root elongation rates over time.
Image analysis for one condition (e.g., control to 100 nM IAA switch)
Determining the timeframe associated with the treatment arrival:
In our imaging conditions, we observed a small drop in background fluorescence during medium switches. This allowed us to determine which timeframe should be considered as time zero for the treatment. This prevents the use of fluorescent tracers such as Fluorescein dextran (see Recipes if a tracer is needed in your experimental conditions).
Open one microscopy image stack in ImageJ/Fiji.
Check if scale settings are correct (Menu: Analyze > Set Scale).
Set measurements to at least Mean gray value (Menu: Analyze > Set Measurements).
Select a region of interest (ROI) in the background as far as possible from the root (Figure 4a).
Figure 4. Measurements and plotting microfluidic experiments. (a) Selection of a background region of interest to determine the treatment arrival. (b) Rectangular selection of the root tip to determine root elongation rates over time. (c) Measured and normalized DISBAC2(3) fluorescence reflecting the membrane potential and (d) root elongation; values of Col-0 observed 5 min in control and then 10 min with IAA 100 nM. For (c) and (d), negative values represent the root being exposed to the control medium, and positive values represent the root being exposed to the treatment medium.
Add this ROI to the ROI manager (Shortcut Ctrl + t or Analyze/Tools/ROI Manager).
Make sure that the result table window is not open or is empty.
Select the added selection in the ROI manager and click on more >> Multi Measure. A window will pop up; ensure that the options Measure all xx slices and One row per slice are ticked.
Then, click OK.
Copy and paste the results from the result table into an Excel table.
Note: The arrival of the treatment is marked by either a clear drop in background fluorescence or a small but sharp increase, followed by a drop in background fluorescence. If you have trouble determining the treatment arrival in your conditions, see Recipes for fluorescent tracer.
Stabilization of the root on the image stack:
Select the segmented line tool (right-click on the default straight line).
Increase line width of the segmented line tool to 20 pixels (double-click on the segmented line tool).
Display the first timeframe of the image stack.
Adjust brightness to comfortably see the root transition zone cells (shortcut Control + Shift + C or in the menu: Image > Adjust > Brightness/Contrast).
Optional: Set a LUT to display intensities as color gradients (Main bar: Lut, e.g., Green Fire blue or Fire).
Draw a segmented line on the transition zone cells on the right side of the root, centered on the membrane between root cortex and epidermis (as in Figure 3b, left-click to add segment, right-click to end segmented line selection).
Start the Correct 3D drift plugin (Plugins/Registration/Correct 3D drift).
In the window, select the channel for registration (the fluorescence channel works well, but we also had great results with brightfield), and then click OK.
Add your transition zone ROIs (left and right sides) to the registered stack into the ROI Manager and check that the selection is always on the transition zone over time.
Note: If the selections are not stable over time, try the registration with the Sub pixel drift correction, and/or use the other side of the root and/or register on one side and then the other.
Make sure that the result table window is not open or is empty.
Once the ROIs are stable over time, click more >>/Multi measure in the ROI Manager window.
Copy and paste the results from the result table into an Excel table (only keeping the Mean1 and Mean2 columns.
Repeat the process of 1a and 1b for all your image stacks in control and treatment condition(s).
Root elongation measurements:
Go back to the original image stack (not registered).
Make a rectangular selection on the root tip (Figure 4b).
Open the Correct 3D drift plugin (Plugins/Registration/Correct 3D drift).
In the window, select the channel for registration and then tick Only compute drift vectors?
Rename the measure text file with the name of your raw image stack (or desired name) and save it.
Data analysis
Fluorescence analysis:
Organize the measured data to fit Table 2 (the name of the column Time is essential for the script to recognize the column).
Open the python script fluorescence_analysis.py in the Spyder IDE.
Modify the user inputs at the beginning of the code and run.
The script will import, normalize, and plot your data (Figure 4c).
Note: The normalization process is explained in the Materials and Methods of Serre et al. (2021).
Table 2. Characteristics of table with measured membrane potential fluorescence for analysis. Negative times represent the control window, zero is the timeframe of the media switch, and positive times represent the treatment window.
Time (min) Root1 Root2 Root3 Root4 Root5 Root6 Root7 Root8
-5 408.908372 389.499513 392.495975 327.607 411.428 395.879762 403.292463 411.977079
-4.5 405.668564 386.243762 393.183715 327.672 413.428 397.335688 404.113862 413.522185
-4 403.082014 386.306109 392.140659 332.045 413.523 395.057337 404.527451 417.715058
-3.5 396.392234 385.716733 390.918841 333.657 414.2265 396.265252 402.175133 418.270171
-3 395.523104 380.711613 392.716895 335.763 416.6785 396.086036 401.628771 419.469708
-2.5 397.419366 381.983792 396.580456 339.207 420.0695 396.374816 412.618216 423.596701
-2 392.097153 381.825249 399.140425 342.854 418.4595 396.809199 405.392976 420.864414
-1.5 399.883876 379.197157 401.737308 347.443 418.183 399.659632 399.678631 421.315916
-1 396.454276 377.647971 403.093227 347.078 418.119 399.218287 392.094359 419.257999
-0.5 398.013425 378.172317 403.254221 349.409 418.368 402.997758 404.306832 423.040022
0 400.452519 382.279517 407.598904 349.563 418.121 397.010205 394.462017 422.650659
0.5 419.649339 390.901924 441.873124 358.037 434.2055 415.260263 439.403187 431.782716
1 442.17854 416.22474 475.150674 380.477 463.097 442.370851 490.670599 458.766988
… … … … … … … … …
9.5 398.475274 452.643626 466.316994 405.541 491.053 528.389269 360.38102 406.706359
10 399.328873 452.295109 467.085197 395.029 486.823 534.563718 344.475839 403.269758
Root elongation analysis:
Regroup all the shifts text files into a dedicated folder.
Retrieve scale (in pixels/μm) from one of your stacks of images (Fiji > Analyze > Set Scale).
Open the python script elongation_compilation.py in the Spyder IDE.
Modify the user inputs at the beginning of the code and run the script to create a table with all your root elongation rates calculated in μm/min at each time point.
Note: This script takes the x and y coordinates shifts over time and 1) calculates the distance between the coordinates time +1 and time +0, 2) transform those coordinates in μm using the user scale, and 3) divides the data by the duration of the timeframes in minutes to obtain μm/min.
Open the saved table into an Excel table and manually add the Time (name is essential) column as a first column to resemble Table 2.
Open the python script elongation_plotting.py in the Spyder IDE.
Modify the user inputs at the beginning of the code and run the script to create the line plot (Figure 4d).
Note: For plotting more than two conditions in microfluidics (e.g., control to 100 nM IAA and control to 1,000 nM IAA), the scripts fluorescence_analysis.py and elongation_compilation.py should be run on the individual conditions. We then provide another python script: plotting_fluo_elongation_2andmore.py.
Validation of protocol
The development and validation of this method is described in detail in Serre et al. (2021).
Notes
Reproducibility
The DISBAC2(3) staining method can be very reproducible if the user 1) respects the dye dilution steps and 2) minimizes mechanical stress during transfer and 3) the recovery time before imaging.
Particularities we observed
Dead cells shine considerably more than living cells, as the dye is able to freely stain membranes inside and outside (e.g., dead root cap cells).
The outer cells (epidermis/cortex) fluoresce more, but observations by multi-photons microscopy revealed that the dye actually penetrates the whole root (Serre et al., 2021).
DISBAC2(3) tends to stick to microfluidics tubing (such as Tygon or PTFE), especially when the flow is stopped. We observed lower fluorescence intensities switching from one control medium to another control medium. This emphasizes the need for proper controls and the relative aspect of this method.
Rapidity of response
Depolarizations can be very fast and transient and might not be visible after a few minutes [e.g., agar imaging with low concentration of IAA (< 100 nM)]. The IAA-induced depolarization is statistically still visible in agar after 25 min at strong concentrations above 100 nM.
Vertical microscopy
If using a vertical microscope, always keep your seedlings oriented toward the gravity (while growing, moving the plate, on the bench, during transfer to agar patches, and in the microscopy chamber). This will prevent auxin fluxes from being triggered, which could bias measurements.
Recipes
DISBAC2(3) master stock [for a 100 mg bottle of DISBAC2(3) powder]
Add 2.290 mL of DMSO directly into the bottle (final concentration of 100 mM).
Vortex until fully dissolved.
Keep at -20 °C.
DISBAC2(3) working aliquots
Add 392 μL of DMSO into a 1.5 mL tube.
Add 8 μL of DISBAC2(3) master stock at 100 mM (Recipe 1) to obtain a 2 mM final concentration.
Vortex.
Aliquot in 20 μL in 500 μL tubes.
Keep at -20 °C.
70% ethanol (approximately) prepared from 96% ethanol
Pour 175 mL of 96% ethanol into a graduated cylinder.
Pour distilled water up to 250 mL.
Keep at room temperature.
500 mL of buffered liquid 1/2 MS
In a 500 mL beaker, weigh and add:
1.1 g of Murashige and Skoog mix
0.3 g of MES hydrate
Add approximately 450 mL of Milli-Q or distilled water to the beaker.
Add magnetic rod and mix with a magnetic mixer until all powders are dissolved.
Raise pH to 5.8 with 1 M KOH using a pH meter.
Remove magnetic rod and transfer to a 500 mL graduated cylinder.
Add water up to 500 mL.
Filter in a 0.5 L glass bottle using a siphon filter or a syringe filter.
Keep at room temperature.
500 mL of buffered solid 1/2 MS
In a 500 mL beaker, weigh and add:
1.1 g of Murashige and Skoog mix
5 g of sucrose (final concentration of 1% m/v)
0.3 g of MES hydrate
Add approximately 450 mL of Milli-Q or distilled water to the beaker.
Add magnetic rod and mix with a magnetic mixer until all powders are dissolved.
Raise pH to 5.8 with 1 M KOH using a pH meter.
Remove magnetic rod and transfer to a 500 mL graduated cylinder.
Add water up to 500 mL.
Add 4 g of plant agar to a 0.5 L glass bottle.
Add the medium from step 5e.
Autoclave with cap half unscrewed.
Keep bottle in a 50/60 °C incubator to keep it liquid, use directly after autoclave, or let solidify and dissolve agar in microwave before use (up to two times).
IAA 10 mM stock
Let the IAA powder bottle come to room temperature (30 min on the bench).
Using a precision scale, weigh 17.518 mg of powder and add to a 15 mL tube.
Add 10 mL of 70 % ethanol (Recipe 3) in the 15 mL tube (final concentration of 10 mM).
Vortex until dissolved.
Cover the tube with aluminum foil and store at -20 °C for up to one month.
IAA 100 μM solution
Add 990 μL of liquid 1/2 MS (Recipe 4) into a 1.5 mL tube.
Add 10 μL of IAA 10 mM (Recipe 6) to obtain a final concentration of 100 μM.
Vortex.
Discard after one day at room temperature.
Fluorescent tracer (to follow treatment arrival in microfluidics)
Weigh 10 mg of Fluorescein Dextran 10 K MW into a 1.5 mL tube.
Add 1 mL of distilled water for a 10 mg/mL final concentration.
Vortex until fully dissolved.
Add 0.6 μL per mL of control liquid medium.
Add 0.3 μL per mL of treatment liquid medium.
The fluorescent tracer is visible with a 515 and 488 nm excitation source.
Caution: We advise to alternate the two concentrations in the control/treatment media to be sure that the observed effects are not from the tracer. It is important to add the tracer in two different concentrations in both media to minimize potential effects of the dye, as minimal but visible effects on root elongation can be observed.
Acknowledgments
This work was supported by the European Research Council (grant no. 803048). This protocol is a detailed version of the material and methods and reproduces results published in Serre et al. (2021).
Competing interests
The authors declare no competing interests.
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Serre, N. B. C., Kralík, D., Yun, P., Slouka, Z., Shabala, S. and Fendrych, M. (2021). AFB1 controls rapid auxin signalling through membrane depolarization in Arabidopsis thaliana root. Nat. Plants 7(9): 1229–1238.
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Plant Science > Plant cell biology > Cell imaging
Plant Science > Plant developmental biology > General
Cell Biology > Cell imaging > Live-cell imaging
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Production and Purification of Cell Culture–generated Hepatitis B Virus by Transient Transfection and Density Gradient
AM Asako Murayama
HA Hirofumi Akari
TK Takanobu Kato
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4779 Views: 517
Reviewed by: Saskia F. Erttmann Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Sep 2022
Abstract
An efficient cell culture system for hepatitis B virus (HBV) is indispensable for research on viral characteristics and antiviral agents. Currently, for HBV infection assays in cell culture, HBV genome-integrated cell line–derived viruses are commonly used. However, these viruses are not suitable for the evaluation of polymorphism-dependent viral characteristics or resistant mutations against anti-viral agents. To detect the infection of cell culture–generated HBV (HBVcc) by the transient transfection of the HBV molecular clone, a large amount of purified viruses is needed, because such viruses exhibit limited infection efficiencies in cell culture. Here, we describe how to generate and purify HBVcc by the transient transfection of HBV molecular clones. This system provides a powerful tool for studying the infection and propagation of HBV and for developing anti-viral agents against HBV.
Keywords: HBV Cell culture Molecular clone NTCP Virus generation Virus purification
Background
Hepatitis B virus (HBV) infection is a significant cause of chronic liver disease, including cirrhosis and hepatocellular carcinoma. Although effective vaccines against HBV infection are available in many countries, the global prevalence of HBV is estimated to be over 290 million. The eradication of chronic HBV infection by the current treatment strategy is not expected, because HBV covalently closed circular DNA in hepatocytes cannot be eliminated. To explore novel anti-HBV agents, a system for the infection and replication of HBV in cell culture is indispensable. Sodium taurocholate cotransporting polypeptide (NTCP) was identified as an HBV receptor, and NTCP-transduced HepG2 or HuH-7 cells contributed to the observation of HBV infection and replication in cell culture (König et al., 2019; Otoguro et al., 2020). In such cell culture systems for HBV, viruses derived from HBV genotype D genome-integrated cell lines, such as HepG2.2.15 or HepAD-38, are used. However, such viruses are not suitable for investigating the effects of strain-specific characteristics or resistance-related polymorphisms on the effectiveness of anti-HBV agents. For these purposes, the viruses obtained by the transient transfection of the HBV molecular clone are used. However, these viruses have limited infection efficiencies in cell culture. Therefore, methods for the efficient production of cell culture–generated HBV (HBVcc) and for the purification of infectious viruses will be important (Washizaki et al., 2022).
Materials and reagents
Production of HBVcc
HBV molecular clone plasmid
A replication-competent HBV molecular clone with a 1.38-fold genome length of HBV genotype C strain (accession number: AB246345) (Figure 1) (Murayama et al., 2021) inserted into the pUC19 plasmid is prepared. This clone is introduced with a precore stop mutation (G1896A) to evaluate the production of HBc proteins by measuring the HBcrAg level excluding HBeAg. HBeAg is not produced from this construct.
Figure 1. Structure of the hepatitis B virus (HBV) molecular clone for cell culture-generated HBV (HBVcc). Structure of the plasmid of the HBV molecular clone. This plasmid contains a 1.38-fold HBV genome (4,438 bp) of the HBV genotype C strain. This construct generates pregenomic RNA and expresses all HBV proteins.
Sodium taurocholate cotransporting polypeptide (NTCP)-transduced cells
HepG2-NTCPsec+ (provided by Dr. Marc Peter Windisch; Institut Pasteur Korea, Seoul, South Korea) (König et al., 2019). These cells were cultured with HepG2-NTCPsec+ culture medium (see Recipes) supplemented with Blasticidin (5µ g/mL)
Dulbecco’s modified Eagle medium (DMEM) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 044-29765)
Fetal bovine serum (FBS) (Sigma-Aldrich, catalog number: F7524)
MEM non-essential amino acids solution (100×) (NEAA) (Thermo Fisher Scientific, Gibco, catalog number: 11140050)
HEPES (1 M) (Thermo Fisher Scientific, Gibco, catalog number: 15630130)
Sodium pyruvate (100 mM) (Thermo Fisher Scientific, Gibco, catalog number: 11360070)
Penicillin-streptomycin (10,000 U/mL) (Thermo Fisher Scientific, Gibco, catalog number: 15140122)
Blasticidin S hydrochloride, HEPES solution (Blasticidin) (10 mg/mL) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 022-18713)
Opti-MEM I reduced serum medium (Thermo Fisher Scientific, Gibco, catalog number: 31985070)
Lipofectamine 3000 transfection reagent (Thermo Fisher Scientific, Invitrogen, catalog number: L3000015)
Collagen-coated T225 flask (Corning, catalog number: NCO431082)
Dulbecco’s phosphate buffered saline (PBS) (Nacalai Tesque, catalog number: 14249-24)
Dimethyl sulfoxide (DMSO) (Nacalai Tesque, catalog number: 09659-14)
Syringe-top filter unit; Millex-HV Syringe Filter Unit, 0.45 μm, PVDF, 33 mm (Merck Millipore, catalog number: SLHVR33RS)
Amicon Ultra-15 centrifugal filter units (100 kDa) (Merck Millipore, catalog number: UFC910096)
Purification of HBVcc
Optiprep (60% iodixanol solution) (Serumwerk Bernburg, catalog number: AXS-1114542)
Open-top thinwall ultra-clear tube (14 mm × 89 mm) (Beckman Coulter, catalog number: 344059)
DNase; RQ1 RNase-free DNase (Promega, catalog number: M6101)
QIAamp DNA Mini kit (Qiagen, catalog number: 51306)
Lumipulse G HBsAg-Quant (Fujirebio, catalog number: 296851)
Lumipulse G HBcrAg (Fujirebio, catalog number: 294109)
Luna Universal qPCR Master Mix (New England Biolabs, catalog number: M3003)
Primers and probe for the real-time PCR targeting the HBs region:
Forward primer: 5′-CTTCATCCTGCTGCTATGCCT-3′
Reverse primer: 5′-AAAGCCCAGGATGATGGGAT-3′
Probe: 5′-FAM-ATGTTGCCCGTTTGTCCTCTAATTCCA-TAMRA-3′
TNE buffer (see Recipes)
Titration of HBVcc
Collagen-coated 96-well culture plate (Corning, catalog number: NCO3585)
Polyethylene glycol average mol wt 8,000 (PEG8000) (Sigma-Aldrich, catalog number: P2139)
4% paraformaldehyde phosphate buffer solution (4% PFA) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 163-20145)
Block ACE (Bio-Rad, catalog number: BUF029)
Triton X-100 detergent (Calbiochem, Merck Millipore, catalog number: 648466)
Anti-HBc antibody; anti-hepatitis B virus core antigen IgG fraction (polyclonal) (AUSTRAL Biologicals, catalog number: HBP-023-9)
Goat anti-rabbit IgG (H+L) highly cross-adsorbed secondary antibody, Alexa Fluor Plus 555 (Thermo Fisher Scientific, catalog number: A327320
DAPI (4’,6-Diamidino-2-phenylindole dihydrochloride) (FUJIFILM Wako Pure Chemical Corporation, catalog number: 342-07431)
4% PEG8000 (see Recipes)
Recipes
HepG2-NTCPsec+ culture medium
DMEM
10% FBS
Penicillin-streptomycin (100 U/mL)
NEAA (1×)
HEPES (10 mM)
Sodium pyruvate (1 mM)
TNE buffer
150 mM NaCl
10 mM Tris HCl (pH 7.5)
1 mM EDTA
4% PEG8000
Dissolve in water to 4% (w/v) in a water bath at 60 °C.
Equipment
Refrigerated centrifuge with swing rotor (TOMY, model: AX-310)
Automated chemiluminescent enzyme immunoassay system (Fujirebio, model: LUMIPULSE G1200)
Ultracentrifuge apparatus, Optima L-90K with an SW-41 Ti rotor (Beckman Coulter, model: Optima L-90K)
Analytical balance (Sartorius Lab Instruments, model: SECURA124-1SJP)
Real-time PCR system (Thermo Fisher Scientific, model: StepOnePlus Real-Time PCR System)
Fluorescence microscope (KEYENCE, model: BZ-X710)
Procedure
Production of HBVcc
Detach and count HepG2-NTCPsec+ cells.
Place the cells (5 × 107 cells resuspended with 2–5 mL HepG2-NTCPsec+ culture medium) into a 50 mL conical tube.
Prepare the transfection mixture in 50 mL conical tubes (Table 1).
Table 1. Composition of transfection reagent
Reagent Amount
Mixture A
Opti-MEM I reduced serum medium 2 mL
HBV molecular clone plasmid (2 μg/μL) 40 μL
P3000 enhancer reagent (included in Lipofectamine 3000 Transfection Reagent kit) 80 μL
Mixture B
Opti-MEM I reduced serum medium 2 mL
Lipofectamine 3000 reagent 80 μL
Mix the prepared mixtures A and B and incubate for 5 min following the manufacturer’s protocol.
Transfer the generated mixture into HepG2-NTCPsec+ cells in the 50 mL conical tube and mix well by vortex.
Seed the transfected cells into a collagen-coated T225 flask with 45 mL of HepG2-NTCPsec+ culture medium.
One day after transfection, wash the transfected and seeded cells with 12 mL of PBS three times and add 45 mL of HepG2-NTCPsec+ culture medium with 2% DMSO. The addition of DMSO improves the yield of HBVcc.
Culture the transfected cells for an additional seven days, monitoring HBsAg levels in the culture medium by the automated chemiluminescent enzyme immunoassay system (Murayama et al., 2019) (Figure 2).
Figure 2. Production of HBsAg in the culture medium of hepatitis B virus (HBV) molecular clone-transfected cells. HBsAg levels in the culture medium of the HBV molecular clone-transfected cells were monitored. A time-dependent increase was observed.
Eight days after transfection, harvest the culture medium into a 50 mL conical tube and centrifuge (2,380× g, 5 min, 4 °C) to precipitate the cell debris.
Pass the culture medium through a 0.45 μm syringe-top filter unit to remove cell debris.
Concentrate the culture medium approximately 30-fold by centrifugation (2,380× g, 90 min, 4 °C) with Amicon Ultra-15 centrifugal filter units (100 kDa) (Table 2). If the recovery rate is quite low, confirm that the centrifugal filter unit is intact, and the condition of the centrifugation is appropriate. The generated HBVcc in the culture medium can be stored at -80 °C before or after concentration.
Table 2. Concentration of HBVcc in culture medium by using two centrifugal filter units
Before After
Volume 30 mL 1 mL
HBsAg 86.3 IU/mL 2,190 IU/mL
HBsAg recovery rate - 84.6%
HBV DNA 4.1 × 109 copies/mL 7.2 × 1010 copies/mL
HBV DNA recovery rate - 58.5%
Purification of HBVcc
Dilute 60% iodixanol (Optiprep) to 40%, 30%, 20%, and 10% with TNE buffer (Table 3).
Table 3. Composition of mixtures of iodixanol and TNE buffer
Iodixanol concentration Optiprep TNE buffer Total volume
40% 4 mL 2 mL 6 mL
30% 3 mL 3 mL 6 mL
20% 2 mL 4 mL 6 mL
10% 1 mL 5 mL 6 mL
Prepare a stepwise iodixanol density gradient by layering the 40%–10% iodixanol solutions (2 mL each) in the ultracentrifuge tube (open-top thinwall ultra-clear tube) (Figure 3).
Figure 3. Preparation of stepwise iodixanol density gradient. Two milliliters each of the 40%–10% iodixanol solutions were layered. After that, the concentrated culture medium was put at the top of the iodixanol density gradient.
Layer the concentrated culture medium (2–3 mL) on top of the 10%–40% iodixanol gradient.
Centrifuge the culture medium on the gradient at 178,000× g for 16 h at 4 °C in an ultracentrifuge apparatus with an SW-41 Ti rotor.
After centrifugation, collect 500 μL of 20–22 fractions, depending on the volume of the layered concentrated culture medium, from the top of the density gradient. The collected HBVcc in the fractions can be stored at -80 °C.
Take a 100 μL aliquot of each fraction and measure the weight of the 100 μL aliquot using the analytical balance to determine the density of each fraction.
Dilute the collected fraction 200-fold with HepG2-NTCPsec+ culture medium and measure the HBsAg and HBcrAg levels for each fraction by the automated chemiluminescent enzyme immunoassay system as described in the manufacturer’s protocol (Figure 4).
Figure 4. Profile of cell culture–generated hepatitis B virus (HBVcc) in an iodixanol density gradient. HBVcc in the culture medium was applied to an iodixanol density gradient, the titers of HBsAg and HBcrAg were measured in each fraction, and the HBV DNA titer was quantified by real-time PCR.
Treat the 50 μL of diluted fraction with DNase (RQ1 RNase-free DNase) at a concentration of 50 unit/mL, 37 °C, 1 h. Extract the total DNA using a QIAamp DNA Mini kit following the manufacturer’s protocol.
Measure the HBV DNA levels by real-time PCR targeting the HBs region by using the plasmid containing the PCR fragment amplified by primers for the real-time PCR as the standard (Tables 4 and 5) (Figure 4) (Honda et al., 2021).
Table 4. Composition of real-time PCR
Reagent Concentration Volume
Luna Universal qPCR Master Mix 2× 12.5 μL
Forward primer 10 μM 0.5 μL
Reverse primer 10 μM 0.5 μL
Probe 15 μM 0.5 μL
H2O - 6 μL
Extracted DNA - 5 μL
Table 5. Program for real-time PCR
Step Temperature Time Data collection
1 50 °C 2 min off
2 95 °C 10 min off
3 95 °C 20 s off
4 60 °C 1 min on
Repeat steps 3 and 4 for 40 cycles
Titration of HBVcc
Seed HepG2-NTCPsec+ cells in collagen-coated 96-well culture plates at a density of 3 × 104 cells/well in 100 μL of HepG2-NTCPsec+ culture medium supplemented with DMSO (2%).
Prepare the inoculum with an aliquot of each fraction (Table 6) and infect into HepG2-NTCPsec+ cells one day after seeding by replacing the culture medium with the inoculum.
Table 6. Composition of inoculum with purified HBVcc
Amount (per well)
Fraction 5 μL
HepG2-NTCPsec+ culture medium 39 μL
DMSO 1 μL
40% PEG8000 5 μL
One day after infection, wash the infected cells with 100 μL of PBS three times and add 100 μL of HepG2-NTCPsec+ culture medium containing 2% DMSO.
Culture the infected cells by changing the culture medium containing 2% DMSO every three or four days.
Collect culture medium 12 days after infection and measure the HBsAg level as described in step A7 (Figure 5A). In this case, fraction 15 was the fraction with the peak infectivity.
After washing with 100 μL of PBS twice, fix the infected cells 12 days after infection by replacing the culture medium with 4% paraformaldehyde in PBS (4% PFA) for 30 min at room temperature (RT).
After washing with 100 μL of PBS twice, block and permeabilize the fixed cells using Block ACE with 0.3% Triton X-100 for 1 h at RT.
Stain the infected cells with anti-HBc antibody (1 μg/mL diluted with PBS) for 1 h at RT.
After washing with 100 μL of PBS three times, stain the infected cells with Alexa Fluor 555-conjugated anti-rabbit IgG (1 μg/mL diluted with PBS) for 1 h at RT in the dark.
After washing with 100 μL of PBS three times, stain the nuclei with DAPI (1 μg/mL diluted with PBS) (Figure 5B).
Figure 5. Determination of infectivity of purified cell culture–generated hepatitis B virus (HBVcc). (A) Purified HBVcc was used to infect HepG2-NTCPsec+ cells, and the HBsAg levels in the culture medium at 12 days after infection were measured. (B) The infected cells were fixed and visualized with anti-HBc antibody, and nuclei were visualized by DAPI.
To confirm the infectivity of generated viruses in the peak fraction, infect the HBVcc in the peak fraction of infectivity (fraction 15) at concentrations of one genome equivalent (GEq)/cell (3 × 104 copies/well), 10 GEq/cell (3 × 105 copies/well), 100 GEq/cell (3 × 106 copies/well), and 1,000 GEq/cell (3 × 107 copies/well), following the inoculum composition (Table 6).
Measure the HBsAg levels in the culture medium of the infected cells and stain the infected cells with anti-HBc antibody 12 days after infection (see steps A7 and C6–C10) (Figure 6).
Figure 6. Determination of the infectivity titer of purified cell culture–generated hepatitis B virus (HBVcc). (A) HBVcc in the peak fraction of infectivity was used to infect HepG2-NTCPsec+ cells at concentrations of 1, 10, 100, and 1,000 genome equivalent (GEq)/cell. The HBsAg level in the culture medium at 12 days after infection was measured. (B) The infected cells were fixed and visualized with anti-HBc antibody, and nuclei were visualized by DAPI.
Notes
The HBVcc should be handled according to the regulation of infectious agents. HBV is infectious for humans and is designated as a level 2 infectious agent. It should be handled in BSL2 facilities. All liquid and solid wastes should be disposed of after inactivation.
Acknowledgments
This protocol was adapted from our previous works: Murayama et al. (2021), Honda et al. (2021), and Washizaki et al. (2022). This work was supported by the Program on the Innovative Development and the Application of New Drugs for Hepatitis B (JP22fk0310517 and JP22fk0310503).
Competing interests
The authors declare that they have nothing to disclose.
References
Honda, T., Yamada, N., Murayama, A., Shiina, M., Aly, H. H., Kato, A., Ito, T., Ishizu, Y., Kuzuya, T., Ishigami, M., et al. (2021). Amino acid polymorphism in hepatitis B virus associated with functional cure. Cell. Mol. Gastroenterol. Hepatol. 12(5): 1583–1598.
König, A., Yang, J., Jo, E., Park, K. H. P., Kim, H., Than, T. T., Song, X., Qi, X., Dai, X., Park, S., et al. (2019). Efficient long-term amplification of hepatitis B virus isolates after infection of slow proliferating HepG2-NTCP cells. J. Hepatol. 71(2): 289–300.
Murayama, A., Momose, H., Yamada, N., Hoshi, Y., Muramatsu, M., Wakita, T., Ishimaru, K., Hamaguchi, I. and Kato, T. (2019). Evaluation of in vitro screening and diagnostic kits for hepatitis B virus infection. J. Clin. Virol. 117: 37–42.
Murayama, A., Yamada, N., Osaki, Y., Shiina, M., Aly, H. H., Iwamoto, M., Tsukuda, S., Watashi, K., Matsuda, M., Suzuki, R., et al. (2021). N‐Terminal preS1 sequence regulates efficient infection of cell‐culture–generated Hepatitis B Virus. Hepatology 73(2): 520–532.
Otoguro, T., Tanaka, T., Kasai, H., Kobayashi, N., Yamashita, A., Fukuhara, T., Ryo, A., Fukai, M., Taketomi, A., Matsuura, Y., et al. (2020). Establishment of a cell culture model permissive for infection by hepatitis B and C viruses. Hepatology Commun. 5(4): 634–649.
Washizaki, A., Murayama, A., Murata, M., Kiyohara, T., Yato, K., Yamada, N., Aly, H. H., Tanaka, T., Moriishi, K., Nishitsuji, H., et al. (2022). Neutralization of hepatitis B virus with vaccine-escape mutations by hepatitis B vaccine with large-HBs antigen. Nat. Commun. 13(1): e1038/s41467-022-32910-z.
Article Information
Copyright
© 2023 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 biochemistry > Protein
Cell Biology > Cell-based analysis > Viral infection
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4,780 | https://bio-protocol.org/en/bpdetail?id=4780&type=0 | # Bio-Protocol Content
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Three-color dSTORM Imaging and Analysis of Recombination Foci in Mouse Spread Meiotic Nuclei
LK Lieke Koornneef
MP Maarten W. Paul
AH Adriaan B. Houtsmuller
WB Willy M. Baarends *
JS Johan A. Slotman *
(*contributed equally to this work)
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4780 Views: 718
Reviewed by: Xiaokang WuJames H. Crichton Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in PLOS Genetics Jul 2022
Abstract
During the first meiotic prophase in mouse, repair of SPO11-induced DNA double-strand breaks (DSBs), facilitating homologous chromosome synapsis, is essential to successfully complete the first meiotic cell division. Recombinases RAD51 and DMC1 play an important role in homology search, but their mechanistic contribution to this process is not fully understood. Super-resolution, single-molecule imaging of RAD51 and DMC1 provides detailed information on recombinase accumulation on DSBs during meiotic prophase. Here, we present a detailed protocol of recombination foci analysis of three-color direct stochastic optical reconstruction microscopy (dSTORM) imaging of SYCP3, RAD51, and DMC1, fluorescently labeled by antibody staining in mouse spermatocytes. This protocol consists of sample preparation, data acquisition, pre-processing, and data analysis. The sample preparation procedure includes an updated version of the nuclear spreading of mouse testicular cells, followed by immunocytochemistry and the preparation steps for dSTORM imaging. Data acquisition consists of three-color dSTORM imaging, which is extensively described. The pre-processing that converts fluorescent signals to localization data also includes channel alignment and image reconstruction, after which regions of interest (ROIs) are identified based on RAD51 and/or DMC1 localization patterns. The data analysis steps then require processing of the fluorescent signal localization within these ROIs into discrete nanofoci, which can be further analyzed. This multistep approach enables the systematic investigation of spatial distributions of proteins associated with individual DSB sites and can be easily adapted for analyses of other foci-forming proteins. All computational scripts and software are freely accessible, making them available to a broad audience.
Key features
• Preparation of spread nuclei, resulting in a flattened preparation with easy antibody-accessible chromatin-associated proteins on dSTORM-compatible coverslips.
• dSTORM analysis of immunofluorescent repair foci in meiotic prophase nuclei.
• Detailed descriptions of data acquisition, (pre-)processing, and nanofoci feature analysis applicable to all proteins that assemble in immunodetection as discrete foci.
Graphical overview
Keywords: Meiosis RAD51 DMC1 Super-resolution microscopy dSTORM Single-molecule analysis SYCP3 Spermatocytes Nuclear spreading
Background
Many studies describe analyses using spread meiotic nuclei and immunofluorescence to visualize proteins involved in chromosome pairing and DNA double-strand break (DSB) repair. This mostly involves (fluorescent) light microscopy, which has the benefit of multi-color imaging but limited spatial resolution, restricted to ~250 nm. Electron microscopy overcomes this diffraction limit but has its pitfalls in complicated time-consuming sample preparation and limited labeling possibilities. Still, electron microscopy has revealed details of the two lateral elements of the synaptonemal complex (SC), forming between two homologous chromosomes during meiotic prophase and spaced ~200 nm apart, that were not resolved in light microscopy (Barlow et al., 1993).
To elucidate the spatiotemporal localization of proteins in meiosis, combining high resolution with imaging of multiple proteins is critical, and this possibility became a reality upon the development of super-resolution microscopy techniques (Cremer and Cremer, 1978). One type is single-molecule localization microscopy (SMLM), where blinking fluorophores are precisely localized in a sample, currently yielding the highest resolution in light microscopy (~5–20 nm). Direct stochastic optical reconstruction microscopy (dSTORM) is an indirect SMLM technique using fluorophores conjugated to antibodies bound to target proteins (Betzig et al., 2006; Rust et al., 2006).
dSTORM microscopy is very suitable for applications on spread meiotic nuclei due to the thin-layered chromatin preparations, low background achieved through the loss of most freely diffusing proteins, and the availability of a toolbox of well-characterized antibodies. We are interested in the localization patterns of two proteins involved in meiotic DNA DSB repair, RAD51 and DMC1, in the context of chromosome pairing, using SYCP3 as a marker of the lateral elements of the SC. Here, we describe an improved version of the nuclear spreading protocol that originates from Peters et al. (1997) in terms of optimal spreading quality, reproducibility, and suitability for dSTORM microscopy. For example, the requirement for coverslips rather than microscopy slides asked for some adaptations to ensure adherence of nuclei to the coverslip (coating step) and to avoid handling the vulnerable coverslips as much as possible.
Previously, we performed two-color dSTORM of recombinases RAD51 and DMC1 (Slotman et al., 2020) and recently expanded this to three-color dSTORM in combination with semi-automated selection of recombination foci (Koornneef et al., 2022). Here, we present the detailed protocol for this optimized pipeline, including the improved spreading procedure, immunofluorescent sample preparation, dSTORM data acquisition, pre-processing, and data analysis. Three-color dSTORM analyses allow high-resolution features to be discriminated, such as the precise position of each lateral element relative to the recombinase(s), to assess whether a particular recombinase is in close proximity with these components of the SC or localizes in the central area or outside the SC. The updated nuclear spreading protocol is specifically of interest to the meiosis community but might also be tested for other cell types. In addition, irrespective of the sample fixation protocol, the data acquisition, pre-processing, and data analysis described in this protocol are generally applicable and of interest to a broad public working on super-resolution microscopy in different fields.
Materials and reagents
Note: Materials and reagents not provided with company and catalog number information can be ordered from any qualified company for this experiment.
Biological materials
Laboratory-bred mice
Note: Mice were socially housed in individually ventilated cages with food and water ad libitum, in 12:12 h light/dark cycles.
Reagents
Paraformaldehyde (PFA) (Sigma-Aldrich, catalog number: P6148), storage temperature: 4 °C
dH2O
Sodium hydroxide (NaOH) (Honeywell, catalog number: 06203)
Sodium tetraborate decahydrate (Na2B4O7·10H2O) (Honeywell, catalog number: 31457)
Triton X-100 (Sigma-Aldrich, catalog number: T9284), storage temperature: room temperature (RT)
DPBS (Gibco, catalog number: 14190094), storage temperature: RT
Calcium chloride dihydrate (CaCl2·2H2O) (Sigma-Aldrich, catalog number: C7902)
Magnesium chloride hexahydrate (MgCl2·6H2O) (Honeywell, catalog number: M9272)
Sodium DL-lactate (Sigma-Aldrich, catalog number: L4263), storage temperature: 4 °C
Sucrose (Sigma-Aldrich, catalog number: 84097), storage temperature: RT
Tris base (Tris) (Sigma-Aldrich, catalog number: T6066)
Sodium citrate tribasic dihydrate (Sodium citrate) (Sigma-Aldrich, catalog number: 71406)
Ethylenediaminetetraacetic acid disodium salt dihydrate (EDTA) (Sigma-Aldrich, catalog number: E1644)
DL-Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D0632), storage temperature: 4 °C
Hydrochloric acid (HCl) (Honeywell, catalog number: 30721)
CO2 gas
Photo Flo (Kodak, catalog number: 5010640), storage temperature: RT
Sodium phosphate dibasic dihydrate (Na2HPO4·2H2O) (Sigma-Aldrich, catalog number: 71645)
Potassium dihydrogen phosphate (KH2PO4) (Sigma-Aldrich, catalog number: 04243)
Potassium phosphate monobasic (KCl) (Honeywell, catalog number: 60220)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: 71380)
Bovine serum albumin fraction V (BSA) (Roche, catalog number: 10735086001), storage temperature: 4 °C
Skim milk powder (Sigma-Aldrich, catalog number: 70166), storage temperature: RT
Mouse monoclonal anti-DMC1 (Abcam, catalog number: ab11054), aliquot storage temperature: -80 °C, stock concentration: 1 mg/mL
Note: After thawing an aliquot, the remainder is refrozen and then stored at -20 °C. Aliquot volumes were calculated to be sufficient for approximately 3–4 immunostainings to limit the number of freeze-thaw cycles.
Rabbit polyclonal anti-RAD51 [gift from R. Kanaar, previously generated and described by Essers et al. (2002)], aliquot storage temperature: -80 °C
Note: This antibody was stored at 4 °C after thawing.
Guinea pig anti-SYCP3 [gift from R. Benavente, previously generated and described by (Alsheimer and Benavente, 1996)], aliquot storage temperature: -20 °C
Notes:
This antibody was stored at 4 °C after thawing.
As an alternative to visualize SYCP3, we recommend using a mouse monoclonal antibody (Abcam, catalog number: ab97672).
Goat serum (Sigma-Aldrich, catalog number: G9023), storage temperature: -20 °C
Goat anti-guinea pig IgG Alexa 647 (Abcam, catalog number: ab150187), aliquot storage temperature: -80 °C, stock concentration: 2 mg/mL
Goat anti-rabbit IgG CF568 (Sigma-Aldrich, catalog number: SAB4600310), aliquot storage temperature: -20 °C, stock concentration: ~2 mg/mL
Goat anti-mouse IgG Atto488 (Rockland, catalog number: 610-152-121S), aliquot storage temperature: -20 °C, stock concentration: 1 mg/mL
Note: All secondary antibodies were stored at 4 °C after thawing. Please note that the quality of all antibodies decreases over time.
TetraSpeckTM microspheres 100 nm (Thermo Fisher, catalog number: T7279)
D-(+)-Glucose anhydrous (Sigma-Aldrich, catalog number: 49139)
Catalase (Sigma-Aldrich, catalog number: C9322)
Glucose oxidase (Sigma-Aldrich, catalog number: G2133)
Cysteamine hydrochloride (MEA) (Sigma-Aldrich, catalog number: M6500)
Immersion oil: ImmersolTM 518 F (Carl Zeiss, Jena, catalog number: 4449640000000)
Solutions
1 M NaOH (see Recipes)
50 mM borate buffer (see Recipes)
1.1 M CaCl2·2H2O (see Recipes)
0.5 M MgCl2·6H2O (see Recipes)
100 mM sucrose (see Recipes)
Hypobuffer (see Recipes)
Coated coverslips (see Recipes)
0.08% photo Flo (see Recipes)
20× PBS (see Recipes)
1× PBS (see Recipes)
Blocking buffer 1 (see Recipes)
Primary antibody buffer (see Recipes)
Blocking buffer 2 (see Recipes)
5 M NaCl (see Recipes)
1 M Tris-HCl pH 8 (see Recipes)
5× glucose (see Recipes)
dSTORM enzymes (see Recipes)
25 mM MEA (see Recipes)
Recipes
1 M NaOH
40 g of NaOH
Add dH2O up to 1 L
50 mM borate buffer
3.81 g of Na2B4O7·10H2O
Add dH2O up to 200 mL
Adjust the pH to 9.2 with 1 M NaOH
1.1 M CaCl2·2H2O
1.62 g of CaCl2·2H2O
Add dH2O up to 10 mL
Prepare 0.5 mL aliquots and store them at RT
0.5 M MgCl2·6H2O
1.02 g of MgCl2·6H2O
Add dH2O up to 10 mL
Prepare 0.5 mL aliquots and store them at RT
100 mM sucrose
3.42 g of sucrose
Add dH2O up to 100 mL
Prepare 1 mL aliquots and store them at -20 °C
Hypobuffer
1 mL of 600 mM Tris HCl pH 8.2 (0.73 g of Tris base in 10 mL of dH2O)
2 mL of 500 mM sucrose pH 8.2 (3.42 g of sucrose in 20 mL of dH2O)
2 mL of 170 mM sodium citrate pH 8.2 (0.50 g of sodium citrate in 10 mL of dH2O)
200 μL of 500 mM EDTA pH 8.2 (1.86 g of EDTA in 10 mL of dH2O)
100 μL of 100 mM DTT (0.15 g of DTT in 10 mL of dH2O)
14.7 mL of dH2O
Add dH2O up to 5 mL
Prepare 1 mL aliquots and store them at -20 °C
Note: All components for this buffer should be individually calibrated to a pH of 8.2 (except for DTT).
Coated coverslips
Boil the coverslips in dH2O for 20 min in the microwave (900 W)
Let them air dry
Coat the dry coverslips with 0.01% poly-L-lysine
Mix 3 mL of 0.1% poly-L-lysine with 27 mL of dH2O in a 15 cm Petri dish and put the dry boiled coverslips in the solution for 5 min at RT
Let the coverslips dry in an incubator at 60 °C for 1 h by placing them against the side of a 6-well plate
0.08% photo Flo
160 μL of photo Flo
Add dH2O up to 200 mL
Note: Prepare this just prior to use.
20× PBS
178 g of Na2HPO4·2H2O
24 g of KH2PO4
20 g of KCl
800 g of NaCl
Add dH2O up to 4 L
Adjust the pH to 7.4 with NaOH
Add dH2O up to a final volume of 5 L
1× PBS
45 mL of 20× PBS
Add dH2O up to 900 mL
Blocking buffer 1
0.25 g of BSA
0.25 g of skim milk powder
Add 1× PBS up to 50 mL
Make 1 mL aliquots and store them at -20 °C
Note: Thaw before use.
Primary antibody buffer
1 g of BSA
Add 1× PBS up to 10 mL
Make 1 mL aliquots and store them at -20 °C
Note: Thaw before use.
Blocking buffer 2
0.5 g of skim milk powder
Add 1× PBS up to 10 mL
Centrifuge for 10 min at maximum speed in an Eppendorf centrifuge
Transfer the supernatant to a new tube and add 10% of normal goat serum
Note: Prepare this just prior to use.
5 M NaCl
29.2 g of sodium chloride
Add dH2O up to 100 mL
1 M Tris-HCl pH 8
12.11 g of Tris
Add dH2O up to 80 mL
Adjust the pH to 8 with HCl
Add dH2O up to a final volume of 100 mL
5× glucose
20 g of glucose
10 mL of 1 M Tris-HCl pH 8
400 μL of 5 M NaCl
15 mL of dH2O
Dissolve glucose and add dH2O up to a final volume of 40 mL
Notes:
Thaw before use.
Heating can speed up the dissolving process.
dSTORM enzymes
56 mg/mL glucose oxidase, 3.4 mg/mL catalase in 50 mM Tris-HCl pH 8 and 10 mM NaCl
Mix 5 μL of 5 M NaCl, 125 μL of 1 M Tris-HCl pH 8, and 2.37 mL of dH2O.
Let the catalase thaw completely. When thawed, weigh 3.4 mg of catalase and dissolve it in 1 mL of NaCl/Tris-HCl solution. For 10KU glucose oxidase: check the U/g (in our case, 224890 U/g) to calculate the quantity in grams. 10,000/224,890 = 44.5 g of glucose oxidase.
For 56 mg/mL, use 44.5/56 = 795 μL of the catalase/NaCl/Tris-HCl solution. Add this volume of solution to the storage tube of glucose oxidase and dissolve well.
Prepare aliquots of 30 μL and store at -20 °C. Thaw before use.
25 mM MEA
10 μL of 5 M NaCl
250 μL of 1 M Tris-HCl pH 8
0.57 g of MEA
Add dH2O up to 5 mL
Prepare 100 μL aliquots and store them at -20 °C
Note: Thaw before use.
Laboratory supplies
Note: Laboratory supplies not provided with company and catalog number can be ordered from any qualified company for this experiment.
Disposable gloves
Aluminum foil
Pipette tips
Crushed ice
0.2 μm syringe filter (Whatman, catalog number: WHA10462300)
50 mL syringe (B.Braun, catalog number: 4616502F)
1.5 mL reaction tube (Greiner Bio-One, catalog number: 616201)
50 mL tube (Greiner Bio-One, catalog number: 227285)
10 cm Petri dish (Sarstedt, catalog number: 83.3902)
15 mL tube (Sarstedt, catalog number: 62554502)
10 mL pipette (Greiner Bio-One, catalog number: 607180)
24 mm diameter No. 1.5H (170 ± 5 μm) high precision round coverslips (Marienfield, catalog number: 0117640)
0.1% poly-L-lysine (Sigma-Aldrich, catalog number: P8920), storage temperature: RT
15 cm Petri dish (Sarstedt, catalog number: 83.3903)
6-well plate (Greiner Bio-One, catalog number: 657165)
Microscopic slide (MLS, catalog number: JK41301)
35 mm Petri dish (Falcon, catalog number: 353001)
3MTM tape (Permanento, number: 202)
Parafilm (Sigma-Aldrich, catalog number: P7793)
Equipment
Note: Equipment not provided with company and catalog number can be ordered from any qualified company for this experiment.
250 mL beaker
100 mL graduated cylinder
Pipettes
Microwave
Magnetic stirring bar
Magnetic stirring plate with heating capacity (IKA, model: RET B)
Fume hood
pH meter
Timer
Nutator (Clay Adams, model: 421106)
Euthanasia chamber for mice
Scissors and tweezers to dissect the testis
Curved tweezers with fine points for disrupting the tubuli (e.g., VWR, catalog number: 232-0110)
Pipette controller
Benchtop centrifuge (Eppendorf, model: 5810R)
Counting chamber (VWR, catalog number: BRND718920)
Phase contrast & dark field microscope (Olympus, model: BX41)
Tweezer for handling the coverslip (e.g., VWR, catalog number: 232-0174)
Incubator (Lab-Line Instruments, model: 308-1)
Microscope slide box (with wet paper inside to make a humid chamber) (Kartell, catalog number: 278)
Ultra-low temperature freezer (Sanyo, model: MDF-794)
Wheaton Coplin staining jar (Merck, catalog number: S6016)
Tilt shaker (shaking plate) (Edmund Bühler, catalog number: WS-10)
Dark box to store 6-well plate
Microcentrifuge (Thermo Scientific, model: Pico 17)
AttofluorTM cell chamber (microscopic ring) (Thermo Fisher, catalog number: A7816)
Ultrasonic cleaner (Brandson, model: B200)
Microscope [Zeiss Elyra PS1 system with a 100× 1.46 NA oil immersion objective, Andor iXon DU897 EMCCD camera (512 × 512)]
Note: Any TIRF microscope with high-power lasers > 100 mW and a sensitive camera, sCMOS, or EMCCD can be used. However, the workflow described here using the ZEN software only runs on the Zeiss Elyra microscope.
Software and datasets
ZEN2012 SP5 FP3 (Carl Zeiss, version 14.0.18.201)
Fiji/ImageJ (version > 1.53t, National Institutes of Health, USA, https://imagej.nih.gov/ij/) (Schindelin et al., 2012)
R (version >3.0) (R CoreTeam, 2018)
RTools (version 4.0, https://cran.r-project.org/bin/windows/Rtools/rtools40.html)
RStudio (Posit, 2015, version 4.0.3)
SMoLR (Paul et al., 2019) (available at https://github.com/ErasmusOIC/SMoLR , on the GitHub page you can find details on how to install the software)
Fiji Plugin “STORM_Tools.jar” (available at https://github.com/ErasmusOIC/STORM_Tools/tree/Publication)
Custom data analysis pipelines in Fiji (available at https://github.com/ErasmusOIC/STORM_Tools/tree/Publication/Scripts/)
Fiji script “Align_dSTORM_channels.ijm”
Fiji script “Create_reference.ijm”
Fiji script “Create_multi-color_dSTORM_image.ijm”
Fiji script “ROI_selection.ijm”
Fiji script “Manual_ROI_selection.ijm”
Custom data analysis pipelines in R (available at https://github.com/ErasmusOIC/STORM_Tools/tree/Publication/R_Scripts/)
R script “R scripts for recombination foci analysis using dSTORM.R”
A test dataset for the data analysis pipeline is available via BioImage Archive (http://www.ebi.ac.uk/bioimage-archive) under accession number S-BIAD627. This data is part of the study described before (Koornneef et al., 2022).
Procedure
Spreading spermatocyte nuclei for dSTORM
Notes:
This spreading procedure is based on a previously described protocol (Peters et al., 1997).
This procedure can also be used for spread nuclei preparations on rectangular microscopy slides for other types of microscopy purposes. For details about spreading on these slides, see notes within the procedure.
Make 1% PFA solution as follows: place 1 g of PFA in a 250 mL beaker and cover it with aluminum foil. Add 90 mL dH2O and one drop of 1 M NaOH (Recipe 1). Remove aluminum foil and warm 2 × 10 s in the microwave (900 W) with careful shaking in between. Place the magnetic stirring bar in the solution and cover it with aluminum foil. Warm the solution to 50 °C for 10 min by placing the beaker on a magnetic stirring plate with a heating capacity of ~75 °C (in the fume hood). The solution becomes clear (see Note 1). Cool the solution by placing the glass beaker on ice for 15 min. Adjust the pH to 9.2–9.5 by adding 2 mL of 50 mM borate buffer (Recipe 2). Add more 1 M NaOH if necessary. Adjust the volume with dH2O to 100 mL. Filter the solution with a 0.2 μm syringe filter and a 50 mL syringe into a new beaker (see Note 2). Dissolve 150 μL of Triton X-100 in the solution using magnetic stirring (see Note 3).
Note 1: If the solution does not become clear, incubate longer. It could be that the temperature of the solution is too low, but make sure that the NaOH was added in advance.
Note 2: It can be useful to replace the filter once after filtering half of the PFA solution to avoid clogging of the filter.
Note 3: Cut a small part of the plastic tip to pipette Triton X-100 more easily; add this tip to the solution to ensure that the whole volume is transferred into the solution. Remove and discard the tip when the Triton X-100 has dissolved.
Prepare PBS+ as follows: mix 50 mL of DPBS, 50 μL of 1.1 M CaCl2·2H2O (Recipe 3), 50 μL of 0.5 M MgCl2·6H2O (Recipe 4), and 25 μL of sodium DL-lactate to a 50 mL tube. Place the PBS+ on a nutator until use.
Note: Cut the tip to facilitate pipetting the sodium DL-lactate solution.
Thaw the following solutions: 0.5 mL of 100 mM sucrose (Recipe 5) and 1 mL of Hypobuffer (Recipe 6).
Sacrifice a mouse (at least three weeks old) using CO2 gas followed by cervical dislocation.
Note:
Mice can also be younger or older; but take into account that the composition of the testes changes while the mice are going through puberty.
The use of frozen material is discouraged because of the poor quality of the meiotic nuclear spreads obtained. If only frozen material is available, progress as described below but take into account that the yield will be lower [this will be observed during cell counting (step A15)]. Therefore, only a few slides can be made from one frozen testis.
Dissect the testis and place it in a drop of PBS+ in a 10 cm Petri dish (Figure 1).
Figure 1. Testis collection. A. Place the sacrificed mouse on its back, wet the skin of the abdomen with 70% ethanol, and open the skin of the lower part of the abdomen with a small cut using scissors. Pinch the skin between the fingers of each hand just above and below the small cut and pull up and downward (along the anterior-posterior axes) to tear the skin further and visualize the peritoneum. This helps to avoid hair contamination. B. Use a tweezer to grasp the peritoneum and use a scissor to cut it open and to enlarge the opening laterally. C. The organs in the lower part of the abdominal cavity are visible at this point (intestine and bladder). The testis is descended and located in the scrotum. Most often, they are not yet visible. To retrieve them from the scrotum, use tweezers to grasp adipose tissue on either side of the bladder and softly pull each of them upwards. D. This will expose the testis, visible as an egg-shaped tissue that should be separated from the surrounding tissue such as the epididymis, blood vessels, etc., using tweezers and scissors to carefully isolate it.
Remove the tunica albuginea using tweezers and a fine scissor. This is a fibrous tissue covering the testis. Its removal generates an amorphous mass of testis tubules, connected by interstitial tissue.
Repeatedly place small parts of this tissue mass between two curved tweezers with fine points and rub these against each other in a drop of PBS+ to obtain a cell suspension (for ~5 min) (Video 1, Figure 2A and 2B).
Video 1. Generation of a cell suspension using curved tweezers with fine points
Figure 2. Spreading spermatocyte nuclei for direct stochastic optical reconstruction microscopy (dSTORM). A. Rubbing tissue fragments between two curved tweezers with fine points. B. Cell suspension after ~5 min of repeated rubbing of tissue fragments between the tweezers. C. Example of the result after the remaining tissue fragments have settled down. Only transfer the supernatant. D. Counting chamber. E and F. Example of testicular cells in the counting chamber. Only the large, round cells indicated in F are counted within the red rectangle or crossing the upper and/or right line (E). Scale bar represents 0.05 mm. G. Drops of sucrose in the top lid of a 10 cm dish. H. Adding the sucrose/cell suspension to the PFA drop. I. Humid chamber. J. Coverslip (the edge is accentuated with a white dotted line) with PFA and cells that is positioned on a microscope slide in the humid chamber. For the first hour of incubation, the lid of the humid chamber will be closed. K. Use pipettes to open the humid chamber for the last hour of fixation and drying. L. After the photo Flo wash step, dry the coverslip completely by placing it against the side of a 6-well plate.
Bring the whole cell suspension in a 15 mL tube using a pipette and wash the Petri dish with PBS+ several times to collect all cells. Add PBS+ up to a total volume of 10 mL.
Invert the tube twice and wait a few minutes for the larger tissue fragments to settle to the bottom (Figure 2C).
Transfer the supernatant to a clean 15 mL tube using a 10 mL pipette.
Centrifuge for 5 min at 617× g (1,000 rpm).
Remove the supernatant.
Add 1 mL of PBS+ and gently pipette up and down to carefully resuspend the pellet.
Load 10 μL of the cell suspension in the counting chamber (Figure 2D).
Using a phase contrast microscope, count the number of spermatocytes (large, round cells) that are visible at 40× magnification in vertical large rectangles (including cells that cross the upper and/or right side) and calculate the average of 10 rectangle counts. This average is multiplied by 106 to obtain the concentration of cells/mL (Figure 2E and 2F).
Note: When the number of spermatocytes per rectangle is low (e.g., many rectangles with a score of zero), count 20 rectangles to arrive at a more trustworthy average.
Add 1 mL of Hypobuffer (always an equal volume of Hypobuffer and cell suspension) to the cell suspension and wait for 8 min.
Add PBS+ to a total volume of 10 mL and centrifuge for 5 min at 617× g (1,000 rpm).
Remove the supernatant and resuspend the pellet in PBS+ in a volume that yields a concentration of 15 × 106 cells/mL. For example: the average cell count at step A15 was 2; this corresponds to a concentration of 2 × 106 cells/mL. To obtain a concentration of 15 × 106 cells/mL, divide 2 × 106 by 15 × 106 to obtain the resuspension volume (in mL) of PBS+. In this example, you add 133 μL of PBS+ to the pellet.
Make separate drops of 20 μL of 100 mM sucrose in a 10 cm Petri dish and add 10 μL of the cell suspension to each drop (pipette up and down to create a homogenous solution) (Figure 2G).
Hold the coated coverslip (Recipe 7) with a tweezer and scoop it through the PFA to end with a drop of PFA on the coverslip (Video 2).
Video 2. Scoopingthe coverslip into the PFA
Pipette the 30 μL cell suspension with sucrose in the PFA drop and disperse the solution by slowly moving the coverslip circularly (Figure 2H, Video 3).
Note: This is easiest if two people work together, one handling the coverslip and the other adding the cell suspension.
Video 3. Dispersing the solution by circular movements of the coverslip
Place the coverslip with the cells upwards on a microscope glass slide in a humid box (Figure 2I and 2J).
Leave the coverslips for 1 h in the humid box with the lid closed.
Generate a small opening by placing the lid on two pipettes and allow the coverslips to slowly dry for 1 h (Figure 2K).
Notes:
When making spreads on microscope glass slides, this second drying step only takes 30 min.
This is a critical step; it is necessary to always check your coverslip under the microscope to assess if cells have attached to the coverslip before continuing. If they are mostly still floating, incubate for 15 or 30 min longer until the majority of cells appear to have attached.
Always remember which side of the coverslip contains the cells, since this is not visible. Once you grab the coverslip with the tweezers, you have to keep track of which side should remain up. Grab the coverslip with a tweezer and wash in 200 mL of 0.08% photo Flo (Recipe 8) by slowly moving it through the solution for 10 s.
Let the coverslips air dry by placing them against the side of a 6-well plate (Figure 2L).
When dry, the coverslips are ready for immunocytochemistry (pause point). For long time storage, place the coverslips in a small Petri dish (with cells upwards) closed with small pieces of tape and store at -80 °C, or continue directly with the immunocytochemistry procedure.
Note: When making spreads on rectangular microscope glass slides, dried slides can be packaged back-to-back in aluminum foil and also stored at -80 °C.
Immunocytochemistry on nuclear spread preparations
Thaw the coverslip for 10 min at RT.
Place the coverslip into a 6-well plate with dH2O between the wells in a box with wet paper (humid chamber for immunocytochemistry). Keep the 6-well plate as much as possible in the box with wet paper during all the next steps.
Wash the coverslip in 2 mL of 1× PBS (Recipe 10) on a shaking plate for 10 min and repeat this two times.
Note: When using spreads on microscope glass slides, use a Coplin jar for washing.
Block the chromatin on the coverslip by adding 700 μL of blocking buffer 1 drop by drop on the coverslip (Recipe 11) and incubate for 20 min.
To perform the three-color staining of SYCP3, RAD51, and DMC1, add 0.5 μL of SYCP3 antibody, 1 μL of DMC1 antibody (from a 1:10 dilution, which is 1 μL of antibody and 9 μL of 1× PBS), and 1 μL of RAD51 antibody (from a 1:10 dilution, which is 1 μL of antibody and 9 μL of 1× PBS) to 100 μL of primary antibody buffer (Recipe 12).
Remove blocking buffer 1 using a pipette. Subsequently, add the primary antibody buffer including antibodies as prepared in step B5.
Cover the coverslip with a small circle of Parafilm that has approximately the same size as the coverslip and place the Parafilm circle on the coverslip with buffer using tweezers for equal distribution.
Incubate the coverslip with the primary antibody overnight at RT.
Note: It is possible to perform incubation also at 4 °C for two nights but, before storing at 4 °C, the coverslip should be incubated for at least 1 h at RT. Before continuing with the protocol on the next day, it should also be incubated at RT for at least 1 h.
Gently remove the Parafilm using tweezers.
Wash the coverslip in 2 mL of 1× PBS on a shaking plate for 10 min and repeat this two times.
Block the coverslip by adding 700 μL of blocking buffer 2 (Recipe 13) and incubate for 20 min.
To perform the three-color staining of SYCP3 (Alexa 647), RAD51 (CF568), and DMC1 (Atto488), add 2 μL of goat anti-guinea pig Alexa 647 antibody (from a 1:10 dilution, which is 1 μL of antibody and 9 μL of 1× PBS), 2 μL of goat anti-rabbit CF568 antibody (from a 1:10 dilution, which is 1 μL of antibody and 9 μL of 1× PBS), and 0.4 μL of goat anti-mouse Atto488 to 100 μL of blocking buffer 2 (Recipe 13). Protect this mix from light before use.
Gently remove the blocking buffer 2 and add the 100 μL of blocking buffer solution including antibodies that was prepared in step B12.
Cover the coverslip with Parafilm (see step B7).
Incubate the coverslip with the secondary antibody solution for 2 h at RT.
Gently remove the Parafilm.
Wash the coverslip in 2 mL of 1× PBS for 10 min on a shaking plate, repeat this twice, and protect it from light by wrapping it in aluminum foil.
Store the coverslip in 2 mL of 1× PBS in a 6-well plate (add water between the wells) in a dark box with wet paper (or a white box covered in aluminum foil) and store at 4 °C until further use.
Notes:
It is recommended to use a fresh immunostained sample for dSTORM. The sample, when stored in a dark box (or a 6-well plate kept in the dark), can last for months, but sample quality can decrease over time.
Two milliliters of PBS is enough for short-time usage but could evaporate when stored longer; please keep in mind to increase this volume when storing the coverslip for longer.
Addition of fiducials to the sample
Sonicate the vial containing the fiducials (TetraSpecksTM) for 20 s.
Pipette 0.5 mL of 1× PBS in an Eppendorf tube.
Place the coverslip in the AttofluorTM cell chamber and close the chamber carefully by screwing.
Note: Be careful with closing the chamber because closing it too loose will cause leakage and closing it too tight will cause the coverslip to break.
Pipette the fiducials at least 10 times up and down.
Add 0.35 μL of the fiducials to the PBS in the Eppendorf tube and mix thoroughly.
Add the fiducials with PBS to the sample in the AttofluorTM cell chamber and make sure that the liquid is dispersed equally. If not, softly tap to the side of the chamber.
Note: The distribution pattern of fiducials among different fields of view is often very variable, despite the thorough mixing of the solution beforehand. Selection of cells with a suitable number of fiducials before starting the actual dSTORM imaging is therefore essential (see step E2).
Add 0.5 mL of 1× PBS to the chamber.
Place the chamber in a 10 cm Petri dish and protect it from light.
Store it at 4 °C overnight to allow fiducials to settle.
Remove the solution with fiducials the next day and replace it with 1 mL of 1× PBS.
Note: The fiducials need to settle down; if this procedure is shortened, not enough fiducials will be visible on the sample, which makes the alignment of the different channels after imaging impossible.
dSTORM microscope setup
Turn on the microscope 1 h before imaging.
Start ZEN Software and turn on all lasers.
Prepare dSTORM buffer: mix 200 μL of 5× glucose (Recipe 16), 10 μL of dSTORM enzymes (Recipe 17), 25 μL of MEA (Recipe 18), and 765 μL of dH2O.
Note: Enzyme and MEA can expire over time, which decreases the blinking capacity during imaging.
Remove PBS from the sample in the AttofluorTM cell chamber and add 1 mL of dSTORM buffer.
Close the chamber with a coverslip.
Notes:
To make sure that the chamber is closed off from air, gently tap against the AttofluorTM cell chamber so that air bubbles that are present rise and become visible just below the top coverslip. Having a small air bubble in the chamber is not harmful and shows that the chamber is properly sealed due to capillary action.
The dSTORM buffer works for a maximum of 4–5 h after preparation. Thereafter, the blinking capacity will decrease and a new dSTORM buffer needs to be prepared and added to the sample.
Add a drop of oil to the 100× 1.46 NA objective.
Place the chamber in the microscope 30 min before imaging to let the dSTORM buffer establish its equilibrium at a stabilized temperature.
dSTORM imaging
Focus on the sample by eye through the oculars using the mercury lamp and appropriate filters.
Search for a nucleus. A good nucleus or cell is characterized by a strong signal of all fluorophores, little to no background signal of fluorophores, and an optimal number of fiducials (at least 5–10, maximum 50). Too few fiducials make the alignment difficult/impossible and too many fiducials interfere with your signal to be imaged.
Check the infrared signal using the camera (continuous mode) since it is not visible through the ocular lens.
Set up the microscope for three channels; in our case, by selecting the laser 488, 561, or 642 (“on” in Figure 3) and the corresponding filter block BP 495-575, BP 570-650, or LP655 (the latter is visibly selected in Figure 3).
Figure 3. Screenshot of ZEN software. Screenshot of ZEN software during direct stochastic optical reconstruction microscopy (dSTORM) imaging and pre-processing to reconstruct dSTORM image. The yellow numbers indicate the relevant location on the screen corresponding to the steps described in Section E. The yellow rectangle indicates the relevant location on the screen related to Data analysis “Pre-processing to reconstruct dSTORM image.” i, ii, and iii indicate screen parts that become (completely) visible when sliding down the bar that is on the right side of the acquisition parameters column.
Only for the first nucleus: set the TIRF angle by focusing on the sample using the AF647 channel and select EPI as illumination mode. Move the TIRF angle slider to the right until the signal disappears; this setting is just beyond the optimal TIRF angle. Then, move the slider to the left and stop when the penetration depth indicated in the software is between 190 and 200 nm. At this TIRF angle, only a small region just above the coverslip is illuminated, giving high signals and no background from out-of-focus contributions. Repeat this for CF568 and ATTO488.
Note: A penetration depth between 190 and 200 nm should illuminate the full sample but not more. This can be achieved by moving from no signal until the brightest signal of the stained protein is visible.
Take a snapshot (z-stack) of the nucleus: select all channels one by one (Figure 3, screen i) and set the following settings: laser power = 2% (indicated with the yellow triangle), exposure time = 100 ms, EMCCD gain = 100 a.u., light path (Figure 3, main screenshot, under Imaging setup bar): switch track every = z-stack, Under acquisition mode bar: averaging = 2. Select the option Z-stack (Figure 3, main screenshot, top left, and Figure 3, screen ii), ensure that the image is in focus, and use the z-stack Center option. Thereafter, press center to put the z-stack in the middle of the cell. Select settings to make a z-stack with 35 slices of 0.110 μm.
Notes:
When the signal is saturated, lower the gain.
These settings depend on the specific microscope and camera used. This is a low laser power setting to avoid monitor bleaching.
Press Start Experiment .
Save the snapshot as a .czi file using file save (Exp1_snap.czi). The use of an experiment name is essential for the data analysis. In this example, we choose to use Exp1 as our experiment name.
Continue with the dSTORM imaging by selecting the following: Online Processing PALM (peak mask size = 9 pixels, peak intensity noise = 6) (Figure 3, main screenshot, under Online processing Options ), and under acquisition mode bar: averaging = 1, deselect “Z-Stack” (Figure 3, main screenshot, top left) and instead select Time series (12,000 frames).
Focus on the observed image with the AF647 track selected.
Adjust settings as follows (Figure 3, screen i): laser power = 100%, exposure time = 33 ms, EMCCD gain = 300 a.u., and press Continuous (Figure 3, main screenshot, top left). This will directly increase the signal intensity followed by an overall decrease of the signal, and finally in clearly blinking fluorophores on the screen. Press Stop when all fluorophores blink well, which can depend on the dSTORM buffer quality or the type of fluorophore. Under normal conditions, this takes a few minutes for Alexa 647.
Note: These settings may have to be adapted depending on the microscope and camera. This setting pushes all fluorophores into a dark state, but high laser power can also bleach fluorophores. If too many fluorophores are bleached, lowering the laser power is essential. The camera is most sensitive when the gain is set at maximum (for EMCCD cameras) and when an exposure time is chosen that allows recording of individual separated blinking events. A longer exposure time will record more photons and thus lead to increased localization precision, but also yields a higher chance of recording multiple overlapping blinking events, which should be prevented.
Set the experiment to time series (Figure 3, screen iii) and define a series of 12,000 frames with 0 ms interval.
Press Start Experiment .
During the ongoing recording of 12,000 frames, make sure that the number of detections per frame (First Frame) stays approximately constant. When detection frequency decreases, this may be due to loss of focus or bleaching of fluorophores. Loss of focus can be assessed in the wide field image, and focus should be adjusted when necessary. If focus is corrected and detection frequency still decreases, bleaching of fluorophores can be slowed down by lowering the laser power to 90%, or an even lower percentage, but not lower than 70%.
Notes:
For Alexa 647, most often it is not necessary to adjust the laser power.
Systems with hardware autofocus are highly recommended.
Save the movie as a .czi file (Exp1_Ch1.czi).
Repeat steps E8–E15 for the CF568 (Exp1_Ch2.czi).
Note: Proper dSTORM imaging of the CF568 fluorophore normally requires a bit longer waiting time before it starts to blink properly, compared with Alexa 647 (5–8 min). Make sure not to start recording too early, to ensure the optimal yield of blinking events.
Repeat steps E8–E15 for ATTO488 (Exp1_Ch3.czi).
Notes:
Atto488 bleaches very fast. Therefore, when the high laser power is switched on at step E11, wait a few seconds and then directly start the experiment (steps E12–E13). During imaging, you will rapidly see a drop in the number of localization events per frame, and therefore the laser power should be lowered in steps of 10% to a minimum of 70% already during the first 1,000–1,500 frames. Green fluorophores are less suitable than longer wavelength fluorophores for dSTORM imaging, and therefore green dyes will give lower resolution images. This should be taken into account when choosing target proteins and their corresponding fluorescent label.
Another option to stimulate the blinking of the green dye is to use the 405 nm laser at very low power to release molecules from their dark state and increase the number of localization events.
Data analysis
Pre-processing steps to reconstruct the dSTORM image
Notes:
Install the Plugin STORM_Tools.jar in Fiji and RTools and SMoLR in R. Also, open the custom data analysis pipelines in Fiji and R.
Test data for the data analysis pipeline is available via BioImage Archive (http://www.ebi.ac.uk/bioimage-archive) under accession number S-BIAD627.
The use of an experiment name (e.g., the name of the experiment like Exp1 in the test data) is essential for the scripts to work properly.
Additional information about the SMoLR package can be found at https://github.com/ErasmusOIC/SMoLR and https://htmlpreview.github.io/?https://github.com/ErasmusOIC/SMoLR_data/blob/master/SMoLR.html . Additional help on each SMoLR function can be accessed by entering a “?” followed by the name of the specific function (i.e., ?SMOLR_PLOT).
Re-open the movies in the ZEN Software.
Perform a drift correction as follows. PAL drift: select model-based approach and press apply two times (Figure 3 yellow square). Save the table as a .txt file by right-clicking on the table. Repeat this for every channel (Exp1_Ch1.txt, Exp1_Ch2.txt, Exp1_Ch3.txt).
Note: The drift correction is needed to correct for small drifts that occurred during imaging.
Perform grouping as follows. PAL Group: max on time = 50, off gap = 2, capture radius = 4 pixels (Figure 3 yellow square). Save the table as a .txt file by right-clicking on the table. Repeat this for every channel (Exp1_ Ch1_g.txt, Exp1_Ch2_g.txt, Exp1_Ch3_g.txt).
Note: Grouping is essential to couple blinking events from the same fluorophore within a specific time range.
The pre-processing continues in Fiji, where you start with seven files: a z-stack image (Exp1_snap.czi) and, per channel, a drift-corrected localization file (Exp1_Ch1.txt, Exp1_Ch2.txt, Exp1_Ch3.txt) and a drift-corrected and grouped localization file (Exp1_Ch1_g.txt, Exp1_Ch2_g.txt, Exp1_Ch3_g.txt). Align the different channels in Fiji using the script Align_dSTORM_channels by using a gap of 50 and a track length of 70 (Figure 4A). The ungrouped localization files (Exp1_Ch1.txt, Exp1_Ch2.txt, Exp1_Ch3.txt) are used to identify fiducials, and the grouped location files (Exp1_Ch1_g.txt, Exp1_Ch2_g.txt, Exp1_Ch3_g.txt) are used for further data analysis. This generates two transformation files (Exp1_Ch2_transformation.txt and Exp1_Ch3_transformation.txt) containing the transformation matrix to align Ch2 and Ch3 with Ch1 as a template and two localization files containing the transformed localizations from Ch2 and Ch3 (Exp1_Ch2_gt.txt and Exp1_Ch3_gt.txt).
Notes:
Name the channels based on wavelength, whereby channel 1 corresponds to the highest wavelength. In case you only have two channels, exclude channel 3.
When the number of fiducials found is lower than 3 (or the alignment turned out not to be correct), increase the gap size or decrease the track length.
The script Align_dSTORM_channels is based on the DoM Plugin (Katrukha et al., 2022).
The directory asked for in the script is the location where the .txt files are stored.
Figure 4. Flowchart of pre-processing to reconstruct direct stochastic optical reconstruction microscopy (dSTORM) image. A. Start with the alignment of dSTORM channels by aligning the localization tables in Fiji using the script Align_dSTORM_channels . An example of a fiducial before and after alignment is shown (steps A1–A4). B. Next, determine the outline of the nucleus in the z-stack image to generate a reference in Fiji using the script Create_reference (step A5). C. The aligned localization tables and the reference are used in R using the R script Create dSTORM image per channel to generate the dSTORM images per channel (step A6). D. Finally, the separate dSTORM images per channel are combined in Fiji to create a multi-color dSTORM image using the script Create_multi-color_dSTORM_image (step A7). An example of a three-color dSTORM image of a spread spermatocyte nucleus immunostained for SYCP3 (magenta), RAD51 (red), and DMC1 (green) is shown, including an enlargement of the region indicated with the dotted square. Scale bar represents 100 nm (A), 250 nm (D enlargement), and 5 μm (B, D).
Create a reference for the dSTORM image by using the z-stack image in Fiji and running the script Create_reference (Figure 4B). In short, select the z-position with the best focus for each channel, merge these single images, and save this image (Exp1_Composite.tif). Determine the border of the nucleus saved as a region of interest (ROI) (Exp1_Roi.roi) including its characteristics (Exp1_Roi_characteristics.csv). This reference is created to reduce computational time in the following steps of the pre-processing.
Notes:
To open the .czi file via Bio-Formats (Plugins/Bio-Formats/Bio-Formats Importer), set the following settings in Import Options (View stack with: Hyperstack, Color mode: Colorized, and select “Autoscale”).
The z-position can be found in the left upper corner of the image (z:1/35).
Use R and the package SMoLR to process the transformed localization files into a dSTORM image per channel by running the script Create dSTORM image per channel of the R script (Figure 2C). In short, the localization files are imported and converted to a data frame. The localizations within the previously selected nucleus ROI are transformed into an image (Exp1_Ch1.tif, Exp1_Ch2.tif, Exp1_Ch3.tif).
Notes:
Manually set the directory in the R script.
Generation of the images in R is a time-consuming step.
Combine the individual images of each channel in Fiji using the script Create_multi-color_dSTORM_image (Figure 4D). In short, this script uses Bio-Formats to import individual images. For each image, the image intensity is converted into a 16-bit image. Also, each image is colored (Ch1 = magenta, Ch2 = red, Ch3 = green) and all individual images are combined into one image, which is saved (Exp1_Ch1_Ch2_Ch3.tif).
Note: The script adjusts the brightness/contrast of the image using the auto settings. In case this is not correct, decrease the maximum value and/or increase the minimum value using the B&C control (Image > Adjust > Brightness/Contrast).
Check if the channels are correctly aligned, as shown in the example in Figure 4A.
Recombination foci analysis
Identify regions of interest (ROIs) that display RAD51 and/or DMC1 signals in Fiji using the script ROI_selection (Figure 5A). The details of this script were previously described in the Material and Methods section under the heading Recombination foci analysis of Koornneef et al. (2022). In short, this script uses the three-color dSTORM image (Exp1_Ch1_Ch2_Ch3.tif) and performs a semi-automatic identification of ROIs containing both RAD51 and/or DMC1 nanofoci using specific thresholds for focus size and SYCP3 signal. The default thresholds for focus size are 500 pixels for RAD51 and 300 pixels for DMC1, and the radius of each ROI is set at 375 nm. This whole procedure generates a set of all identified ROIs (Exp1_allRois.zip), a set of ROIs after manual adaption (Exp1_RoiSet.zip), a transformed set of ROIs after manual adaptation (Exp1_RoiSet_t.zip), and a file containing specific characteristics of each ROI (Exp1_RoiSet_t_characteristics.csv).
Note: To manually add ROIs (after visual inspection of all identified ROIs in the image), use the script Manual ROI selection with a radius of 375 nm. To select a ROI, click on the center of the region. Press T to add this ROI to the ROI Manager. To remove an ROI from the ROI Manager, select the ROI and press Delete. To stop the macro Manual ROI selection, finish the ROI_selection macro or press shift and click at a random position in the image.
Figure 5. Flowchart of recombination foci analysis. A. Select regions of interest (ROIs) containing RAD51 and DMC1 foci in the three-color direct stochastic optical reconstruction microscopy (dSTORM) image in Fiji using the script ROI_selection (step B1). An example of part of a three-color dSTORM image of a spread spermatocyte nucleus immunostained for SYCP3 (magenta), RAD51 (red), and DMC1 (green). ROIs of 750 nm diameter are indicated as yellow circles. B. Combine aligned localization files of different channels into one data frame in R using the script Combine channels (step B2). C. The combined localization file and the ROIs are combined to generate small dSTORM localization files of every ROI, whereafter the clusters of localizations (termed nanofoci) are determined. The number of nanofoci within an ROI determines the DxRy configuration correlating with x DMC1 nanofoci and y RAD51 nanofoci. The example is a kernel density estimation of a ROI, whereby the dashed green and red lines visualize the binary outline of the nanofoci. This ROI has a D2R1 configuration, as it contains two DMC1 nanofoci and a single RAD51 nanofocus. All these steps are performed in R using the script Analysis of ROIs (step B3). D, F. The DxR y configuration can be used to determine the frequency (D, step B3) or create binary images of every ROI (E, step B4). Also, features of RAD51 and DMC1 nanofoci can be measured (F, step B5). Scale bar represents 250 (A) and 100 nm (C, E).
Combine the localization files from each channel into one data frame in R (Exp1_alldata.txt), using the script Combine Channels (Figure 5B).
Process the ROIs in R into DxRy configurations using the script Analysis of ROIs (Figure 5C). In short, a list is generated in which for each ROI all the localization events corresponding to all channels are documented (Exp1_data_sub.RData). Next, a kernel density estimation is performed on the subset of localization data from each ROI to cluster localization events (Exp1_kde.RData). A signal density threshold [0.05 (axes component) or 0.15 (recombinases) localizations/nm2] is used to create binary clusters termed nanofoci. The number of nanofoci within a ROI determines the DxRy configuration and thereby also the DxRy frequency that can be then determined for each nucleus (Figure 5D). Nanofoci smaller than 1,250 nm2 are excluded. Also, a statistics file (Exp1_sf.RData) is generated with details about the ROIs and their DxRy configuration information.
Notes:
Manually set the directory, the threshold for the kernel density estimate, and the threshold for the area in the R script.
Generation of data_sub.RData and kde.RData is a time-consuming step.
Optional: Generate binary images of individual ROIs in R using the script Generate binary DxRy images .
Note: The colors (red, green, and blue) in the generated images represent channels 1, 2, and 3, respectively.
Features of nanofoci (like the position and the number of localizations) can be found in the cluster parameters of kde.RData (e.g., for ROI number 1, use kde[[1]]$clust_parameters). Features of nanofoci (like area and shape) can be found in features (e.g., for channel 3 in ROI number 1, use features[[1]]$channel_3 and “x.0.s.area” or “x.0.m.eccentricity”). Also, other features (like distances to other nanofoci) can be calculated using the data_sub.RData and kde.RData, but require additional processing.
Note: Manually set the directory and threshold for the area in the R script.
Acknowledgments
This protocol was used to obtain the data published in PLOS Genetics (Koornneef et al., 2022).
We would like to thank E. Sleddens-Linkels for help with image acquisition and videos to document the meiotic nuclei spreading procedure. We would like to thank I. de Bruin for testing the protocol including the analysis pipeline as a new user and R. Benavente for providing the SYCP3 antibody.
Competing interests
All authors declare they have no conflicts of interest or competing interests.
Ethical considerations
All procedures were in accordance with the European guidelines for the care and use of laboratory animals (Council Directive 86/6009/EEC). All animal experiments were approved by the local animal experiments committee DEC (Dutch abbreviation: Dier Experimenten Commissie) Consult and animals were maintained under the supervision of the Animal Welfare Officer.
References
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Barlow, A. L., Jenkins, G. and ap Gwynn, I. (1993). Scanning electron microscopy of synaptonemal complexes. Chromosome Res. 1(1): 9–13.
Betzig, E., Patterson, G. H., Sougrat, R., Lindwasser, O. W., Olenych, S., Bonifacino, J. S., Davidson, M. W., Lippincott-Schwartz, J. and Hess, H. F. (2006). Imaging Intracellular Fluorescent Proteins at Nanometer Resolution. Science 313(5793): 1642–1645.
Cremer, C. and Cremer, T. (1978). Considerations on a laser-scanning-microscope with high resolution and depth of field. Microsc. Acta. 81(1): 31–44.
Essers, J., Hendriks, R. W., Wesoly, J., Beerens, C. E., Smit, B., Hoeijmakers, J. H., Wyman, C., Dronkert, M. L. and Kanaar, R. (2002). Analysis of mouse Rad54 expression and its implications for homologous recombination. DNA Repair 1(10): 779–793.
Katrukha, E., Teeuw, J., bmccloin and den Braber, J. (2022). ekatrukha/DoM_Utrecht: Detection of Molecules 1.2.5 (1.2.5). Zenodo.
Koornneef, L., Slotman, J. A., Sleddens-Linkels, E., van Cappellen, W. A., Barchi, M., Tóth, A., Gribnau, J., Houtsmuller, A. B. and Baarends, W. M. (2022). Multi-color dSTORM microscopy in Hormad1-/- spermatocytes reveals alterations in meiotic recombination intermediates and synaptonemal complex structure. PLos Genet. 18(7): e1010046.
Paul, M. W., de Gruiter, H. M., Lin, Z., Baarends, W. M., van Cappellen, W. A., Houtsmuller, A. B. and Slotman, J. A. (2019). SMoLR: visualization and analysis of single-molecule localization microscopy data in R. BMC Bioinf. 20(1): e1186/s12859-018-2578-3.
Peters, A. H., Plug, A. W., van Vugt, M. J. and de Boer, P. (1997). A drying-down technique for the spreading of mammalian meiocytes from the male and female germline. Chromosome Res 5(1): 66–68.
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4,781 | https://bio-protocol.org/en/bpdetail?id=4781&type=0 | # Bio-Protocol Content
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Binging from Food to Alcohol: A Sequential Interaction Between Binging Behaviors in Male Wistar Rats
SC Sergio Cuesta-Martínez *
LR Leandro Ruiz-Leyva *
AJ Ana María Jiménez-García
TA Teresa Aparicio-Mescua
OL Olga López-Guarnido
RP Ricardo Marcos Pautassi
IM Ignacio Morón
CC Cruz Miguel Cendán
(*contributed equally to this work)
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4781 Views: 307
Reviewed by: Alejandro GrauMohammed Mostafizur Rahman Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Addiction Biology Mar 2022
Abstract
The development of excessive alcohol (ethanol) and/or highly palatable food self-administration is an essential task to elucidate the neurobiological mechanisms that underlie these behaviors. Previous work has highlighted that ethanol self-administration is modulated by both the induction of aversive states (i.e., stress or frustration) and by the concurrent availability of appetitive stimuli (e.g., food). In our protocol, rats are food deprived for three days until they reach 82%–85% of their ad libitum weight. After that, rats are exposed daily for 10 days to a brief binge or control eating experience with highly sugary and palatable food (i.e., the ingestion of 11.66 and 0.97 kcal/3 min, respectively), which is followed by a two-bottle-choice test (ethanol vs. water) in their home cages for 90 min. This model induces robust binge eating, which is followed by a selective increase in ethanol self-administration. Therefore, this protocol allows to study: a) behavioral and neurobiological factors related to binge eating, b) different stages of alcohol use, and c) interactions between the latter and other addictive-like behaviors, like binge eating.
Keywords: Alcohol Binge drinking Binge eating Addiction Two-bottle-choice test Wistar rats
Background
Animal models of binge drinking (BD) are crucial for clarifying the neurobiological and psychobiological mechanisms underlying this harmful behavior [see Jeanblanc et al. (2019) for a review]. Considerable advances have been made by forcibly administering ethanol (Becker and Lopez, 2016), employing operant paradigms (Simms et al., 2010), developing strains with a genetic predisposition to ethanol (Colombo et al., 2014), or using self-administration models in Wistar rats where ethanol is presented in an intermittent and temporally restricted fashion (Salguero et al, 2020; Ruiz-Leyva et al, 2020). These animal models, however, do not always mimic the contextual variables surrounding human drinking behaviors or fail to achieve pharmacologically relevant blood ethanol concentrations within the proposed time frame for this type of consumption. Moreover, alcohol use is driven by several factors. For instance, it has been observed that the induction of aversive states in conjunction with ethanol availability generally leads to an increase in ethanol consumption and favors the development of psychiatric diseases—including alcohol use disorders (AUD)—via epigenetic changes [for a review, see Pucci et al. (2019)]. In addition, the induction of frustration and anxiety (Díaz-Morán et al., 2013a and 2013b; Sabariego et al, 2013; Kawasaki et al., 2017; Jiménez-García et al., 2019) is associated with increases in ethanol consumption as a coping strategy (Ramirez-Castillo et al., 2019). Paradoxically, exposure to appetitive stimuli (i.e., food) also enhances ethanol consumption. Seminal studies showed that concurrent access to food and ethanol enhanced drinking behaviors in animals (Mello and Mendelson, 1971; Meisch and Thompson, 1972). In this respect, both substance use disorders (including AUD) and BD appear to be comorbid (or at least closely related) with binge eating (BE) (Ferriter and Ray; 2011; Munn-Chernoff et al., 2020). Conceptually, BE involves ingestion of large amounts of palatable (sugary and/or fatty) food in a short period of time, associated with a feeling of loss of control [see Dingemans et al. (2017) for a review].
The present protocol aims to serve as a guide for researchers interested in the study of the interaction of binge-like behaviors and should help replicability efforts. It is also an improvement over the models described above. An exhaustive description of the novel animal model of BE-BD interaction originally developed by Ruiz-Leyva et al. (2022) is presented. Here, the self-administration model induced remarkably high levels of absolute ethanol consumption (approximately 5 g/kg/90 min) and ethanol preference (80%–90%), much greater than those reported by other preclinical models. For instance, rats exposed to intermittent access to 20% ethanol in a two-bottle-choice procedure achieved 9–10 g/kg, yet they did that in a 24 h period, whereas those tested in the drinking-in-the-dark-multiple-scheduled access rarely exceed 5–6.5 g/kg/day. In this model, food-deprived rats are briefly exposed to a measurable amount of highly palatable sugary pellets immediately followed by a two-bottle-choice test (ethanol vs. water). Here, the authors reported dramatic and specific increases in ethanol intake as the BE behavior develops, leading to pharmacologically relevant blood ethanol concentrations. In addition, ethanol consumption turned compulsive by the last sessions and sensitive to naltrexone administration.
Materials and reagents
Anti-drip bottles with capacity of 150 mL (Classic Drinker de Luxe; Zooplus, Munich, Germany)
Filtered and autoclaved tap water
Rodent Dustless precision pellets (DPPs) (45 mg each, nutritional profile: 59.1% carbohydrate, 18.7% protein, 5.6% fat, 3.6 kcal/g) (Bio-serv, catalog number: F0021). Shelf life: 12 months. Store in a dry and ventilated place. The container should be tightly closed and protected from moisture
Ethyl alcohol (ethanol 96%, v/v; CH3CH2OH; 46.07 g/mol) (PanReac AppliChem, catalog number: 141085). This substance is highly flammable (liquid or vapor) and causes severe eye irritation. Store in a cool, ventilated place away from heat sources such as hot surfaces
Animals: adult male Wistar HAN rats (Envigo Laboratories, Barcelona, Spain), aged 70–80 days and weighing 240 g (± 37) at the beginning of the procedure
Equipment
Rectangular polycarbonate cages (42.5 cm × 26.5 cm × 15 cm) (Figure 1A)
Timer and chronometer (Digital Onstart 100) (Figure 1B)
Small plastic cups for depositing the DPPs
Weighing scale (WLC 1/A2 Precision Balance; Radwag©) (Figure 1C)
Plastic syringes (15 mL) and crystal flasks (600 mL) for diluting ethanol solutions (Figure 1D)
Personal protective equipment (laboratory coat, gloves, masks, etc.)
Figure 1. Equipment. A. Rectangular polycarbonate cage with grid. B. Timer and chronometer. C. Weighing. D. Syringe and flasks.
The alcohol concentrations of 6% or 10% were achieved by pouring 470 (or 450) mL of filtered and autoclaved tap water into 500 mL flasks and then adding 30 (or 50) mL of alcohol with the plastic syringe. Next, the solution was gently stirred to ensure proper dissolution of the alcohol.
Procedure
Stabling conditions, habituation, and general considerations
Stabling
Individualize each animal in polycarbonate standard cages with a bed of sawdust and free access to water and standard chow (i.e., ad libitum conditions). Use anti-drip bottles filled with water from the beginning in order to habituate the animals to them. Moreover, put the bottles in the middle of the cage’s grid to prevent side preferences. Finally, make sure that the animals do not have environmental enrichment in their home cages such as cardboard shavings, cotton strips, or plastic tubes.
Habituation
Once in the stabling module or room, habituate the animals to the new environment for at least five days before starting the procedures. During this period, handle each animal gently and briefly for 3–5 days and preferably by the same person and/or regular experimenter(s). Furthermore, carry out each manipulation at approximately the same time every day. In this respect, each procedure in our protocol is always done during the light cycle and, more specifically, between 9:30 a.m. and 14:00 p.m., so that both handling and weighing should begin between 9:30 a.m. and 10:30 a.m.
Environmental conditions
Monitor environmental conditions throughout the experiment. In this particular case, animals were kept under a 12:12 h light/dark cycle (lights on at 6:00 a.m.) in a room with constant temperature (21 °C) and humidity (50%–60%).
General and methodological considerations
When experimenting with animal models, always try to reduce the number of animals used and minimize suffering. To do that, we recommend the following:
Experiments should always be carried out by qualified personnel with experience in this type of paradigm.
All manipulations must be carried out at the same time every day and preferably by the same experimenter(s).
Animals should be matched by weight and randomly assigned to the different experimental conditions.
Design the experiments in batches of animals, especially when it has not been implemented before. This way, experimenters can become familiar with the protocol, and both the data and experimental conditions are gradually completed.
Taking these principles into account, also make sure that you have enough statistical power when reporting effects and making inferences.
Induction of the interaction between BE-BD behaviors
Food deprivation
Once the animals are completely habituated to the stabling room as well as to the experimenter(s), the food deprivation process can begin (see Figure 2, sessions 1–3) by weighing and labeling each rat.
On this day, register the initial weight of each animal and remove all food from the cage. After that, return the animals to the stabling room and leave them there until the day of habituation.
Figure 2. Schematic representation of the experimental procedure. Sessions 1–3 comprise the food deprivation period. Session 4 corresponds to the habituation day. Sessions 5–7 refer to the binge eating (BE)-like behavior acquisition period, while sessions 8–14 cover the BE-binge drinking (BD) interaction establishment period. The solid line represents body weight scores, while the long dash line represents the Dustless precision pellets (DPP) consumption (%) exhibited by the rats all along the experimental timeline. The dotted line marks the beginning of the BE-like DPP consumption (i.e., the ingestion of ≥ 80% of the pellets). Extracted and modified from Ruiz-Leyva et al. (2022) with permission from the authors.
Additional tips for the deprivation procedure
In this section you will find some recommendations that will help to improve the quality of this procedure.
Note that the food deprivation procedure allows rats to acclimatize to the experimental room so that there will be no additional sources of stress on the day of habituation (i.e., session 4).
In adult Wistar rats aged 70–80 days (10–11 weeks), it takes approximately three days of complete deprivation to reach 82%–85% of the ad libitum body weight.
During this time (i.e., sessions 1–3), check the health status of rats daily in order to detect some signs of distress such as piloerection, body dirt, or dehydration. In such case, provide five additional grams of food.
As always, try to carry out this procedure at the same time every day.
Considerations prior to the habituation day
In this section, we provide relevant information to optimize time during the habituation day.
The day before of the habituation day (i.e., session 3), clean and arrange as many empty cages (i.e., without sawdust) with grid (hereinafter referred to as BE cages) as rats that can be assessed simultaneously. These are similar to home cages (see Equipment).
Dispose and arrange the essential material for preparing ethanol solutions (i.e., ethanol 96% v/v, syringes, flasks, weighing scale…). Also, store as many bottles of water as needed to fill every anti-drip bottle that will be used in the subsequent two-bottle-choice test.
Habituation day
On the habituation day (i.e., session 4, see Figure 2), prepare fresh ethanol solutions and fill each pair of anti-drip bottles with a given concentration of ethanol or water, as appropriate. Do not forget to attach a label to each pair of bottles with the corresponding rat identifier.
Once the bottles are prepared, weigh each pair of the anti-drip bottles (do it daily), remove every anti-drip daily water bottle from the home cages, and weigh each rat. At this point, check whether every rat is on the fixed weight interval (82%–85%).
Now, arrange the BE cages in a way that: (1) there is space to place a rat cage in between and (2) each BE cage corresponds to a specific rat or rats if using multiple batches (this must remain constant throughout the experiment). Once so disposed, register the time, prepare the stopwatch, and begin the timer as soon as the first rat touches the floor of their corresponding BE cage. After placing each rat, close the cage’s grid and wait for 3 min. After this, remove the rats from their BE cages, put them back into their home cage, and return them to their corresponding position in the racks. If there is urination or defecation in the box, it should be wiped with a 70% alcohol wipe.
Finally, the two-bottle-choice test will take place. To do that, carefully put each pair of anti-drip bottles on each side of the corresponding home cage grid (i.e., ethanol left and water right, or vice-versa) (see Figure 3A and 3B).
Figure 3. Cage and bottle arrangement in the two-bottle-choice test. A. Upper frontal vision. B. Lateral frontal vision. C. Positioning of Dustless precision pellets (DPPs) and rats during the binge eating (BE) episode.
After 90 min, carefully remove each pair of anti-drip bottles. Then, put back the anti-drip daily water bottles in the center of the cage’s grid. Now, weigh again each pair of anti-drip bottles and subtract the loss value of the corresponding ghost bottle (see the section below) from the difference in weights. Finally, feed each rat with the approximate amount of normal chow that it needs to maintain the desired range of body weight and deliver six DPPs to each, in order to avoid neophobia in the following sessions.
Adjustment of the drip loss over the bottle weight difference
The positioning of the bottles during the 90 min two-bottle-choice test (see Figure 3A and 3B) may lead to some very slight losses of contents, which could affect the actual difference in weight (i.e., bottle weight before and after the test). To control this, weigh two extra anti-drip bottles (one filled with the desired ethanol concentration and the other one with water) and place them for 90 min in the grid of an empty rat cage (see Equipment). The positioning of the bottles should be similar to Figure 3.
As a note, these bottles that are meant to control for leakage or spillage are often referred to as ghost bottles.
Desired body weight range considerations and food adjustments
At this point, it is necessary to advise that these considerations depend on the age, sex, strain, and even the supplier, so they must be taken with caution and adjusted for each situation.
It is possible that, at the first weighing after the end of the three days of food deprivation, experimenters may find that not all rats are in the expected range (i.e., 82%–85% of their ad libitum weights). This is perfectly plausible, since the proposed deprivation interval (i.e., three days) for adult Wistar rats of this age and weight is only approximate and is based on previous evidence obtained in our laboratory. Thus, although it is not strictly necessary for all rats to be in this weight range at this moment, it may be relevant when adjusting the amount of food to be administered to each rat at the end of the session (see the section below).
Weight maintenance
Adult Wistar rats of these characteristics (see Materials and reagents) require approximately 15–20 g (±1.5 g) of normal chow to maintain body weight. However, as mentioned above, this may vary depending on two variables:
Current body weight:
• If rats are within the required weight (82%–85%), take the standard feed value (15–20 ± 1.5 g) and adjust the quantity according to the eating condition (binge or control).
• If rats are above the required weight range, reduce* the standard feeding value according to the grams that fall outside the desired range (i.e., upper limit or 85%) and according to the eating condition.
• If rats are below the required weight, increase* the standard value of the feed according to the grams that deviate from the desired range (i.e., lower limit) and according to the intake condition.
An example of weight maintenance is presented below (Table 1).
Table 1. Example of weight maintenance
Initial weight (g) 82% weight (g) Maintenance food (g) control/binge
350 287 16 ± 1.5 g/14 ± 1.5 g
300 246 14 ± 1.5 g/11 ± 1.5 g
280 229 13 ± 1.5 g/10 ± 1.5 g
Eating condition:
Take into account (specially from session 5 onward) that the rats have a differential amount of DPPs depending on the intake group to which they belong (i.e., 72 DPPs for binge and 6 DPPs for control) and will therefore be able to ingest a certain maximum amount of calories during the binge episode (i.e., 11.66 kcal/3 min for the binge group and 0.97 kcal/3 min for the control group)*.
*Note: Only six DPPs are available at the end of the session (delivered to their home cages), but the BE cages remain empty. In this sense, this variable has to be considered, particularly during the rest of the sessions, where they are already exposed to the binge context with their corresponding DPPs.
Prevent side preferences and control for potential order effects
When carrying out the two-bottle-choice test, remember to change the position of the anti-drip bottles every day. Thus, if on the habituation day (i.e., session 4) the ethanol bottle is on the right and the water bottle on the left, the next day (i.e., session 5) the position should be reversed and so on until the end of the procedure.
If you use batches of animals in each BE session, keep in mind that you must counterbalance the order of the batches across the sessions. Let us consider an example in which 20 animals are tested in four batches (1, 2, 3, 4) of five animals. On even days, the first batch would be run first, followed by batches 2–4. However, on odd days, batch four would be run first and batch one would close the experimental day.
Acquisition of BE-like behavior as well as the interaction between BE-BD behaviors
This phase begins 24 h after the habituation session and should last 10 consecutive days (see Figure 2, sessions 5–7 and sessions 8–14). It is similar to the habituation session, yet it features some crucial manipulations.
Once all bottles and rats are all weighed, arrange the BE cages as done in the habituation. Next, place the labeled plastic cups containing different amounts of DPPs (i.e., 72 or 6 DPPs for binge or control eating condition, respectively) next to the BE cages in a way that each cage corresponds to one (or more) cups, depending on whether the experiment is being done in batches or all at once. Now, move each rat to its corresponding BE cage and place the DPPs (without the cup) just in front of the wall furthest from where the rat is left and stacked in the middle (see Figure 3C).
Close each cage’s grid as you place each rat. During the 3 min of the BE episode, please ensure that the rats are not interrupted in their ongoing behavior and keep environmental noise to a minimum.
After 3 min, put each rat back in their home cage and put each pair of anti-drip bottles into their corresponding home cage (do not forget to change daily the position of the pairs in order to prevent side preferences) for 90 min.
At the end of the test, sequentially remove each pair of anti-drip bottles and replace them for the anti-drip daily water bottles. Then, weigh again the bottles and apply the correction to the difference of weights. Finally, feed each animal with its corresponding amount of standard chow.
Current body weight status
Be sure to check whether the rats are within the required weight range (and if possible, close to 82%). This is particularly relevant at this point, as it may determine, in part, how quickly the BE behavior develops.
Collect relevant data concerning the progression of the BE behavior
When the rats have gone through the binge episode, collect the remaining DPPs of each particular cage and put them back into their corresponding and labeled plastic cup. Once you have collected them all, weigh the remaining DPPs of each cup to calculate the number of DPPs ingested (i), the % of DPPs ingested over total (ii), and the kcal/3 min for each rat (iii).
DPP ingested: You can calculate the number of DPPs consumed based on the remaining weight. Given that the maximum weight for the binge group is 3.24 g (i.e., 72 DPPs) and for the control group is 0.27 (i.e., 6 DPPs), you only have to subtract the remaining weight in the cup.
For example: if a rat has given up 1.80 g of DPPs during the binge eating episode and the maximum possible is 3.24 g, equal to 72 DPPs, then: (1.80 × 72)/3.24 = 40. This tells us how many DPPs it has given up without consuming, so: 72 - 40 = 32.
*To apply this example to the control intake group, just modify the value of the maximum weight of DPPs to be consumed (0.27 g instead of 3.24 g) and the maximum number of pellets (from 72 DPPs to 6 DPPs)
% DPPs consumed: To get the ratio or percentage of DPPs consumed, you simply divide the number of DPPs ingested by the total (i.e., 72 DPPs for the binge group) and multiply this value by 100.
Based on the previous example, if the animal has consumed 32 DPPs and the total is 72, then: (32/72) × 100 = 44.4%.
Kcal ingested in 3 min: multiply the number of DPPs consumed by the caloric value of each DPP (0.162 kcal). Returning to the initial example, if the animal consumes 32 DPPs, then: 32 × 0.162 = 5.18 kcal/3 min.
Data analysis
Exclusion and inclusion criteria
Binge eating behavior progression:
As mentioned in Ruiz-Leyva et al. (2022), out of a total of 142 adult male Wistar rats used for the experiments, seven were eliminated from the subsequent analyses because they did not acquire the BE behavior (i.e., they completely avoided the DPPs). In this regard (and at least when using males), make sure that rats exhibit the BE behavior from day 4 (i.e., session 8) onwards (i.e., ingestion of ≥ 80% DPPs) or at least show signs that DPP intake is significantly increasing and consistently approaching this percentage.
Preliminary, unpublished data from our laboratory indicate that this is an achievable task for adult male rats, but ingestion must be performed rapidly and without interruption. Furthermore, we have observed that male rats show an average intake of 80% of the available DPP even when adulterated with a bitter flavor (quinine, see Figure 4).
Figure 4. Progression in the consumption of palatable sugar pellets [i.e., Dustless precision pellets (DPPs)] of adult male Wistar rats assigned to the binge eating condition (i.e., ingestion of 72 DPP or 11.66 kcal/3 min). Percent of DPP intake (% DPPs consumed over total) across sessions of the values obtained in 7–9 animals. Dashed lines mark the percentage of DPP consumption in the binge eating range model.
Ethanol consumption
Researchers should check the ethanol consumption exhibited by each rat even when they exhibit binge eating behaviors towards sugary pellets. This is due to the existence of rats that completely avoid alcohol consumption and the fact that binge eating is motivated, in part, by calorie restriction. To this end, use statistical criteria for detecting potential outliers. Finally, note that opposite the big amount of ethanol consumed, no taste aversion is development to the pellets (Gallo et al., 1999).
Data calculation
Calculation of the main dependent variable data
There are three dependent variables whose units you can (and should) obtain from the habituation day (i.e., session 4). These are (1) the weight-adjusted water consumption (grams per kilogram of body weight), (2) the weight-adjusted net ethanol consumption (grams of ethanol per kilogram of body weight), and (3) the percentage preference of ethanol over water.
First of all, obtain the difference in weights of the same bottle before (i.e., first weight or PRE) and after the two-bottle-choice test (i.e., second weight or POST) within the same session. To this difference, apply the correction for possible fluid loss.
Water consumption:
Ethanol intake:
See Figure 5 for a complete overview of the results.
Figure 5. Ethanol intake exhibited by male Wistar rats as a function of ethanol concentration (0%, 2%, 6%, 10%, or 14% w/w), eating condition [binge or control Dustless precision pellets (DPP) exposure], and session. (A) and (B) represent ethanol consumption (net gEtOH/kg) in the acquisition phase (i.e., sessions 5–7) and in the establishment phase (i.e., sessions 8–14), respectively. (C) Depicts the same data as (A) and (B) yet collapsed across sessions. Each point or bar and vertical line represent the mean ± SE obtained in 7–12 animals per group. (A and B) Statistically significant differences between the values obtained in binge and control groups: *p < 0.05, **p < 0.01; and between the values obtained in binge groups compared with 10% ethanol concentration: #p < 0.05, ##p < 0.01. (C) Statistically significant differences between the values obtained in binge and control groups in each ethanol concentration: &&p < 0.01; and between the values obtained in binge groups compared with 10% ethanol concentration: $$p < 0.01. Extracted with permission from Ruiz-Leyva et al. (2022).
Preference scores for ethanol over water:
Notes
Up to this point, we have described in detail every aspect of the model and the protocol that has been implemented in our laboratory and that has given us the best convergence of results to date. However, there are certain factors that need to be clarified to improve the reproducibility of the model.
Age and sex
In the published article, all male rats used were young adults (i.e., PD70-80). In the case of using rats in other age range or female rats, it may be necessary to adapt the procedure. Of course, if after day 3 of habituation the animal is below 82%, the amount of food can be increased by 5 g to increase the animal’s weight by 2 g.
Strain differences
To date, we have only used Wistar HAN rats. Despite this, it would be interesting to test the model in other strains or in different intraspecies phenotypes that are more or less vulnerable to relevant variables such as stress, or to psychoactive effects of drugs such as sugar or alcohol.
Ethanol concentration
As shown in Figure 5, ethanol intake exhibited by males is concentration dependent, with 6% and 10% (w/w) being the most consumed.
Food composition
Knowing the macro- and micronutrient composition of the food used to induce binge eating behavior is essential to determine which elements may contribute to it and interact with alcohol and/or its effects. The palatable pellets (i.e., DPPs) used here have a high carbohydrate value (i.e., 59.1%) and sugary taste. However, it would be interesting to try other types of foods (e.g., high fat or mixed) or to try artificial sweeteners (e.g., saccharin) with a much lower caloric value. In this way, the mechanisms underlying the interaction between these addictive behaviors and between the substances involved could be understood in more detail.
Acknowledgments
This paper was partially supported by the Spanish Ministry of Health (Government Delegation for the National Plan on Drugs: PNSD 2020-049), the Junta de Andalucía (grants CTS109 and HUM784), and the University of Granada (PP2022.PP-16). The work was also supported by PICT-2019- 00180 and PICT-2018-00597 of FONCyT-Argentina. This protocol and figures are extracted and/or modified from our previous work (Ruiz-Leyva et al., 2022).
Competing interests
The authors declare no conflicts of interest or competing interests.
Data availability statement
The data from the published paper are available from the corresponding author (see Ruiz-Leyva et al., 2022) upon reasonable request.
Ethics considerations
Animals were maintained according to the EU Directive 2010/63/EU for animal experiments. The experimental protocol was approved by the University of Granada Research Ethics Committee (Protocol Number 09/08/2019/138).
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Fabrication of Microfluidic Devices for Continuously Monitoring Yeast Aging
RO Richard O’Laughlin
EF Emerald Forrest
JH Jeff Hasty
NH Nan Hao
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4782 Views: 429
Reviewed by: Chiara AmbrogioPrajita PandeyShun Yu Jasemine Yang
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Original Research Article:
The authors used this protocol in eLIFE Oct 2022
Abstract
For several decades, aging in Saccharomyces cerevisiae has been studied in hopes of understanding its causes and identifying conserved pathways that also drive aging in multicellular eukaryotes. While the short lifespan and unicellular nature of budding yeast has allowed its aging process to be observed by dissecting mother cells away from daughter cells under a microscope, this technique does not allow continuous, high-resolution, and high-throughput studies to be performed. Here, we present a protocol for constructing microfluidic devices for studying yeast aging that are free from these limitations. Our approach uses multilayer photolithography and soft lithography with polydimethylsiloxane (PDMS) to construct microfluidic devices with distinct single-cell trapping regions as well as channels for supplying media and removing recently born daughter cells. By doing so, aging yeast cells can be imaged at scale for the entirety of their lifespans, and the dynamics of molecular processes within single cells can be simultaneously tracked using fluorescence microscopy.
Key features
• This protocol requires access to a photolithography lab in a cleanroom facility.
• Photolithography process for patterning photoresist on silicon wafers with multiple different feature heights.
• Soft lithography process for making PDMS microfluidic devices from silicon wafer templates.
Keywords: Aging Yeast replicative aging Saccharomyces cerevisiae Microfluidics Microfabrication Photolithography Soft lithography Polydimethylsiloxane (PDMS)
Background
Progress in aging research has been greatly accelerated in recent years due to the development of new tools and techniques for single-cell analysis (Dulken et al., 2019; Li et al., 2020; Tabula Muris, 2020; Trapp et al., 2021; Roux et al., 2022). Microfluidic technologies play pivotal roles in these applications, as they allow single cells to be captured in droplets for sequencing (Matuła et al., 2020) or isolated for long-term imaging (Allard et al., 2022). For studying replicative aging in Saccharomyces cerevisiae, the traditional method of manual microdissection of mother and daughter cells in order to count replicative lifespan has been largely supplanted by the use of microfluidic devices, which efficiently trap individual mother cells in place while removing newly budded daughter cells [reviewed in Chen et al. (2017); Gao et al. (2020); O’Laughlin et al. (2020)]. Since mother cells growing in microfluidic devices can be imaged at regular intervals, and various molecular and biochemical processes can be monitored in real-time with fluorescence microscopy, this approach has led to a number of significant advances in understanding yeast aging (O’Laughlin et al., 2020). Prominent among these is the discovery of two divergent trajectories that single cells take as they age, which are marked by distinct morphological features and the breakdown of different cellular functions (Li et al., 2017 and 2020; Jin et al., 2019; Paxman et al., 2022). Recently, our group has used these devices to reveal that cells undergoing one of the aging trajectories display a reduction in protein homeostasis, with RNA binding proteins aggregating after diminished control of chromatin silencing at the rDNA locus (Paxman et al., 2022). Here, we report a detailed protocol for constructing the microfluidic devices for yeast replicative aging experiments that have facilitated the discovery and characterization of these trajectories.
Materials and reagents
100 mm diameter silicon wafers (University Wafer, catalog number: 452)
SU-8 2005 (Kayaku Advanced Materials)
SU-8 2010 (Kayaku Advanced Materials)
SU-8 2015 (optional) (Kayaku Advanced Materials)
SU-8 Developer (Kayaku Advanced Materials)
Isopropanol (e.g., Spectrum Chemical, catalog number: 67-63-0)
Acetone (e.g., Spectrum Chemical, catalog number: 67-64-1)
Ethanol (e.g., Koptec, catalog number: 64-17-5)
Methanol (e.g., Sigma-Aldrich, catalog number: 179337)
Heptane (e.g., Sigma-Aldrich, catalog number: 494526)
Deionized water (from purification system, e.g., Thermo Scientific, model: 7143)
Trichloro(1H,1H,2H,2H-perfluorooctyl)silane (Fisher Scientific, catalog number: 448931-10G)
Chrome glass masks, custom-made according to the user’s AutoCAD design file and ordered from a company such as HTA Photomask (https://htaphotomask.com/)
Pyrex dish for development (Fisher Scientific, Pyrex, catalog number: 08-741F)
Pyrex dish for wafer handling (Fisher Scientific, Pyrex, catalog number: 08-747E)
Plastic weighing dishes (e.g., Fisherbrand, Fisher Scientific, catalog number: S67090A)
SYLGARD 184 Silicone Elastomer kit (PDMS) (Dow, catalog number: 2646340 or purchase from Fisher Scientific, catalog number: NC9285739)
Kapton tape (e.g., Uline, Fisher Scientific, NC0912751)
Adhesive labels (e.g., Fisher Scientific, catalog number: 15922)
Plastic bottles for aliquoting photoresist (e.g., Thermo Scientific Nalgene, Fisher Scientific, catalog number: 02-925-3C)
Microcentrifuge tubes (e.g., Thomas Scientific, Fisher Scientific, catalog number: NC9448938)
Scotch Magic tape (3M, catalog number: B0000DH8HQ)
Aluminum foil (e.g., Thomas Scientific, catalog number: 1181K86)
Single edge razor blades (e.g., Fisher Scientific, Fisherbrand, catalog number: 12-640)
0.5 mm biopsy puncher (World Precision Instruments, catalog number: 504528)
Cutting pad (e.g., Qiagen Harris Cutting Mat, catalog number: WB100020)
Cover glass slides 45 × 50 mm (Oracle, Brain Research Laboratories, catalog number: 4550-1.5D)
Glass stir rod (e.g., Millipore Sigma, catalog number: Z549768)
Timer (e.g., Fisher Scientific, Fisherbrand, catalog number: 14-649-17)
Equipment
Wafer tweezers (e.g., Electron Microscopy Sciences Rubis Style 39S-4, Fisher Scientific, catalog number: 50-239-33)
Tweezers (e.g., for an assorted set, Kaisi, catalog number: B07GLJ7627)
Scissors (any generic brand, e.g., Fisher Scientific, catalog number: S173182)
Spin coater (e.g., Laurell Technologies Corporation, model: WS-650S-6NPP/UD2)
Wafer alignment tool (Laurell Technologies Corporation)
Mask aligner (e.g., EV Group, model: EVG620 Lithography/NIL System)
Hot plates (e.g., Wenesco)
Nitrogen spray gun (supplied in cleanroom facilities, e.g., Terra Universal, catalog number: 2002-21)
Profilometer (e.g., Veeco, Dektak 150)
Upright microscope (e.g., Zeiss, Axio Imager)
Vacuum desiccator (e.g., Nalgene, Thermo Fisher Scientific, catalog number: 5310-0250)
Stereomicroscope (e.g., Amscope, catalog number: SM-4B)
UVO-Cleaner (Jelight, model: 42)
Oven (e.g., Fisher Scientific, catalog number: 13246516GAQ)
Software
AutoCAD from AutoDesk (https://www.autodesk.com/products/autocad). Requires license but this is free to download for students and educators.
Procedure
Obtain chrome glass masks for photolithography
As part of this protocol, we have created an AutoCAD file with only the designs for the microfluidic devices used in Paxman et al. (2022) and included it in the Supplement. See Figure 1 for the design of the microfluidic device. Email the .dwg AutoCAD file to a company such as HTA photomask, which makes chrome glass masks. Order two 5 × 5 inch chrome photomasks made of quartz glass. Use ± 0.25 μm tolerance for the first mask containing the cell traps and ± 0.5 μm tolerance for the second mask containing the media channels. Have the closed objects on the .dwg file printed as clear on the mask and have the printing as right reading down with the chrome side facing down.
Figure 1. Layout and design of yeast aging microfluidic device. (A) Screenshot of the AutoCAD design file of the device. (B) Schematic of device layout with the inlet ports and outlet/waste ports labeled. The inlet ports are where cells are loaded into the device and afterward a fresh syringe with media is connected. Four units comprise a single device. (C) Single unit with 100 cell traps. Cell traps were designed to be 4.5 μm tall and media channels were designed to be 15 μm tall. D. Dimensions of cell traps.
Prepare the wafer for photolithography
Cleaning the wafer (optional):
Sonicate the wafer in acetone for 15 min.
Rinse wafer with methanol and then sonicate in methanol for 5 min.
Rinse wafer with isopropanol and then sonicate in isopropanol for 5 min.
Rinse wafer with deionized water.
Note: While this step may be useful for certain kinds of silicon wafers, we have found it to be unnecessary if using the wafers included in the Materials and Reagents section (100 mm wafers). Therefore, if using these wafers, we recommend skipping this step.
Dehydration bake:
Bake the wafer at 150 °C for 15 min followed by a 15 min cool down to room temperature (RT).
Photolithography for cell trapping layer (Layer 1)
See Figure 2 for an overview of the process for Layer 1.
Figure 2. Overview of steps for photolithography process of Layer 1 for the cell trapping layer. (A) The wafer is placed on the spinner chuck using the alignment tool. This ensures that the wafer is centered on chuck. (B) SU-8 2005 is poured on the wafer, covering its entire surface. (C) Spin coating of the wafer with photoresist. (D) Soft bake of the wafer. (E) The mask is loaded onto the EVG620 lithography system as shown, followed by the wafer. (F) Recipe used on the EVG620 lithography system showing important parameters for the process for Layer 1, including the use of vacuum contact and an exposure dose of 75 mJ/cm2. (G) Post-exposure bake of the wafer. (H) Development of the wafer in SU-8 Developer to remove photoresist that has not been crosslinked via ultraviolet (UV) light exposure. (J) To complete the process, the wafer is hard baked at 95 °C for 5 min.
Lift the spin coater lid and use the wafer alignment tool to place the wafer on the center of the chuck.
Flood the wafer with SU-8 2005 photoresist so that the entire wafer is covered in resist.
Set the time on the spin coater to 40 s and set the spin speed to 3,000 rpm. For the acceleration, select the preset value that is closest to 3,000 rpm/s, so that the wafer will reach its desired spin speed in 1 s.
Note: We note that these spin parameters are unconventional, as data sheets for SU-8 2000 series photoresists recommend a first step at a speed of 500 rpm and acceleration of 100 rpm/s. However, since the target height of the cell traps is 4.5 μm, we found it more reliable to achieve this height by omitting this first step and spinning immediately at the high speeds than by following the recommended protocol from the manufacturer, in which we found it difficult to reliably spin this photoresist down to less than 5 μm. Therefore, in our experience, modulating the initial acceleration is a powerful method for tuning the photoresist height.
When spin coating has been completed, soft bake the wafer at 65 °C for 9 min. Place the lid of a Pyrex dish over the wafer while it is on the hot plate. Position the lid so that it slightly hangs over the end of the hotplate. When time is up, remove the Pyrex dish lid and use wafer tweezers to remove the wafer from the hot plate. Place the wafer on the lid and allow it to cool down to RT for 3 min.
Note: Placing the lid over the wafer is done to give the wafer a surface of equal temperature to cool on, so that it does not cool too fast. This is done for all baking steps in this protocol.
Expose the wafer to UV light on the EVG620 using the Layer 1 chrome glass mask without a filter in place at 75 mJ/cm2 using the vacuum contact setting.
Remove the wafer from the mask aligner.
Perform a 2 min post-exposure bake at 95 °C. Arrange the Pyrex dish lid over the wafer as before. Allow 3 min to cool down to RT.
Pour SU-8 Developer in a large Pyrex dish and develop the wafer for 2 min. During this time, gently shake the dish back and forth and side to side. After 2 min, take the wafer out of the dish and rinse it with fresh developer for ~10 s; then, rinse with isopropanol for ~20 s. Blow the wafer dry using a nitrogen spray gun.
Hard bake the wafer at 95 °C for 5 min with a 3 min cool down to RT afterward.
Measure the height of the layer using a profilometer and assess feature integrity under a microscope. While there is inherent variability in the spin coating process, it is important to verify that the measured height is close to that of the design.
Note: When assessing the first layer height and feature integrity under an upright microscope, ensure that trap dimensions are close to those shown in Figure 1C and 1D and that there are no cracks or dents in the photoresist layer. Successfully built wafers should have dimensions within ±10% of the design specifications for the first layer.
Photolithography for media channel layer (Layer 2)
Tape over the alignment markers on the wafer (Figure 3). To do this, cut out a piece of Kapton tape and cut out a small square from the adhesive labels that is large enough the cover the alignment markers. Using tweezers, place the sticky side of the cut-out label square onto the sticky side of the Kapton tape. Place this over the alignment markers on the wafer so that the non-sticky side of the label is covering the alignment markers. Use a tweezer to press down on the tape surrounding the alignment markers so that it sticks to the wafer. Trim Kapton tape hanging off the wafer but leave enough excess to grab with a tweezer (tape will be removed after spin coating).
Figure 3. Taping over alignment markers on Layer 1. (A) Location of alignment markers on wafer, including a zoomed in image (top right). (B) Tape setup for covering alignment markers. (C) View of loading wafer onto the spin coater with alignment markers covered with tape. (D) Pouring on second layer photoresist, SU-8 2010. (E) Removal of tape covering alignment markers after spin coating.
Pour approximately 4 mL of SU-8 2010 onto the wafer.
Set the time on the spin coater to 40 s and set the spin speed to 1,450 rpm. For the acceleration, select the preset value that is closest to 1,450 rpm/s, so that the wafer will reach its desired spin speed in 1 s.
Note: In the original build for this wafer, these settings were used to maintain consistency with the method of spinning the first layer. However, in later builds we have switched to a more standard approach for achieving a 15 μm layer by using SU-8 2015 and spinning in two steps: step 1 is at 500 rpm with a 136 rpm/s acceleration for 10 s, and step 2 at 3,000 rpm with a 272 rpm/s acceleration for 40 s. We recommend this latter approach for the build.
With a tweezer, carefully remove the tape over the alignment markers on the wafer.
Soft bake the wafer at 65 °C for 15 min with a 3 min cool down to RT afterward.
Alignment of Layer 1 to the mask for Layer 2:
See Figure 4 for the design of the alignment markers, the goal of the alignment process, and example images during alignment. Align Layer 1 and Layer 2 alignment markers by adjusting the X, Y, and Theta knobs on the EVG620 mask aligner. The most efficient way to accomplish this is to first tune the focus and position of the lenses on the EVG620 to locate the alignment markers on the Layer 2 mask. Then, turn the knobs to locate the alignment markers on the wafer. Then, to align the two sets of markers, correct half in the Y direction and half in the Theta direction until all sets of squares are fully aligned (see Figure 4). Perform any intermittent corrections in the X direction as needed during this process.
Figure 4. Alignment of Layer 1 and Layer 2. (A) Schematic of the final goal of the alignment process with a single set of alignment markers on the wafer and on the Layer 2 mask shown correctly aligned. (B) Progressing views of alignment process. Markers on the wafer and the mask begin out of alignment (top) but are gradually put into correct alignment by tuning the X, Y, and Theta knobs on the mask aligner. The largest markers are aligned first (middle), followed by further improvement of alignment by correctly orienting the smallest set of markers (bottom), thereby completing the alignment process.
Expose the wafer on the EVG620 using the Layer 2 chrome glass mask without a filter at 125 mJ/cm2 using the hard contact setting.
Post-exposure bake at 95 °C for 5 min with a 3 min cool down to RT.
Pour fresh SU-8 Developer into the large Pyrex dish and develop the wafer for approximately 5 min with gentle shaking. Rinse with fresh developer and then isopropanol and blow dry with a nitrogen spray gun.
Hard bake at 95 °C for 5 min with a 3 min cool down to RT.
Measure the height of the media channels with a profilometer and assess feature integrity of the final wafer using an upright microscope (Figure 5A and 5B).
Note: Ensure that the photoresist layer is free of large cracks and dents when analyzing the wafer under an upright microscope. Error tolerability in height for the second layer is larger than that for the first; however, aim for ±20% from the design specifications. Importantly, make sure the traps are well aligned between the channels as in Figure 5B.
Figure 5. Final wafer, silanization, and application of the polydimethylsiloxane (PDMS) microfluidic device. (A) Fully constructed wafer. (B) Check the alignment between the two layers of the final wafer under a microscope. (C) Wafer placed in the silanization chamber with two Eppendorf tubes, each containing 20 μL of silane placed on either side. Black arrows show the placement of the tubes. (D) Wafer in the closed silanization chamber with the vacuum pump turned on. (E) Phase contrast image from Paxman et al. (2022) of cells growing in the PDMS microfluidic device (scale bar = 10 μm). (F) Fluorescence image from Paxman et al. (2022) of cells with a nuclear iRFP marker growing in the PDMS microfluidic device (scale bar = 10 μm).
Wafer silanization
In a chemical fume hood, place the wafer in a vacuum desiccator and place in two microcentrifuge tubes. Pipette 20 μL of trichloro(1H,1H,2H,2H-perfluorooctyl)silane into each tube (Figure 5C). Start the vacuum and allow the wafer to be exposed to the silane for 7 min (Figure 5D). For more details on our vacuum desiccator setup, see Ferry et al. (2011). Caution: Silane is toxic, and this step must be done within a chemical fume hood with appropriate personal protective equipment.
Soft lithography using PDMS
Weigh out 30 g of SYLGARD 184 Base; then, in the same plastic weighing dish, add 3 g of SYLGARD 184 curing agent. Thoroughly mix together with a glass stir rod.
Place PDMS mixture in a vacuum desiccator for approximately 30 min to remove bubbles.
Wrap the wafer in aluminum foil so that the aluminum foil forms a bowl around the wafer and the wafer is sitting flat.
Pour PDMS onto the wafer.
Place the wafer back in the vacuum chamber for approximately 30 min or until all bubbles have been cleared.
Bake the PDMS at 80–90 °C for at least 1 h (overnight optional).
Use a razor blade to remove aluminum foil from the back of the wafer. Gently peel up the aluminum foil and PDMS on the flat edge of the wafer and peel off the PDMS.
Cut out individual devices with a razor blade.
Use a 0.5 mm biopsy puncher to punch the inlet and outlet ports on each device under a stereomicroscope.
Bonding PDMS devices to glass coverslips
Rinse PDMS devices with ethanol and then deionized water. Blow dry with a nitrogen or air spray gun.
Clean the PDMS devices by placing a piece of Scotch tape on the feature side of the PDMS device and using the blunt end of a tweezer to go over the tape in a back-and-forth motion. Apply enough pressure to ensure that the tape works its way into the features and withdraws any dust. Repeat this process at least four times on the feature side and at least once on the non-feature side.
Clean glass coverslips with heptane. Rinse with methanol and then with deionized water. Blow dry with a nitrogen or air spray gun.
Turn on the UVO-Cleaner and run it for 5 min without any samples in.
Place the cleaned glass slides and PDMS devices, feature side facing up, into the UVO-Cleaner and run for 3 min.
Flip PDMS devices over and place the feature sides of the devices in contact with the glass slides.
Place devices in an 80–90 °C oven overnight to bond. After this point, devices should be used for experiments within two weeks [see Paxman et al. (2022) for details on experimental setup]. As a reference, yeast cells growing in the device can be seen in Figure 5E and 5F.
Validation of protocol
Paxman et al. (2022). Age-dependent aggregation of ribosomal RNA-binding proteins links deterioration in chromatin stability with challenges to proteostasis. eLife (Figure 1, panels A–D; Figure 3, panels A–C; Figure 4, panels A–D; Figure 5, panels A and B; Figure 6, panel B).
Acknowledgments
This work was performed in part at the San Diego Nanotechnology Infrastructure (SDNI) of UCSD, a member of the National Nanotechnology Coordinated Infrastructure (NNCI), which is supported by the National Science Foundation (Grant ECCS-2025752). This work was supported by NIH R01 AG068112 (to NH), AG056440 (to NH, JH), GM144595 (to NH, JH), and GM111458 (to NH).
This protocol was derived from the original work of Paxman et al. (2022).
Competing interests
The authors declare no competing interests.
References
Allard, P., Papazotos, F. and Potvin-Trottier, L. (2022). Microfluidics for long-term single-cell time-lapse microscopy: Advances and applications. Front. Bioeng. Biotechnol. 10: e968342.
Chen, K. L., Crane, M. M. and Kaeberlein, M. (2017). Microfluidic technologies for yeast replicative lifespan studies. Mech. Ageing Dev. 161: 262–269.
Dulken, B. W., Buckley, M. T., Navarro Negredo, P., Saligrama, N., Cayrol, R., Leeman, D. S., George, B. M., Boutet, S. C., Hebestreit, K., Pluvinage, J. V., et al. (2019). Single-cell analysis reveals T cell infiltration in old neurogenic niches. Nature 571(7764): 205–210.
Ferry, M., Razinkov, I. and Hasty, J. (2011). Microfluidics for Synthetic Biology. Meth. Enzymol. 497: 295–372.
Gao, Z., Xu, J., Chen, K., Wang, S., Ouyang, Q. and Luo, C. (2020). Comparative Analysis of Yeast Replicative Lifespan in Different Trapping Structures Using an Integrated Microfluidic System. Adv. Mater. Technol. 5(12): 2000655.
Jin, M., Li, Y., O’Laughlin, R., Bittihn, P., Pillus, L., Tsimring, L. S., Hasty, J. and Hao, N. (2019). Divergent Aging of Isogenic Yeast Cells Revealed through Single-Cell Phenotypic Dynamics. Cell Syst. 8(3): 242–253.e3.
Li, Y., Jiang, Y., Paxman, J., O’Laughlin, R., Klepin, S., Zhu, Y., Pillus, L., Tsimring, L. S., Hasty, J., Hao, N., et al. (2020). A programmable fate decision landscape underlies single-cell aging in yeast. Science 369(6501): 325–329.
Li, Y., Jin, M., O’Laughlin, R., Bittihn, P., Tsimring, L. S., Pillus, L., Hasty, J. and Hao, N. (2017). Multigenerational silencing dynamics control cell aging. Proc. Natl. Acad. Sci. U.S.A. 114(42): 11253–11258.
Matuła, K., Rivello, F. and Huck, W. T. S. (2020). Single‐Cell Analysis Using Droplet Microfluidics. Adv. Biosyst. 4(1): 1900188.
O’Laughlin, R., Jin, M., Li, Y., Pillus, L., Tsimring, L. S., Hasty, J. and Hao, N. (2020). Advances in quantitative biology methods for studying replicative aging in Saccharomyces cerevisiae. Transl. Med. Aging 4: 151–160.
Paxman, J., Zhou, Z., O’Laughlin, R., Liu, Y., Li, Y., Tian, W., Su, H., Jiang, Y., Holness, S. E., Stasiowski, E., et al. (2022). Age-dependent aggregation of ribosomal RNA-binding proteins links deterioration in chromatin stability with challenges to proteostasis. eLife 11: e75978.
Roux, A. E., Yuan, H., Podshivalova, K., Hendrickson, D., Kerr, R., Kenyon, C. and Kelley, D. R. (2022). The complete cell atlas of an aging multicellular organism. bioRxiv: e496201.
Tabula Muris, C. (2020). A single-cell transcriptomic atlas characterizes ageing tissues in the mouse. Nature 583(7817): 590–595.
Trapp, A., Kerepesi, C. and Gladyshev, V. N. (2021). Profiling epigenetic age in single cells. Nat. Aging 1(12): 1189–1201.
Supplementary information
The following supporting information can be downloaded here:
The AutoCAD file with the designs for the microfluidic devices used in Paxman et al. (2022).
Article Information
Copyright
© 2023 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
Developmental Biology > Cell growth and fate > Ageing
Cell Biology > Cell imaging > Live-cell imaging
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SIMBA Method—Simultaneous Detection of Antimicrobial and Anti-biofilm Activity of New Compounds Using Salmonella Infantis
MS Meta Sterniša
JS Jerica Sabotič
NJ Nika Janež
TC Tomaž Curk
AK Anja Klančnik
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4783 Views: 599
Reviewed by: Shailesh Kumar Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in International Journal of Food Microbiology Mar 2022
Abstract
The development of antimicrobial resistance and the formation of Salmonella biofilms are serious public health problems. For this reason, new natural compounds with antimicrobial and anti-biofilm activity are being sought, and wild fungi represent an untapped potential. Various extraction agents, including organic solvents and aqueous buffers, can be used to obtain bioactive compounds from natural sources. To evaluate their bioactivity, extensive screening studies are required to determine antimicrobial and anti-biofilm activity using methods such as broth microdilution or crystal violet assay, respectively, but none of these methods allow simultaneous evaluation of both activities against bacteria. Cold water extraction from wild fungi offers the advantage of extracting water-soluble compounds. The SIMultaneous detection of antiMicrobial and anti-Biofilm Activity (SIMBA) method combines the testing of both types of activity against bacteria with the evaluation of the 20 h growth curve of the Salmonella Infantis ŽM9 strain determined with absorbance measurements at 600 nm in a 96-well plate. SIMBA method thus shortens the time to determine the bioactivity of extracts, reduces material consumption, and eliminates the need for additional reagents. SIMBA enables rapid selection of bioactive extracts for their fractionation and shortens the time to determine new natural products with antimicrobial and anti-biofilm activity.
Graphical overview
Keywords: Salmonella Infantis Antimicrobial Anti-biofilm Growth curve Absorbance New compounds High-throughput Screening
Background
Biofilm formation is a persistence strategy for bacteria that plays a role both for biotic surfaces causing chronic infections and for colonization and survival on abiotic surfaces. After attachment to different surfaces, Salmonella spp. can form a protective matrix of extracellular polymeric substances (EPS) that protects bacteria from environmental factors, facilitates gene transfer, and increases nutrient availability and bacterial resistance to antimicrobial agents. Therefore, Salmonella biofilm cells exhibit higher environment survival and increased antimicrobial resistance, both of which are associated with more human infections (Steenackers et al., 2012; MacKenzie et al., 2017; González et al., 2018; Tassinari et al., 2019). In addition, antimicrobial resistant Salmonella are widespread in animals and on various surfaces associated with animal production and food processing (Vestby et al., 2009; Pate et al., 2019; Tassinari et al., 2019). Alternative control strategies for these Salmonella are being sought due to their resistance to environmental stress and to commercial antibacterial agents.
Higher fungi represent an underexplored source of antimicrobial compounds; fruiting bodies of cultivated and wild mushrooms have been shown to contain various antimicrobial compounds, ranging from secondary metabolites such as steroids, terpenes, and quinolines, to peptides and proteins (Alves et al., 2012; Erjavec et al., 2012). Different solvents have been used for the extraction of new potential antimicrobial compounds. Organic solvents such as methanol, ethanol, ethyl acetate, chloroform, acetone, ether, xylene, cyclohexane, or dichloromethane are the most commonly used, while aqueous extraction with hot water or cold aqueous buffers is rarely used (Alves et al., 2012; Klančnik et al., 2017; Gómez Román et al., 2020). The advantage of using water for extraction is that water-soluble compounds can be extracted, which is beneficial for their potential use as antimicrobials. Water solubility is a very important parameter for drug formulations, as poorly soluble drugs are associated with poor absorption and low bioavailability and often require higher doses (Savjani et al., 2012).
Antimicrobials inhibit bacterial growth or kill bacteria, while specific anti-biofilm agents can interact with biofilm-related metabolic processes without affecting bacterial growth (Steenackers et al., 2012). These two concepts must be considered separately, as compounds that interfere with growth may also contribute to faster development of bacterial resistance. Because of their importance in bacterial persistence, there is a need to control biofilms and their formation without affecting bacterial growth (Dieltjens et al., 2020). Currently, different methods are used to test antimicrobial activity and anti-biofilm activity. Methods performed in microtiter plates are based on similar principles, and determination of minimum inhibitory concentration in the form of broth microdilution is used to evaluate antimicrobial activity. Determination of anti-biofilm activity requires more manipulation because the biofilm is formed on the surface, and therefore steps to wash and detach the cells are required, followed by determination of the culturable cells by dilution and plating. In addition, various indicators can be used as colorimetric indicators of microbial metabolic activity of microorganisms to evaluate viability in antimicrobial activity analysis, for example, tetrazolium salts such as 2,3,5-triphenyltetrazolium chloride (TTC), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenlytetrazolium bromide (MTT), or 2-(4-iodophenil)-3-(4-dinitrophenyl)-5-phenyltetrazolium chloride (INT). Resazurin can also be used to determine the viability of microorganisms, measured as a fluorescence signal or with some commercially available kits such as PrestoBlueTM. The bioluminescence signal can also be used to determine the ATP activity of microorganisms using commercially available kits such as BacTiter-GloTM. Similarly, dye assays can also be used to assess anti-biofilm activity, such as crystal violet, safranin, or acridine orange (Klančnik et al., 2010 and 2021; Eloff, 2019).
All of these methods require manipulation after initial incubation; some use reagents that are hazardous to the environment, and two methods must be performed to evaluate both activities. Here, we present a method for SIMultaneous detection of antiMicrobial and anti-Biofilm Activity (SIMBA) published in Sterniša et al. (2022a). It is a 96-well microplate-based method that, compared to other antimicrobial and anti-biofilm assays, does not require additional plate manipulations after plate preparation, does not require reagents, and provides results within 20 h. We additionally provide the workflow to predict antibacterial or anti-biofilm activity using freely available software Orange (Demšar et al., 2013). A learner on predicting the activity of new compounds is trained on a provided dataset consisting of experimentally verified data and then a workflow is provided to predict the activity of the unknown compound. This presents a step forward to analyzing growth curves by advanced algorithms further supporting high-throughput methods like SIMBA. Namely, SIMBA was shown to reduce costs by up to 50% and time to completion in the laboratory by up to 80%. Interpretation of results is based on a computer program, making it objective and straightforward. This protocol can be used for screening extracts for novel antimicrobial and/or anti-biofilm compounds, as well as for bioactivity-guided purification of such compounds. Other areas where this protocol can be useful for faster analysis of antimicrobial and anti-biofilm activities include testing of commercial products such as food or feed additives, pesticides, biocides, or disinfectants and wastewater analysis.
Materials and reagents
Ice
600 mL glass beaker (Brand, catalog number: BR90648)
Disposable viscose cloths (Tosama, catalog number: 23073)
Large potato ricer
HiPrep 26/60 Sephacryl® S-200 HR column (Cytiva, catalog number: 17-1195-01)
Liquid chromatography column 3 × 100 cm (custom made)
0.2 μm syringe filters (Minisart NML, Sartorius, catalog number: 16534)
10 mL syringe (BD Emerald, catalog number: 307736)
50 mL centrifuge tube (Sarstedt, catalog number: 62.547.004)
Transparent flat-bottomed 96-well microplates (Thermo Scientific Nunc, catalog number: 266120)
15 mL centrifuge tube (Sarstedt, catalog number: 62.554.502)
1.5 mL microcentrifuge tubes (Eppendorf, catalog number: 0030 120.086)
1–10 mL tips (Eppendorf, catalog number: 0030 000.765)
50–1,000 μL tips (Eppendorf, catalog number: 0030 000.927)
2–200 μL tips (Eppendorf, catalog number: 0030 000.870)
Cuvettes (Brand, catalog number: 7590 15)
Tryptic soy broth (TSB) (Biolife, catalog number: 4021552)
Tryptic soy agar (TSA) (Biolife, catalog number: 4021502)
Phosphate buffered saline (PBS) (Oxoid, catalog number: BR0014G)
Na2HPO4 (Sigma-Aldrich, catalog number: S9763)
NaCl (Sigma-Aldrich, catalog number: S9888)
KH2PO4 (Sigma-Aldrich, catalog number: P0662)
KCl (Sigma-Aldrich, catalog number: P3911)
NaOH (Sigma-Aldrich, catalog number: S5881)
HCl (Sigma-Aldrich, catalog number: 258148)
Tris (Serva, catalog number: 37180)
Salmonella enterica subsp. enterica Infantis ŽM9 (culture collection of the Laboratory for Food Microbiology at the Department of Food Science, Biotechnical Faculty; GenBank accession number JARDYW000000000)
20× PBS (concentrated phosphate buffered saline) (see Recipes)
30 mM Tris-HCl, pH 7.5 with 0.4 M NaCl (see Recipes)
2× TSB (see Recipes)
Equipment
Refrigerated centrifuge (Thermo Fisher Scientific, Sorwall Lynx 4000)
ÄKTA pure protein purification system (Cytiva 29018224)
Microplate reader (Thermo Fisher Scientific, Varioskan Lux)
Incubator (Kambič, I-105CK)
Vortex (Domel, Vibromix 10)
Pipette set (Eppendorf, catalog number: 3120 000.836)
1–10 mL pipette (Eppendorf, catalog number: 3123 000.080)
10–100 μL multichannel pipette (Eppendorf, catalog number: 3125 000.044)
Biosafety cabinet (Iskra Pio, SMBC 122AV)
Ultra-low temperature freezer (Haier, DW-86L728ST)
Freezer (LTH, HG 5.1Z)
Spectrophotometer (Perkin Elmer, Lambda Bio+)
Microplate mixer (Eppendorf, ThermoMixer C)
Software
SkanIt RE 5.0 Plate Reader Software (Thermo Fisher Scientific, https://www.thermofisher.com/si/en/home/life-science/lab-equipment/microplate-instruments/plate-readers/software.html)
Excel (Microsoft Corporation, https://office.microsoft.com/excel)
R: A language and environment for statistical computing (R Core Team, http://www.R-project.org/)
Growthcurver: simple metrics to summarize growth curves (Sprouffske and Wagner, 2016, https://github.com/sprouffske/growthcurver, https://cran.r-project.org/web/packages/growthcurver/vignettes/Growthcurver-vignette.html)
Orange Data Mining (Demšar et al., 2013, University of Ljubljana, Faculty of Computer and Information Science, https://orange.biolab.si)
Procedure
The overall procedure is divided into several parts, with the fungal extracts first prepared (section A) and analyzed by the SIMBA method (sections C–F) and the data analyzed as described in the data analysis sections. For selected extracts showing antimicrobial and/or anti-biofilm activity, the extracts are then fractionated (section B), followed by another round of SIMBA method (sections C–F) and data analysis.
Obtaining cold water fungal extracts (CWFE) and their fractionation
Cold water extraction of fruiting bodies of wild or cultivated fungi
Collect and clean fresh fruiting bodies and store in a freezer at -20 °C (see Note 1).
Thaw frozen fruiting bodies overnight at 4–8 °C.
Place a beaker on ice.
Wrap the thawed fungal sample in the viscose cloth and place inside the potato ricer.
Press the potato ricer firmly over the beaker on ice and rotate the cloth with the sample several times to extract all the liquid that collects in the beaker below.
Measure the volume of crude extract in the beaker.
Add 1/20 of the volume of 20× PBS to obtain a concentration of 1× PBS in the extract and mix well.
Transfer to a suitable centrifuge tube that has been chilled on ice (see Note 2).
Centrifuge at 10,000× g for 10 min at 4 °C to remove insoluble material.
Carefully transfer the supernatant to suitable centrifuge tubes (on ice) (see Note 3).
Filter sterilize the supernatant, which is the cold water fungal extract (CWFE), using a 0.2 μm syringe filter.
Store CWFE in aliquots in the freezer at -20 or -80 °C until analysis with SIMBA method (Note 4).
Fractionation of CWFE with confirmed antimicrobial and/or anti-biofilm activity (see Note 5)
Thaw selected CWFE (with antimicrobial and/or anti-biofilm activity confirmed by SIMBA method) on ice.
Work with refrigeration (4–8 °C) either by using a refrigerated automated liquid chromatography system or in a cold room (see Note 6).
Equilibrate the Sephacryl S-200 column in 30 mM Tris-HCl, pH 7.5 with 0.4 M NaCl.
Load CWFE sample (at up to 2% of the total column volume) and elute in the same buffer at a flow rate of 1 mL/min.
Collect 5 mL fractions and measure the absorbance at 280 nm (Figure 1).
Figure 1. Examples of size-exclusion chromatograms of two different cold water fungal extracts (CWFEs) showing absorbance at 280 nm in fractions.
The chromatogram of CWFE 1 shows better resolution than that of CWFE 2, where all proteins elute in a single peak. Fractions are subjected to the SIMBA method, and fractions showing antimicrobial and/or anti-biofilm activity are then selected for further analysis or fractionation.
Filter sterilize fractions using 0.2 μm syringe filters.
Store fractions at 4–8 °C until analysis with SIMBA method (see Note 7).
SIMBA method—Determination of antimicrobial and anti-biofilm activity of CWFE and fractions of CWFE with confirmed antimicrobial and/or anti-biofilm activity
Revival of S. Infantis
Work aseptically.
Remove a vial containing a permanent culture of S. Infantis ŽM9 from the ultra-low temperature freezer and place it on ice.
Scrape off the frozen culture and transfer it to a TSA plate using a sterile loop.
Incubate at 37 °C for 24 h.
Preparation of the inoculum of S. Infantis
Work aseptically.
Prepare 5 mL of PBS in a 15 mL tube.
With a cotton swab, add the biomass (of S. Infantis growth on plate) and vortex for 10 s.
Add 1 mL of the bacterial suspension to a cuvette and measure the absorbance at 600 nm on the spectrophotometer. Use 1 mL of PBS as a blank.
The measured absorbance should be 0.100 ± 0.010, which corresponds to (1–2.5) × 107 CFU/mL. If the measured value is lower than the target value, add more culture. If the measured value is higher than the target value, add PBS.
Transfer 100 μL of the prepared bacterial suspension to 9.9 mL of 2× TSB.
Use the prepared inoculum within 20 min of preparation.
Setup of the microplate
Work aseptically.
Remove the CWFE/CWFE fraction sample from the freezer and place it on ice to thaw.
Vortex the thawed CWFE/CWFE fraction for 10 s.
Pipette the thawed CWFE/CWFE fraction into the wells of the microplate (see also Note 8).
Transfer 100 μL of CWFE/CWFE fraction (row A, columns 1, 2, and 3 of the microplate schematic in Figure 2) if you want to test only a 50% concentration of the extract.
Figure 2. Microplate preparation scheme before adding the prepared bacterial inoculum and 2× TSB
Transfer 200 μL of CWFE/CWFE fraction (row A, columns 4, 5, and 6 of the microplate schematic in Figure 2) if you wish to test 50% concentrations and two-fold dilutions of the extract (see Note 9).
i. Pipette 100 μL of PBS (rows B–H, columns 4, 5, and 6 of the microplate schematic in Figure 2).
ii. Transfer 100 μL of CWFE/CWFE fraction from row A to row B and mix by pipetting. Continue these transfers to row H and discard the remaining 100 μL.
Pipette 100 μL of PBS (rows B and C, columns 1, 2, and 3 of the microplate schematic in Figure 2) for positive (row B) and negative (row C) control.
Pipette 100 μL of the prepared inoculum into all wells filled with CWFE/CWFE fraction or its dilutions.
Pipette 100 μL of the prepared inoculum for positive control (row B, columns 1, 2, and 3).
Pipette 100 μL of 2× TSB for the negative control (row C, columns 1, 2, and 3).
Shake the prepared microplate on a microplate shaker at 600 rpm for 2 min.
Absorbance measurement
Set the microplate reader for kinetic measurement.
Set the temperature to 37 °C.
Set 61 kinetic intervals of 20 min, giving a total of 20 h of measurements.
Set 5 s of continuous shaking at 600 rpm before measurement.
Measure the absorbance at 600 nm.
Insert the microplate when the temperature reaches 37 °C and start the kinetic measurements.
Export the absorbance measurements to Excel.
Data analysis
All measurements should be repeated in three biological replicates, each of which is repeated in three technical replicates.
Visualization of obtained data
Open Excel file and mark the samples that were in the individual wells.
Review the initial measured absorbances and discard data where the initial measured absorbance is ≥ 0.500 (see Note 9).
For the repetitions of each extract and controls, calculate the average for each time point (see Note 10).
Plot scatterplot with smooth lines to visualize data. Put time (h) on x-axis and absorbance (600 nm) on y-axis.
Based on obtained curves of S. Infantis growth (Figure 3), visually evaluate the results (see Note 11).
Figure 3. Visual analysis of growth curve of Salmonella Infantis ŽM9. Reprinted from Sterniša et al. (2022a) with permission from Elsevier 5472521010052.
Quantitative evaluation of antimicrobial activity
Use only measurements obtained from 0 to 10 h of incubation.
Prepare data in Excel.
Name the first column as time. The name of other columns should follow the 96-well microtiter plate well marks, i.e. A1, A2, A3, etc.
Insert hours of measurements in the first column and measured absorbances at 600 nm in the corresponding well.
Use a dot to specify a decimal number and save data as tab-delimited .txt file.
Use R package Growthcurver (Sprouffske and Wagner, 2016).
Install Growthcurver package (https://github.com/sprouffske/growthcurver).
Upload data and do analysis as described (https://cran.r-project.org/web/packages/growthcurver/vignettes/Growthcurver-vignette.html).
Compare the obtained parameters of the growth curves of Salmonella exposed to the extracts to the control (growth without the extract). This includes the growth rate, the doubling time, and the carrying capacity.
Quantitative evaluation of anti-biofilm activity
Evaluate difference in endpoint measured absorbances at 600 nm.
Use only endpoint measured absorbances at 600 nm after 20 h of incubation.
Calculate average and standard deviation from all replicas for each extract.
Calculate the percent difference compared to the positive control.
Prediction of antibacterial and anti-biofilm activity from the measurements using Orange
Data formatting:
Export data to Excel.
Calculate differences between neighboring OD600 measurements (first derivative) by subtracting the measurement at time 0 from the measurement at time 1, the measurement at time 1 from the measurement at time 2, and so on until the last measurement for each sample. Please check also the provided template for data formatting.
If necessary, transpose the table in a way that the name of the compound and measurements of OD600 are listed in columns and the time points are listed in the first row, as shown in Table 1.
Table 1. Example of data formatting for prediction of antibacterial or anti-biofilm activity. Data should be formatted in a table where a list of the active compound names and corresponding measurements are presented in columns [first column: name of your sample, second column: value, measurements, and below the calculation of differences (first derivatives), third column: absorbance measurement for each compound for each cycle and calculated first derivatives].
Active compound name Value 0 1 2 3 4 5
my sample 1 technical replicate 1 measurement 0.147 0.153 0.155 0.157 0.170 0.178
my sample 1 first derivative 0.006 0.002 0.002 0.013 0.008
Use provided Orange workflow and model training dataset to predict antibacterial or anti-biofilm activity.
Download and install Orange (https://orangedatamining.com/download/#windows), a free tool for machine learning and data visualization. Data analysis in Orange is done by creating custom workflows; for the purpose of antibacterial or anti-biofilm activity prediction, we provide a workflow and model training data. You will only need to provide your data (measurements) formatted as described above. If you are interested in learning more about the usage of Orange, a wealth of short trainings is provided through Orange Data Mining You Tube channel.
Open the Orange. By clicking open, load the provided workflow named “your_compound_biofilm_activity_prediction_against_Salmonella_revised.ows.” The workflow starts with cross-validation to score learners for prediction of antibacterial or anti-biofilm activity using the provided training set composed of experimentally verified data from Sterniša et al. (2022a). This part is given at the upper part of the workflow as “learner selection.” Each widget is accessed by double-clicking on it. The widgets in the workflow are named as written in this protocol.
Double-click on the first widget on the left called “training_set” and load the provided data in the file “Salmonella_transposed_training_set_Sternisa_et_al_2022.” The type and role of data should be assigned as follows (Table 2):
Table 2. Type and role of data that should be assigned to provided data with known activity
Column name Type Role
active compound name text meta
value categorical meta
antibacterial categorical feature
antibiofilm categorical feature
time point 1(and all other time points) numeric feature
Select features and prediction target (learner selection): target feature is activity, either “antibacterial” or “antibiofilm” (target feature is the feature being predicted); meta-features are “name of the compound,” “value,” and the “activity” that is not target at the moment. You can move features by dragging them into different categories by mouse. The time points should be listed as “features.”
Select curves (learner selection): set a condition “value” “is” “first derivative.” The provided learners (random forrest, kNN, etc.) will then be tested and the widget “score and test” will gather the information on their performance. Based on the presented parameters, we will choose the best-performing learner to be used in the next part of the workflow to predict the antibacterial or anti-biofilm activity of unknown compound. With the current training set, the learner “Random Forest” performs best to predict antibacterial as well as anti-biofilm activity. By clicking on widgets “Test and Score” and “Confusion Matrix,” information on the performance of the model can be accessed.
Now, we will use the best-performing learner (Random Forest) to predict the antibacterial or anti-biofilm activity of the unknown compound (your data or provided dataset for prediction named Salmonella_test_set_for_prediction). Select features and prediction target (train for prediction): target feature is activity “antibacterial” or “antibiofilm,” this target feature has to be the same for “learner selection” and “train for prediction;” meta-features are “name of the compound,” “value,” and the “activity” that is not target at the moment. You can move features by dragging them into different categories by mouse. The measurements for each time should be listed as “features.”
Select curves (train for prediction): set a condition “value” “is” “first derivative.”
Double-click on the widget in the middle at the bottom called “unknown_active_compounds_set.” Load your formatted data as described above. The type and role of data should be assigned as follows (Table 3):
Table 3. Type and role of data that should be assigned to data with unknown activity
Column name Type Role
active compound name text meta
value categorical meta
time point 1(and all other time points) numeric feature
Select rows: set a condition “value” “is” “first derivative.” The learner “random forest” will be used to predict the antibacterial or anti-biofilm activity of the unknown compound, depending on which of the activities you chose as the target.
Check the predictions of antibacterial or anti-biofilm activity by double-clicking on the widget “Predictions.” A table will be opened and, on the left side in the column “Random Forest,” you can find the prediction results.
You may also view line plots of growth curves by clicking on the widget line plot. The line plots are associated with data tables and the results of the modeling. When you open these widgets simultaneously, you may view the growth curves selected in the table.
Notes
From approximately 50 g of fresh fruiting bodies, 5–40 mL of extract is obtained according to the protocol in section A, depending on the species.
The centrifuge tube should be compatible for use at 10,000× g, but selection depends on the volume of the extract.
Additional optional steps in extract preparation can include:
Dialysis against PBS in case small molecules should be removed (e.g., with cutoff 1 or 3.5 kDa).
Concentration by ultrafiltration or centrifugal concentrators (with concentration, small molecules are also lost).
Extracts should be filter sterilized (e.g., 0.2 μm filters) if they are to be used in bioassays with live (micro)organisms.
Aliquots of at least 300 μL should be prepared, as this is the volume needed for one Simba experiment.
Fractionation can be performed based on different properties; most often, the first type of fractionation is by size (using size exclusion or gel filtration chromatography) or charge (using ion exchange chromatography), but also hydrophobicity (using hydrophobic interaction chromatography) or affinity (using affinity chromatography) can be exploited. Size exclusion chromatography separates large protein complexes from small proteins and peptides, indicating the nature of molecules with bioactivity. The compounds elute with the peptides but may also remain in complex with larger molecules in the extracts.
When preparing extracts and their fractions, always keep your samples and the buffers you are working with chilled on ice.
If you do not use fractions the same or next day, store them in a freezer at -20 or -80 °C.
According to point E of procedure, fill the microplate as optimally as possible based on your samples. The procedure presented under point E is considered for one biological replicate.
Initial absorbances of ≥ 0.500 proved too high for analysis. In such cases, it is necessary to dilute extract under investigation in order to reduce initial absorbance.
If any of the replicates deviates significantly, exclude it from further analysis.
Define activity:
Antimicrobial effect is defined by effect on the first part of the curve—first 10 h—especially as visible prolongation of lag phase and reduced slope of log phase. Growth inhibition can also affect the second part of the curve, but this does not mean true anti-biofilm activity.
Anti-biofilm effect is defined as no effect on the first part of the curve and visible changes in the second part of the curve—from 10 to 20 h of incubation. This is visible as no secondary increase of curve due to biofilm formation compared to control.
Recipes
20× PBS
20× PBS is prepared as a 20× concentrated solution of PBS.
Dissolve the following in 800 mL of dH2O:
28.8 g of Na2HPO4 (Mw 141.96 g/mol)
160 g of NaCl (Mw 58.44 g/mol)
4.8 g of KH2PO4 (Mw 136.09 g/mol)
4 g of KCl (Mw 74.55 g/mol)
Adjust the pH to 7.4 using 10 M NaOH or HCl.
Add dH2O to 1 L.
Filter sterilize or autoclave.
Store at room temperature.
30 mM Tris-HCl, pH 7.5 with 0.4 M NaCl
Dissolve the following in 0.8 L of dH2O:
3.63 g of Tris (Mw 121.1 g/mol)
23.37 g of NaCl (Mw 58.4 g/mol)
Adjust the pH to 7.5 using HCl.
Add dH2O to 1 L.
Store at 4–8 °C.
2× TSB
2× TSB is prepared by use of double weight of TSB dehydrated culture medium in one volume.
Dissolve 60 g of TSB dehydrated culture medium in 800 mL of dH2O.
Adjust pH to 7.3 ± 0.2.
Fill to 1 L with dH2O.
Store at 4–8 °C.
Acknowledgments
This study was funded by the Slovenian Research Agency (Grant Numbers P2-0209, P4-0432, P4-0127, J3-3079, P4-0116, J4-3088, J4-4548, Z4-4551) and Innovation Fund’s project “Simultaneous detection of antimicrobial and anti-biofilm activity of active substances – development of a prototype application SIMBApp.” This protocol was derived from Sterniša et al. (2022a).
Competing interests
Patent application EP2022058548W·2022-03-31 was published WO2022207781A2·2022-10-06 (Sterniša et al., 2022b).
References
Alves, M., Ferreira, I., Dias, J., Teixeira, V., Martins, A. and Pintado, M. (2012). A Review on Antimicrobial Activity of Mushroom (Basidiomycetes) Extracts and Isolated Compounds. Planta Med. 78(16): 1707–1718.
Demšar, J., Curk, T., Erjavec, A., Gorup, C., Hočevar, T., Milutinovič, M., Možina, M., Polajnar, M., Toplak, M., Starič, A., et al. (2013). Orange: Data Mining Toolbox in Python. J. Mach. Learn. Res. 14(1): 2349−2353.
Dieltjens, L., Appermans, K., Lissens, M., Lories, B., Kim, W., Van der Eycken, E. V., Foster, K. R. and Steenackers, H. P. (2020). Inhibiting bacterial cooperation is an evolutionarily robust anti-biofilm strategy. Nat. Commun. 11(1): e1038/s41467-019-13660-x.
Eloff, J. N. (2019). Avoiding pitfalls in determining antimicrobial activity of plant extracts and publishing the results. BMC Complement. Altern. Med. 19: 106.
Erjavec, J., Kos, J., Ravnikar, M., Dreo, T. and Sabotič, J. (2012). Proteins of higher fungi – from forest to application. Trends Biotechnol. 30(5): 259–273.
Gómez Román, M. P., Badillo Mantilla, N., Andrés Carreño Flórez, S., De Mandal, S., Kumar Passari, A., Ruiz-Villáfan, B., Rodríguez-Sanoja, R. and Sánchez, S. (2020). Antimicrobial and Antioxidant Potential of Wild Edible Mushrooms. In: An Introduction to Mushroom.
González, J. F., Alberts, H., Lee, J., Doolittle, L. and Gunn, J. S. (2018). Biofilm Formation Protects Salmonella from the Antibiotic Ciprofloxacin In Vitro and In Vivo in the Mouse Model of chronic Carriage. Sci. Rep. 8(1): e1038/s41598-017-18516-2.
Klančnik, A., Piskernik, S., Jeršek, B. and Možina, S. S. (2010). Evaluation of diffusion and dilution methods to determine the antibacterial activity of plant extracts. J. Microbiol. Methods 81(2): 121–126.
Klančnik, A., Megušar, P., Sterniša, M., Jeršek, B., Bucar, F., Smole Možina, S., Kos, J. and Sabotič, J. (2017). Aqueous Extracts of Wild Mushrooms Show Antimicrobial and Antiadhesion Activities against Bacteria and Fungi. Phytother. Res. 31(12): 1971–1976.
Klančnik, A., Šimunović, K., Sterniša, M., Ramić, D., Smole Možina, S. and Bucar, F. (2021). Anti-adhesion activity of phytochemicals to prevent Campylobacter jejuni biofilm formation on abiotic surfaces. Phytochem. Rev. 20(1): 55–84.
MacKenzie, K. D., Palmer, M. B., Köster, W. L. and White, A. P. (2017). Examining the Link between Biofilm Formation and the Ability of Pathogenic Salmonella Strains to Colonize Multiple Host Species. Front. Vet. Sci. 4: e00138.
Pate, M., Mičunovič, J., Golob, M., Vestby, L. K. and Ocepek, M. (2019). Salmonella Infantis in Broiler Flocks in Slovenia: The Prevalence of Multidrug Resistant Strains with High Genetic Homogeneity and Low Biofilm-Forming Ability. Biomed Res. Int. 2019: 1–13.
Savjani, K. T., Gajjar, A. K. and Savjani, J. K. (2012). Drug Solubility: Importance and Enhancement Techniques. ISRN Pharmaceutics 2012: 1–10.
Sprouffske, K. and Wagner, A. (2016). Growthcurver: an R package for obtaining interpretable metrics from microbial growth curves. BMC Bioinf. 17(1): e1186/s12859-016-1016-7.
Steenackers, H., Hermans, K., Vanderleyden, J. and De Keersmaecker, S. C. (2012). Salmonella biofilms: An overview on occurrence, structure, regulation and eradication. Food Res. Int. 45(2): 502–531.
Sterniša, M., Sabotič, J. and Klančnik, A. (2022a). A novel approach using growth curve analysis to distinguish between antimicrobial and anti-biofilm activities against Salmonella. Int. J. Food Microbiol. 364: 109520.
Sterniša, M., Sabotič, J. and Klančnik, A. (2022b). International patent application WO2022207781A2. Geneva, Switzerland: World Intellectual Property Organization.
Tassinari, E., Duffy, G., Bawn, M., Burgess, C. M., McCabe, E. M., Lawlor, P. G., Gardiner, G. and Kingsley, R. A. (2019). Microevolution of antimicrobial resistance and biofilm formation of Salmonella Typhimurium during persistence on pig farms. Sci. Rep. 9(1): e1038/s41598-019-45216-w.
Vestby, L. K., Møretrø, T., Langsrud, S., Heir, E. and Nesse, L. L. (2009). Biofilm forming abilities of Salmonellaare correlated with persistence in fish meal- and feed factories. BMC Vet. Res. 5(1): e1186/1746-6148-5-20.
Supplementary information
The following supporting information can be downloaded here:
template_for_data_formatting.xlsx
Figure workflow Orange
Salmonella_transposed_training_set_Sternisa_et_al_2022.xlsx
Salmonella_test_set_for_prediction.xlsx
your_compound_biofilm_activity_prediction_against_Salmonella_revised.ows
Article Information
Copyright
© 2023 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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Peer-reviewed
A qPCR Method to Distinguish between Expression of Transgenic and Endogenous Copies of Genes
WB William Bezodis
HP Helen Prescott
DV Daniela Vlad
HD Hugh Dickinson
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4784 Views: 735
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Original Research Article:
The authors used this protocol in bioRxiv Sep 2022
Abstract
Study of gene function in eukaryotes frequently requires data on the impact of the gene when it is expressed as a transgene, such as in ectopic or overexpression studies. Currently, the use of transgenic constructs designed to achieve these aims is often hampered by the difficulty in distinguishing between the expression levels of the endogenous gene and its transgene equivalent, which may involve either laborious microdissection to isolate specific cell types or harvesting tissue at narrow timepoints. To address this challenge, we have exploited a feature of the Golden Gate cloning method to develop a simple, restriction digest–based protocol to differentiate between expression levels of transgenic and endogenous gene copies. This method is straightforward to implement when the endogenous gene contains a Bpi1 restriction site but, importantly, can be adapted for most genes and most other cloning strategies.
Key features
• This protocol was developed to determine the expression level of an ectopically expressed transcription factor with broad native expression in all surrounding tissues.
• The method described is most directly compatible with Golden Gate cloning but is, in principle, compatible with any cloning method.
• The protocol has been developed and validated in the model plant Arabidopsis thaliana but is applicable to most eukaryotes.
Graphical overview
Keywords: qPCR RT-qPCR Golden Gate Arabidopsis Transgene expression
Background
Quantifying specific spatial or temporal expression of a transgenically expressed endogenous gene against a background of high or overlapping expression of the native endogenous sequence remains a long-standing problem in eukaryotic molecular research. To overcome this, we have developed a protocol based on the introduction of a synonymous mutation into the transgenic copy of the endogenous gene to remove a restriction enzyme cleavage site. This is frequently a feature of the assembly of Golden Gate Cloning constructs (the so-called domestication process, involving excision of key Type IIS restriction nuclease sites) (Engler et al., 2008; Weber et al., 2011). This restriction site difference can then be exploited to differentiate between endogenous and transgenic expression by digesting complementary DNA (cDNA) from transgenic individuals with the relevant restriction enzyme, which will cleave endogenous cDNA but will leave the transgene cDNA uncut. Misexpression of an endogenous gene in a tissue where it is not normally expressed using a cell type–specific promoter is a commonly used experimental technique (Prelich, 2012). However, if that cell type is temporally or spatially difficult to isolate, it can be challenging to determine accurately the levels of overexpression that have been achieved without technically complex and laborious dissection. The protocol removes the need to isolate the specific cell types to confirm expression but instead allows this to be done on RNA extracted from bulk tissues.
Essentially, the protocol set out below involves making a domesticated version of the transgene missing a restriction site present in the endogenous sequence and transforming it into the organism under study. Following RNA extraction and standard reverse transcription, an additional digestion step is introduced whereby the cDNA is restricted using the enzyme targeting the site now only present in cDNA from the endogenous gene. Subsequent qPCR, using primers amplifying a region of the gene containing the restriction site, will thus only amplify cDNA specific to the transgene, leaving the endogenous cDNA unaffected—since it can no longer serve as a template for amplification. qPCR on this digested cDNA using primers amplifying a region of a housekeeping gene that does not contain the restriction site allows the calculation of relative expression levels using the 2-ΔΔC(T) method (Livak and Schmittgen, 2001). Although this protocol is simplest to implement in cases where Golden Gate cloning is used and an endogenous Bpi1 restriction enzyme cut site has been removed, with planning during cloning this same principle can also be adapted to any other cloning strategy by the introduction of a synonymous mutation that removes a restriction enzyme cleavage site from the transgene. Overall, this technique allows very rapid and accurate estimation of the transgene expression level, as negligible endogenous cDNA survives cleavage and thus expression levels detected by qPCR reflect only the transgene. cDNA from wildtype organisms can, of course, be used to determine approximate control levels of endogenous gene expression and, in the case of inducible transgenes, a more accurate estimate of endogenous gene expression can be obtained by sampling cDNA from uninduced tissue.
This method was developed by Bezodis et al. (2022) to detect expression from inducible constructs that ectopically express high levels of homeodomain transcription factors in a spatially and temporally restricted region of the germline lineage of the model plant Arabidopsis thaliana. While the transcription factors of interest are normally not expressed in the germline, they are expressed significantly throughout the surrounding somatic tissues. These endogenous transcripts thus constitute a major component of the RNA collected from whole tissues but, by digesting cDNA from the endogenous gene copy carrying the functional restriction site, this method avoids the laborious task of dissecting out cells in which only the transgene is expressed. The combination of this qPCR strategy and the use of imaging markers has permitted both the location and extent of transgene expression to be accurately determined using promoters driving expression in specific germline cell types over a short period. We have found the use of an inducible system to be particularly helpful, as it provides a control expression level to confirm that the digest has worked effectively at removing expression of the endogenous gene and matching construct-driven expression to observed phenotypes (Bezodis et al., 2022). Induction uses a two-component LhGR/pOp6 expression system (Craft et al., 2005) and a protocol for this system has been published (Samalova et al., 2019), including for use in reproductive tissues (Schubert et al., 2022). We note, however, that the protocol does not require an inducible system, nor is it limited to Arabidopsis or, in fact, to plants.
Materials and reagents
Biological materials
Gene synthesis; GeneArt Gene Synthesis (Thermo Fisher Scientific, UK)
Laboratory supplies
RNeasy Plant Mini kit (Qiagen, catalog number: 74904)
RNase-free DNase set (Qiagen, catalog number: 79254)
Bpi1 (Bbs1) restriction enzyme (Thermo Fisher Scientific, catalog number: ER1011)
qPCRBIO cDNA synthesis kit (PCR Biosystems Ltd, catalog number: PB30.11-10)
SuperScriptTM IV First-Strand Synthesis System (Thermo Fisher Scientific, catalog number: 18091050)
qPCR SyGreen Mix (PCR Biosystems Ltd, catalog number: PB20.12-05)
qPCR plates (Applied Biosystems, Thermo Fisher Scientific, catalog number: 4346906)
Microamp Fast Optical 96-well reaction plates with barcode (Thermo Fisher Scientific, catalog number 4346906)
Adhesive PCR plate seals (Thermo Scientific, catalog number: AB-0558)
Equipment
MiniAmp thermal cycler (Thermo Fisher Scientific)
StepOnePlus real-time PCR system (Applied Biosystems, Thermo Fisher Scientific)
NanoDrop (Thermo Scientific, Labtech, ND1000, Thermo Fisher Scientific)
Software
Construct design and sequence analysis was carried out using SnapGene software (www.snapgene.com); a plasmid editor, ApE, is a suitable freely available alternative (Davis and Jorgensen, 2022)
qPCR data analysis was performed using Thermo Fisher Data Connect Design & Analysis Software
Procedure
Transgene design
Key to this method is the removal of a Bpi1 restriction site from the coding region of the gene that is to be expressed as a transgene. This is frequently required for Golden Gate cloning (Engler et al., 2008; Weber et al., 2011) and has been described in previous protocols (Engler and Marillonnet, 2014). For other cloning methods, any suitable restriction site(s) can be removed.
RNA extraction, cDNA synthesis, qPCR
This step is only relevant if using the inducible system as in Bezodis et al. (2022). This protocol may be used for any system in which a transgenic copy of an endogenous gene is expressed.
Our validation experiments on Arabidopsis thaliana (described below) utilised an inducible expression system active in inflorescences (Bezodis et al., 2022; Schubert et al., 2022). At least four biological replicates are recommended for achieving highly reproducible results.
Select Arabidopsis plants with at least four inflorescence apices; mark two inflorescences for pre-induction and two for post-induction expression, with small pieces of tape of different colours.
Collect the two pre-induction inflorescence apices, harvesting all buds that have not clearly been fertilised. Snap freeze in liquid nitrogen and store at -80 °C until all samples are ready for RNA extraction.
Apply induction as described (Schubert et al., 2022) to the other two marked inflorescences, harvested 18 h after induction.
Extract RNA using the method appropriate for the system being used. For the work in which this method was developed, RNeasy Plant Mini kit with on-column DNA digest was used, following manufacturer’s recommended methods (Note 1).
Synthesize cDNA from the resultant RNA. The qPCRBIO cDNA synthesis kit and SuperScript IV Reverse Transcriptase have been tested; the resultant cDNA, following manufacturer’s recommended protocol, proved suitable for use with this method.
cDNA digestion:
Determine approximate cDNA concentration by NanoDrop (Note 2).
Dilute cDNA to 250 ng/μL in molecular biology grade water (Note 3).
Digest cDNA in 96-well PCR plates in 20 μL reactions as below (Table 1):
Table 1. Components for cDNA Restriction Digest
Component Volume
250 ng/μL diluted cDNA 14 μL
Bpi1 4 μL
10× digestion buffer 2 μL
Mix by pipetting and spin down plates in a plate centrifuge at ~1,200× g for 30 s.
Perform digestion in a thermocycler as below, with lid temperature 103 °C (Table 2).
Table 2. Thermocycler run for cDNA Restriction Digest
Thermocycler step Step 1 Step 2 Step 3
Duration 8 h 20 min hold
Temperature 37 °C 65 °C 4 °C
Primer design and qPCR
Primer design
Primers are needed to amplify an endogenous control housekeeping gene and the gene of interest. The amplification product should be in the region of 70–200 bp with an annealing temperature of > 60 °C.
For the housekeeping gene, such as ubiquitin or tubulin, the region amplified should not contain a Bpi1 site (or other restriction enzymes being used), whilst for the gene of interest the region amplified must contain a region that contains a Bpi1 (or other restriction enzyme) site in the endogenous copy that has been removed as described in section A. Selection of housekeeping genes as controls for qPCR has been discussed previously (Czechowski et al., 2005) and depends on the aims of the specific experiment and tissue being used (Dekkers et al., 2012). Protocol validation by Bezodis et al. (2022) used AtUBQ10 (AT4G05320), which is more stable than some other commonly used reference genes (Czechowski et al., 2005).
Once a suitable region for amplification has been chosen, optimal primers can be designed using primer3 (primer3.ut.ee) (Untergasser et al., 2012).
qPCR reaction
Perform qPCR in Microamp Fast Optical 96-well reaction plates with Adhesive PCR plate seals in either 10 or 20 μL reactions; for the latter, volumes below should be doubled (Table 3).
Table 3. Components for qPCR
Component Volume
Digested cDNA 1 μL
2× SyGreen qPCR Master Mix 5 μL
10 μM forward primer and 10 μM reverse primer stock solution 1 μL
Molecular biology grade water 3 μL
Perform qPCR on a StepOnePlus real-time PCR system programmed as shown below (Table 4). A melt curve is used to check primer specificity. Lid temperature set to 103 °C.
Table 4. Real-time PCR system run for qPCR
Thermocycler step Step 1 Step 2
Duration 3 min 15 s 30 s
Temperature 95 °C 95 °C 60 °C
Number of cycles 1 40
Data analysis
qPCR data are analysed either by manual calculation using Excel or more easily with Thermo Fisher Data Connect Design & Analysis Software, which allows quantification of expression using the 2-ΔΔC(T) method (Livak and Schmittgen, 2001). The level of expression of the transgene from digested cDNA is normalised to the level of housekeeping gene used. As described, it is important that the region of the housekeeping gene amplified does not include a restriction enzyme target and, if properly designed as described, the analysis will be no different to any other qPCR experiment. It is important to use the number of biological replicates as the sample size for statistical analyses, as the technical replicates should be averaged and used only to ensure values from the same sample are similar. Various methods of analysis of qPCR data are available, which have been reviewed by Pabinger et al. (2014). Application of this protocol does not change the data analysis.
Notes
Numerous kits from different manufacturers are available and method should follow manufacturer’s recommended protocol.
Sufficient cDNA is needed for at least six qPCR reactions to allow the amplification of a control housekeeping gene for normalisation and the transgenically expressed endogenous gene of interest, at least in triplicate for technical replicates. This is in addition to the biological replicates described in point B1.
This concentration must be lower than the concentration of the most diluted sample; otherwise, RNA extraction and cDNA synthesis should be repeated.
Validation of protocol
Data validating this protocol can be found in our preprint Bezodis et al. (2022) deposited on bioRxiv at doi.org/10.1101/2022.09.15.508103; relevant data are shown in Figure 1D–1G.
Acknowledgments
We thank Vinay Shukla for help with operation of the qPCR machine and data analysis methods and Jasmine Staples for help designing the graphical overview. This work was carried out and funded as part of an Mbiol masters project in the Department of Plant Sciences (now Department of Biology), University of Oxford.
Competing interests
We have no competing interests.
References
Bezodis, W., Prescott, H. and Dickinson, H. (2022). Misexpression of the Homologues of Bryophyte Gametophyte-to-Sporophyte Control Genes in Arabidopsis Results in Germline Reprogramming and Phenotypes that Mirror Apomictic Development. bioRxiv: e508103.
Craft, J., Samalova, M., Baroux, C., Townley, H., Martinez, A., Jepson, I., Tsiantis, M. and Moore, I. (2005). New pOp/LhG4 vectors for stringent glucocorticoid-dependent transgene expression in Arabidopsis. Plant J. 41(6): 899–918.
Czechowski, T., Stitt, M., Altmann, T., Udvardi, M. K. and Scheible, W. R. (2005). Genome-Wide Identification and Testing of Superior Reference Genes for Transcript Normalization in Arabidopsis. Plant Physiol. 139(1): 5–17.
Davis, M. W. and Jorgensen, E. M. (2022). ApE, A Plasmid Editor: A Freely Available DNA Manipulation and Visualization Program. Front. Bioinf. 2: e818619.
Dekkers, B. J. W., Willems, L., Bassel, G. W., van Bolderen-Veldkamp, R. P., Ligterink, W., Hilhorst, H. W. M. and Bentsink, L. (2012). Identification of Reference Genes for RT–qPCR Expression Analysis in Arabidopsis and Tomato Seeds. Plant Cell Physiol. 53(1): 28–37.
Engler, C., Kandzia, R. and Marillonnet, S. (2008). A One Pot, One Step, Precision Cloning Method with High Throughput Capability. PLoS One 3(11): e3647.
Engler, C. and Marillonnet, S. (2014). Golden Gate Cloning. In: Valla, S. and Lale, R. (Eds.). DNA Cloning and Assembly Methods (pp. 119–131). Methods in Molecular Biology. Humana Press, Totowa, NJ.
Livak, K. J. and Schmittgen, T. D. (2001). Analysis of Relative Gene Expression Data Using Real-Time Quantitative PCR and the 2−ΔΔCT Method. Methods 25(4): 402–408.
Pabinger, S., Rödiger, S., Kriegner, A., Vierlinger, K. and Weinhäusel, A. (2014). A survey of tools for the analysis of quantitative PCR (qPCR) data. Biomol. Detect. Quantif. 1(1): 23–33.
Prelich, G. (2012). Gene Overexpression: Uses, Mechanisms, and Interpretation. Genetics 190(3): 841–854.
Samalova, M., Kirchhelle, C. and Moore, I. (2019). Universal Methods for Transgene Induction Using the Dexamethasone‐Inducible Transcription Activation System pOp6/LhGR in Arabidopsis and Other Plant Species. Curr. Protoc. Plant Biol. 4(2): e20089.
Schubert, J., Li, Y., Mendes, M. A., Fei, D., Dickinson, H., Moore, I. and Baroux, C. (2022). A procedure for Dex-induced gene transactivation in Arabidopsis ovules. Plant Methods 18(1): e1186/s13007–022–00879–x.
Untergasser, A., Cutcutache, I., Koressaar, T., Ye, J., Faircloth, B. C., Remm, M. and Rozen, S. G. (2012). Primer3—new capabilities and interfaces. Nucleic Acids Res. 40(15): e115–e115.
Weber, E., Engler, C., Gruetzner, R., Werner, S. and Marillonnet, S. (2011). A Modular Cloning System for Standardized Assembly of Multigene Constructs. PLoS One 6(2): e16765.
Article Information
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Molecular Biology > DNA > Gene expression
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Likelihood-based Modeling of Brassiceae Ancient Whole-genome Triplication with POInT
YH Yue Hao
GC Gavin C. Conant
Published: Aug 20, 2023
DOI: 10.21769/BioProtoc.4786 Views: 79
Reviewed by: Avinash Chandra Pandey Anonymous reviewer(s)
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Abstract
Ancient whole-genome duplication and triplication events have a profound impact on present-day plant genomes. However, sub genome assignments in allopolyploids can be technically challenging. The present study used a likelihood-based tool, POInT (the Polyploid Orthology Inference Tool) to model the resolution of the ancient whole-genome triplication events in Brassiceae. This protocol will showcase the application of POInT for polyploid comparative genomics studies and its potential use for future studies.
Keywords: Allopolyploidy Whole-genome triplication Biased fractionation Synteny Orthology Brassiceae
Background
Ancient polyploidy events are prevalent throughout the evolutionary history of flowering plants (One Thousand Plant Transcriptomes Initiative, 2019) and have contributed to the diversity of complexity of present-day species (van de Peer et al., 2009). When the sub genomes were derived from genetically distant parents, such events are termed as allopolyploidy. Post-polyploidy gene loss in allopolyploids are often unbalanced across different sub genomes (Cheng et al., 2012). The biased fractionation reshaped genomic landscape, and the gene retention/loss pattern has important implications in crop development (Qi et al., 2021). However, sub genome assignment in allopolyploids is still challenging (Edger et al., 2018).
POInT (the Polyploid Orthology Inference Tool) is a likelihood-based tool for modeling ancient polyploidy events (Conant and Wolfe, 2008; Emery et al., 2018; Hao and Conant, 2022; Conant, 2023). POInT supports user-defined ploidy levels, namely whole-genome duplication (Conant and Wolfe, 2008; Emery et al., 2018; Conant, 2020), triplication (Schoonmaker et al., 2020; Hao et al., 2021), or quadruplication (Hao et al., 2022). Using POInT, we have modeled the whole-genome triplication (WGT) events shared by the members of the tribe Brassiceae (Hao et al., 2021). The synteny-based and phylogenetically aware analysis of gene loss across different sub genomes provides statistical support for the two-step hypothesis of hexaploidy and biased fractionation across the LF, MF1, and MF2 sub genomes after the ancient WGT (Cheng et al., 2012; Tang et al., 2012). Here, we present a step-by-step guide to recreate the WGT model comparisons in Hao et al. (2021).
Software and dataset
Software
POInT (Conant and Wolfe, 2008; Emery et al., 2018; Hao and Conant, 2022; version 1.55; http://conantlab.org/POInT/POInT.html)
Input data
The input files for this case study are available at https://doi.org/10.6084/m9.figshare.12277832 (Hao et al., 2021).
Input files include:
Gene orders from four individual polyploid genomes:
i. Brassica_oleracea_POInT_geneorders.txt
ii. Brassica_rapa_POInT_geneorders.txt
iii. Crambe_hispanica_v3_POInT_geneorders.txt
iv. Sinapis_alba_POInT_geneorders.txt
Inferred ancestral pillar order: FourSpp_M2Opt3.txt
Optimal phylogenetic topology: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
WGT models for use with POInT:
- WGT Null: WGT_Null_model.txt
- WGT 1Dom: WGT_2rate_G1Dom_model.txt
- WGT 1DomG3: WGT_3rate_G1Dom_model.txt
- Root-spec. WGT 1DomG3: WGT_3rate_G1Dom_brnspec_model.txt
- Root model: WGT_RootModel.txt
Procedure
Software download and installation
The latest version of the POInT software is available on GitHub (https://github.com/gconant0/POInT) or on the lab website (http://conantlab.org/POInT/POInT.html). For a detailed installation guide, please go to the software installation page (http://conantlab.org/POInT/INSTALL). To download and compile the software:
wget http://conantlab.org/POInT/POInT.tar
tar xvf POInT.tar
cd POInT
# To compile the OpenMP parallel version
./configure.pl -omp
Make
Create a symbolic link in the home directory bin:
# User should check ~/.bash_profile to see how PATH was defined.
# Add “export PATH=$PATH:$HOME/bin” if not already.
cd ~/bin
ln -s /where_POInT_was_downloaded/POInT/POInT
Test to see if the software was successfully compiled:
cd
POInT
You should see the following:
Using 16 threads for this run
Usage: POInT -g:<genome file> -g:<genome file> -o:<ortholog file> -m:<Model file> (-r:<Root model file>) (-t:treefile) (-p:<posteriortrackprobs file>) (-c:<conditional probabilities file> (-no_opt) (-s:<start>:<end>) (-zerolengthfixed) (-x:#TreestoSave)
Basic usage
A basic POInT run requires a minimum of three types of files and a specification of the polyploidy type:
-d: 2 for WGD, 3 for a WGT, and 4 for an octaploidy.
-g: genome files, listing the order of double/triple/quadruple conserved synteny genes in the extant genomes.
-o: order file, an inferred ancestral order of the conserved synteny blocks.
-m: model file, a phylogenetic model of post-polyploidy gene loss.
Additionally, it is very useful to specify an assumed phylogenetic topology using -t. If no tree file is given, POInT will try to search all possible topologies. However, for larger datasets with five or six genomes, this step will be intractable. Here, the optimal topology for the four species in the case study is provided.
Testing different WGT models
The analyses showcased here were performed on the Linux system. Twenty-four cores were requested when submitting the job to the high-performance computing cluster. It takes approximately 17 min to complete a single iteration of likelihood estimation using 96 threads on a 24-core node, and several hours or longer to reach the optimum.
export OMP_NUM_THREADS=96
WGT Null model scenario: Null model with no biased fractionation.
POInT -d:3 -g:Brassica_rapa_POInT_geneorders.txt /
-g:Brassica_oleracea_POInT_geneorders.txt /
-g:Sinapis_alba_POInT_geneorders.txt /
-g:Crambe_hispanica_v3_POInT_geneorders.txt -o:FourSpp_M2Opt3.txt /
-m:WGT_Null_model.txt /
-t: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
WGT 1Dom model scenario: MF1 and MF2 sub genomes are more fractionated than the LF sub genome, but the fractionation rates for MF1 and MF2 are the same.
POInT -d:3 -g:Brassica_rapa_POInT_geneorders.txt /
-g:Brassica_oleracea_POInT_geneorders.txt /
-g:Sinapis_alba_POInT_geneorders.txt /
-g:Crambe_hispanica_v3_POInT_geneorders.txt -o:FourSpp_M2Opt3.txt /
-m:WGT_2rate_G1Dom_model.txt /
-t: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
WGT 1DomG3 model scenario: MF2 is more fractionated than MF1, and MF1 is more fractionated than LF.
POInT -d:3 -g:Brassica_rapa_POInT_geneorders.txt /
-g:Brassica_oleracea_POInT_geneorders.txt /
-g:Sinapis_alba_POInT_geneorders.txt /
-g:Crambe_hispanica_v3_POInT_geneorders.txt -o:FourSpp_M2Opt3.txt /
-m:WGT_3rate_G1Dom_model.txt /
-t: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
Root-spec. WGT 1DomG3 is similar to model 3, but with two sets of parameters: one for the root branch and the other for the remainder of the branches in the phylogenetic tree, modeling the scenario of shifted fractionation rates from root branch to later branches.
POInT -d:3 -g:Brassica_rapa_POInT_geneorders.txt /
-g:Brassica_oleracea_POInT_geneorders.txt /
-g:Sinapis_alba_POInT_geneorders.txt /
-g:Crambe_hispanica_v3_POInT_geneorders.txt -o:FourSpp_M2Opt3.txt /
-m:WGT_3rate_G1Dom_brnspec_model.txt /
-t: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
WGT 1DomG3 + Root is modeling the two-step hexaploidy scenario (Cheng et al., 2012; Tang et al., 2012), in which the MF1 and MF2 merged first following an initial round of gene loss, and the LF sub genome joined later, with more subsequent gene loss.
POInT -d:3 -g:Brassica_rapa_POInT_geneorders.txt /
-g:Brassica_oleracea_POInT_geneorders.txt /
-g:Sinapis_alba_POInT_geneorders.txt /
-g:Crambe_hispanica_v3_POInT_geneorders.txt -o:FourSpp_M2Opt3.txt /
-m: WGT_3rate_G1Dom_model.txt -r:WGT_RootModel.txt /
-t: BrBoSaCh_WGT_3rate_G1Dom_M2Opt3_Top3.tre
Data analysis
Result interpretation
The likelihoods for different WGT models were compared using likelihood ratio tests (LRTs), shown in Figure 2 of Hao et al. (2021). The significant likelihood increases from model 1 WGT Null to model 2 WGT 1Dom (P < 10-10), and from model 2 WGT 1Dom to model 3 WGT 1DomG3 (P < 10-10), showed strong evidence of biased fractionation in the three sub genomes. We also found strong support (P < 10-10) for the two-step hexaploidy model WGT 1DomG3 + Root, suggesting that the first two sub genomes, MF1 and MF2, merged first, and the third sub genome LF joined after a certain amount of time. All of the completed ancient polyploidy analyses can be visualized using an online tool called the POInTbrowse (http://wgd.statgen.ncsu.edu/; Conant and Wolfe, 2008; Siddiqui and Conant, 2023). Users will be able to browse the gene retention/loss patterns in different sub genomes with simple clicks.
Acknowledgments
G.C.C. is supported by U.S. National Science Foundation (NSF) Division of Integrative Organismal Systems (IOS) grant NSF-IOS-1339156. This protocol was derived from Hao et al. (2021) and the POInT website (http://conantlab.org/POInT/POInT.html).
Competing interests
The authors have no conflicts of interest to declare that are relevant to the content of this article.
References
Cheng, F., Wu, J., Fang, L., Sun, S., Liu, B., Lin, K., Bonnema, G. and Wang, X. (2012). Biased gene fractionation and dominant gene expression among the subgenomes of Brassica rapa. PLoS One 7(5): e36442.
Conant, G.C. (2023). POInT: Modeling Polyploidy in the Era of Ubiquitous Genomics. In Y. Van de Peer (Ed.). Polyploidy (pp. 77–90). Methods in Molecular Biology. Humana, New York.
Conant, G. C. (2020). The lasting after-effects of an ancient polyploidy on the genomes of teleosts. PLoS One 15(4): e0231356.
Conant, G. C. and Wolfe, K. H. (2008). Probabilistic cross-species inference of orthologous genomic regions created by whole-genome duplication in yeast. Genetics 179(3): 1681–1692.
Edger, P. P., McKain, M. R., Bird, K. A. and VanBuren, R. (2018). Subgenome assignment in allopolyploids: challenges and future directions. Curr. Opin. Plant Biol. 42: 76–80.
Emery, M., Willis, M. M. S., Hao, Y., Barry, K., Oakgrove, K., Peng, Y., Schmutz, J., Lyons, E., Pires, J. C., Edger, P. P., et al. (2018). Preferential retention of genes from one parental genome after polyploidy illustrates the nature and scope of the genomic conflicts induced by hybridization. PLoS Genet. 14(3): e1007267.
Hao, Y. and Conant, G. C. (2022). POInT: A Tool for Modeling Ancient Polyploidies Using Multiple Polyploid Genomes. In Pereira-Santana, A., Gamboa-Tuz, S. D. and Rodríguez-Zapata, L. C. (Eds.). Plant Comparative Genomics (pp. 81–91). Methods in Molecular Biology. Humana, New York.
Hao, Y., Fleming, J., Petterson, J., Lyons, E., Edger, P. P., Pires, J. C., Thorne, J. L. and Conant, G. C. (2022). Convergent evolution of polyploid genomes from across the eukaryotic tree of life. G3 Genes|Genomes|Genetics 12(6): e1093/g3journal/jkac094.
Hao, Y., Mabry, M. E., Edger, P. P., Freeling, M., Zheng, C., Jin, L., VanBuren, R., Colle, M., An, H., Abrahams, R. S., et al. (2021). The contributions from the progenitor genomes of the mesopolyploid Brassiceae are evolutionarily distinct but functionally compatible. Genome Res. 31(5): 799–810.
One Thousand Plant Transcriptomes, I. (2019). One thousand plant transcriptomes and the phylogenomics of green plants. Nature 574(7780): 679–685.
Qi, X., An, H., Hall, T. E., Di, C., Blischak, P. D., McKibben, M. T. W., Hao, Y., Conant, G. C., Pires, J. C., Barker, M. S., et al. (2021). Genes derived from ancient polyploidy have higher genetic diversity and are associated with domestication inBrassica rapa. New Phytol. 230(1): 372–386.
Schoonmaker, A., Hao, Y., Bird, D. M. and Conant, G. C. (2020). A Single, Shared Triploidy in Three Species of Parasitic Nematodes. G3 Genes|Genomes|Genetics 10(1): 225–233.
Siddiqui, M. and Conant, G. C. (2023). POInTbrowse: orthology prediction and synteny exploration for paleopolyploid genomes. BMC Bioinform 24: 174.
Tang, H., Woodhouse, M. R., Cheng, F., Schnable, J. C., Pedersen, B. S., Conant, G., Wang, X., Freeling, M. and Pires, J. C. (2012). Altered patterns of fractionation and exon deletions in Brassica rapa support a two-step model of paleohexaploidy. Genetics 190(4): 1563–1574.
Van de Peer, Y., Maere, S. and Meyer, A. (2009). The evolutionary significance of ancient genome duplications. Nat Rev Genet 10(10): 725–732.
Supplementary information
Data and code availability: all data and code have been deposited into GitHub: https://github.com/Bio-protocol/POInT-WGT.
Article Information
Copyright
© 2023 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
Computational Biology and Bioinformatics
Plant Science > Plant molecular biology > Genetic analysis
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Likelihood Estimation of Arabidopsis Population Parameters with MAPGD
YH Yue Hao
MA Matthew S. Ackerman
ML Michael Lynch
Published: Aug 20, 2023
DOI: 10.21769/BioProtoc.4787 Views: 110
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Abstract
MAPGD (https://github.com/LynchLab/MAPGD) is a likelihood-based software for population genomics data analysis. Here, we demonstrate a simple tutorial for using MAPGD to analyze Arabidopsis population data. We estimated the allele and genotype frequencies, the average heterozygosity, relatedness, and other population parameters. This pipeline could be applied to further plant population genomic studies.
Keywords: MAPGD Population genomics Arabidopsis SNP Allele frequency Heterozygosity
Background
MAPGD (Ackerman, 2016; Ackerman et al., 2022) is a series of related programs that estimate allele frequency, heterozygosity, Hardy-Weinberg disequilibrium, linkage disequilibrium, and identity-by-descent coefficients from population genomic data using a statistically rigorous maximum likelihood approach. The MAPGD software was primarily designed for analysis of planktonic crustacean Daphnia populations (Lynch et al., 2017; Maruki et al., 2019; Ye et al., 2019). It is most useful for the analysis of low coverage population genomic data or pooled data, where many individuals are used to prepare a single sample (Lynch et al., 2014). Since its release, MAPGD has been widely used in population genomic studies of various organisms, including the nematode Caenorhabditis elegans (Adams et al., 2022), stickleback fish (Jeffries et al., 2022), and human gut microbiota (Shoemaker, 2022). A benchmark study showed that MAPGD could outperform other single nucleotide polymorphism (SNP) callers and has great potential in different applications (Guirao-Rico and González, 2021). Here, we demonstrate the usage of MAPGD in the analysis of a publicly available Arabidopsis population dataset (The 1001 Genomes Consortium, 2016). We show that MAPGD could be used for plant population genomic studies.
Software and datasets
Software
SAMtools (Li et al., 2009; Li, 2011; Danecek et al., 2021; version 1.9; http://www.htslib.org/)
MAPGD (Lynch et al., 2014; Maruki and Lynch, 2014 and 2015; Ackerman et al., 2017 and 2022; version 0.4.40; https://github.com/LynchLab/MAPGD). The complete user manual is available at https://github.com/LynchLab/MAPGD/tree/master/docs/man. Users may also find the software performance testing results at https://github.com/LynchLab/genomics_simulation
R (R Core Team, 2022; 4.1.3, https://www.r-project.org/) for visualization
Input data
The sample bam files were downloaded from the Arabidopsis 1001 Genomes Project (The 1001 Genomes Consortium, 2016). All five bam files used were from the “JGIHeazlewood2011” subset. The subset project page is: https://1001genomes.org/projects/JGIHeazlewood2011/index.html.
Procedure
The analyses showcased here were performed on the Linux system.
Prerequisites
MAPGD depends on the standard GNU Scientific Library (GSL); if GSL is already installed on the system, you should load the GSL module prior to the MAPGD installation. Commonly, pre-installed available modules on high-performance computing clusters can be checked using “module avail.” For this demonstration, we are using GSL version 2.6.
module load GSL/2.6-GCC-9.3.0
SAMtools will be used for bam file processing:
module load samtools/1.9
Installation
Using MAPGD version 0.4.40 for this case study:
wget https://github.com/LynchLab/MAPGD/archive/master.zip
unzip master.zip
cd MAPGD-master/
# To install software in your home directory
./configure --prefix=/home/directory
The configuration step will create “Makefiles” that are later used for software compilation. Once all the configuration is done, the package can then be compiled using the standard commands:
make
make install
Check to see if the software has been successfully installed in the home directory:
cd
mapgd –help
Check usage of a certain command; in this case, the allele command:
mapgd allele -h
Copy useful scripts to home directory bin:
cd /where/MAPGD/was/downloaded/MAPGD-master/extras
cp sub_sample.py /home/directory/bin
Test dataset download
The example bam files were downloaded from the Arabidopsis 1001 Genomes Project subset JGIHeazlewood2011 (The 1001 Genomes Consortium, 2016). Five Arabidopsis thaliana accessions were used, namely Ri-0, Jea, Sakata, Oy-0, and Alc-0. The reads were mapped to the TAIR10 reference genome (Berardini et al., 2015). The previously mapped results were used for subsequent analyses.
Data download command example:
wget https://1001genomes.org/data/JGI/JGIHeazlewood2011/releases/ \
current/TAIR10/strains/Alc-0/Alc-0.bam --no-check-certificate
Population analysis using MAPGD
Sort and filter bam files using SAMtools (http://www.htslib.org/doc/samtools.html). Unmapped reads, PCR duplicates, reads failed QC, and supplementary alignments that are not primary alignments were removed (SAM flag 3844). Create a .mpileup file using SAMtools.
samtools sort -o <file>.sort.bam <file>.bam
samtools view -q 20 -f 3 -F 3844 -b <file>.sort.bam \
> <file>.filtered.sort.bam
samtools index <file>.filtered.sort.bam
# Create header file
samtools view -H <file>.filtered.sort.bam > Arabidopsis.header
# Generate mpileup file
samtools mpileup -q 25 -Q 25 -B <file1>.filtered.sort.bam \
<file2>.filtered.sort.bam <file3>.filtered.sort.bam \
<file4>.filtered.sort.bam <file5>.filtered.sort.bam > Arabidopsis.mpileup
Make a .pro file of nucleotide-read quartets (counts of A, C, G, and T) from the .mpileup file.
mapgd proview -i Arabidopsis.mpileup -H Arabidopsis.header \
> Arabidopsis.pro
Run the allele command to estimate allele and genotype frequencies from the .pro file.
mapgd allele -i Arabidopsis.pro -o Arabidopsis -p Arabidopsis.clean
The output will be Arabidopsis.map, which contains a list of estimated genotypic frequency. Test statistics for polymorphism and Hardy-Weinberg disequilibrium, as well as sequencing error rate and population depth of coverage, are also stored in the .map file.
Extract the coverage information from Arabidopsis.map and filter the data based on the population coverage distribution using the filter command, with log-likelihood ratio of polymorphism greater than 20 (-p 20), minor allele frequency between 0.05 and 0.45 (-q 0.05 -Q 0.45), and the population coverage between 10 and 250 (-c 10 -C 250).
awk '{print $6}' Arabidopsis.map > Arabidopsis_coverage.txt
The minimum and maximum coverage cutoffs are determined by the coverage distribution (Figure 1).
Figure 1. Histogram of the population coverage for all SNPs
mapgd filter -i Arabidopsis.map -p 20 -q 0.05 -Q 0.45 -c 10 -C 250 \
-o filtered_Arabidopsis
# -p: minimum value of the likelihood-ratio statistic for polymorphism
# -q: minimum minor-allele frequency estimate
# -Q: maximum minor-allele frequency estimate
# -c: minimum population coverage
# -C: maximum population coverage
Run the genotype command to generate a file of genotype likelihoods.
mapgd genotype -p Arabidopsis.clean.pro -m filtered_Arabidopsis.map \
> Arabidopsis.genotype
For large datasets, the relatedness calculation will take longer. Users could subsample a group of SNPs using sub_sample.py, which will randomly sample N number of SNPs, with N being a user-defined number.
sub_sample.py F_Arabidopsis.genotype -N 200000 \
> 200K_F_Arabidopsis.genotype
Run the relatedness command.
mapgd relatedness -i Arabidopsis.genotype -o Arabidopsis.rel
Data analysis
Result interpretation
The .map files contain a list of estimated genotypic frequency obtained with the allele command. These files store test statistics for polymorphism and Hardy-Weinberg disequilibrium, as well as useful statistics filtering variants, such as sequencing error rate and population depth of coverage (Figure 2). From the .map file, to extract and examine heterozygosity and allele frequency:
awk '{print $1, $2, $7, $16}' filtered_Arabidopsis.map > Arabidopsis_stats.txt
Figure 2. Average heterozygosity across every 100 kb window on chromosome 1
The output of the relatedness command contains the seven genotypic correlation coefficients for all pairs of individuals and some log likelihood ratio test statistics, as described in Ackerman et al. (2017). However, the current version of MAPGD might incur errors when running the relatedness command. Stable results are pending until the next software release on https://github.com/LynchLab/MAPGD. Users may also find example output figures and software performance testing figures at https://github.com/LynchLab/genomics_simulation/tree/master/figures.
Acknowledgments
M.L. is supported by National Science Foundation Division of Environmental Biology grants 1257806, 1927159, and Division of Biological Infrastructure grant 2119963. This protocol was partly adapted from the MAPGD user manual available at https://github.com/LynchLab/MAPGD.
Competing interests
The authors have no conflicts of interest to declare that are relevant to the content of this article.
References
Ackerman, M. S. (2016). Developing quantitative and population genomic techniques to test evolutionary theories. Indiana University.
Ackerman, M. S., Johri, P., Spitze, K., Xu, S., Doak, T. G., Young, K. and Lynch, M. (2017). Estimating Seven Coefficients of Pairwise Relatedness Using Population-Genomic Data. Genetics 206(1): 105–118.
Ackerman, M. S., Maruki, T. and Lynch, M. (2022). MAPGD: A program for the maximum likelihood analysis of population data (0.4.40).
Adams, P. E., Crist, A. B., Young, E. M., Willis, J. H., Phillips, P. C. and Fierst, J. L. (2022). Slow Recovery from Inbreeding Depression Generated by the Complex Genetic Architecture of Segregating Deleterious Mutations. Mol Biol Evol 39(1): e1093/molbev/msab330.
Berardini, T. Z., Reiser, L., Li, D., Mezheritsky, Y., Muller, R., Strait, E. and Huala, E. (2015). The Arabidopsis information resource: Making and mining the “gold standard” annotated reference plant genome. Genesis 53(8): 474–485.
Danecek, P., Bonfield, J. K., Liddle, J., Marshall, J., Ohan, V., Pollard, M. O., Whitwham, A., Keane, T., McCarthy, S. A., Davies, R. M., et al. (2021). Twelve years of SAMtools and BCFtools. GigaScience 10(2): e1093/gigascience/giab008.
Guirao-Rico, S. and González, J. (2021). Benchmarking the performance of Pool-seq SNP callers using simulated and real sequencing data. Mol. Ecol. Resour. 21(4): 1216–1229.
Jeffries, D. L., Mee, J. and Peichel, C. L. (2022). Convergent recruitment of Amh as the sex determination gene in two lineages of stickleback fish. bioRxiv: e477894.
Li, H. (2011). A statistical framework for SNP calling, mutation discovery, association mapping and population genetical parameter estimation from sequencing data. Bioinformatics 27(21): 2987–2993.
Li, H., Handsaker, B., Wysoker, A., Fennell, T., Ruan, J., Homer, N., Marth, G., Abecasis, G., Durbin, R. and 1000 Genome Project Data Processing Subgroup (2009). The Sequence Alignment/Map format and SAMtools. Bioinformatics 25(16): 2078–2079.
Lynch, M., Bost, D., Wilson, S., Maruki, T. and Harrison, S. (2014). Population-genetic inference from pooled-sequencing data. Genome. Biol. Evol. 6(5): 1210–1218.
Lynch, M., Gutenkunst, R., Ackerman, M., Spitze, K., Ye, Z., Maruki, T. and Jia, Z. (2017). Population Genomics of Daphnia pulex. Genetics 206(1): 315–332.
Maruki, T. and Lynch, M. (2014). Genome-wide estimation of linkage disequilibrium from population-level high-throughput sequencing data. Genetics 197(4): 1303–1313.
Maruki, T. and Lynch, M. (2015). Genotype-Frequency Estimation from High-Throughput Sequencing Data. Genetics 201(2): 473–486.
Maruki, T., Ye, Z. and Lynch, M. (2019). Genomic analyses of population structure reveal metabolism as a primary driver of local adaptation in Daphnia pulex. bioRxiv: 807123.
R Core Team. (2022). R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria.
Shoemaker, W. R. (2022). A macroecological perspective on genetic diversity in the human gut microbiome. bioRxiv: e487434
The 1001 Genomes Consortium. (2016). 1,135 genomes reveal the global pattern of polymorphism in Arabidopsis thaliana. Cell 166(2): 481–491.
Ye, Z., Molinier, C., Zhao, C., Haag, C. R. and Lynch, M. (2019). Genetic control of male production in Daphnia pulex. Proc. Natl. Acad. Sci. U. S. A. 116(31): 15602–15609.
Supplementary information
Code availability: all code has been deposited into GitHub: https://github.com/Bio-protocol/MAPGD-for-plants.
Data availability: intermediate and output files were uploaded to Figsuare: https://figshare.com/s/345244bc98fa0e8a6ab1
.
Article Information
Copyright
© 2023 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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4,788 | https://bio-protocol.org/en/bpdetail?id=4788&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Protocol for the High-quality Plasmid Isolation from Different Recalcitrant Bacterial Species: Agrobacterium spp., Rhizobium sp., and Bacillus thuringiensis
PK Preshobha Kodackattumannil *
SS Shina Sasi *
SK Saranya Krishnan
GL Geetha Lekshmi
MK Martin Kottackal
KA Khaled M. A. Amiri
(*contributed equally to this work)
Published: Vol 13, Iss 15, Aug 5, 2023
DOI: 10.21769/BioProtoc.4788 Views: 843
Reviewed by: Hemant Kumar Prajapati Anonymous reviewer(s)
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Abstract
High yield of good quality plasmid DNA from gram -ve bacteria (Agrobacterium tumefaciens, A. rhizogenes, and Rhizobium sp.) and gram +ve bacterium (Bacillus thuringiensis) is difficult. The widely used plasmid extraction kits for Escherichia coli yield a low quantity of poor-quality plasmid DNA from these species. We have optimized an in-house modification of the QIAprep Spin Miniprep kit protocol of Qiagen, consisting of two extraction steps. In the first, the centrifugation after adding neutralization buffer is followed by ethanol (absolute) precipitation of plasmid DNA. In the second extraction step, the precipitated DNA is dissolved in Tris-EDTA (TE) buffer, followed by an addition of 0.5 volumes of 5 M sodium chloride and 0.1 volumes of 20% (w/v) sodium dodecyl sulfate. After incubation at 65 °C for 15 min, the plasmid DNA is extracted with an equal volume of chloroform:isoamyl alcohol (CIA). RNase (20 mg/mL) is added to the upper phase retrieved after centrifugation and is incubated at 37 °C for 15 min. The extraction of the plasmid DNA with an equal volume of CIA is followed by centrifugation and is precipitated from the retrieved upper phase by adding an equal volume of absolute ethanol. The pellet obtained after centrifugation is washed twice with 70% (v/v) ethanol, air dried, dissolved in TE buffer, and quantified. This easy-to-perform protocol is free from phenol extraction, density gradient steps, and DNA binding columns, and yields high-quality plasmid DNA. The protocol opens an easy scale up to yield a large amount of high-quality plasmid DNA, useful for high-throughput downstream applications.
Key features
• The protocol is free from density gradient steps and use of phenol.
• The protocol is an extension of the QIAprep Spin Miniprep kit (Qiagen) and is applicable for plasmid DNA isolation from difficult-to-extract bacterial species.
• The protocol facilitates the direct transformation of the ligation product into Agrobacterium by skipping the step of E. coli transformation.
• The plasmids isolated are of sequencing grade and the method is useful for extracting plasmids for metagenomic studies.
Graphical overview
Overview of the plasmid isolation protocol (modified QIAprep Spin Miniprep kit) of the present study
Keywords: Agrobacterium Bacillus thuringiensis Modified QIAprep Spin Miniprep kit protocol Plasmid DNA Low-copy number bacterial strains Rhizobium Sodium dodecyl sulfate
Background
Plasmid isolation is an essential procedure in gene cloning, gene expression studies, sequencing, mutagenesis, and several downstream molecular processes. Easy-to-extract proprietary kits at different formats such as mini, midi, and maxi, and several published protocols are best suited for the plasmid extraction from widely used Escherichia coli strains. A simple and reliable method for isolating high-quality plasmid for downstream molecular applications from bacteria such as Agrobacterium tumefaciens, A. rhizogenes, Rhizobium sp., and Bacillus thuringiensis is lacking, due to the recalcitrance of the bacterial strains to cell lysis. Besides, lysozyme in the cell lysis solution is required to circumvent the non-lysis of the cell wall (Marmur, 1961). The Agrobacterium cells are not sensitive to the lysozyme-EDTA-detergent lysis procedures (Marmur, 1961; Clewell and Helinski, 1969), necessitating a relatively long treatment with proteolytic enzymes (Zaenen et al., 1974). Further, it is challenging to isolate Ti-plasmid free of chromosomal DNA (Ledeboer et al., 1976). In the case of agrobacteria, which is used to develop genetically modified and genome-edited plants, if there is no stock culture of E. coli with the desired plasmid, it necessitates retransformation of the plasmid into E. coli and subsequent extraction of plasmids for restriction digestion verification (Wise et al., 2006) and other downstream applications, e.g., biolistic transformation. There is no reliable protocol to extract high-quality plasmid DNA in large amounts from agrobacteria to accomplish downstream applications directly, i.e., without E. coli retransformation. The protocols described for the extraction of plasmid DNA from Agrobacterium, Rhizobium, and Bacillus thuringiensis are relatively lengthy and consist of sucrose gradients, CsCl-dye buoyant density gradients, or ethidium bromide treatment followed by phenol extraction (Ledeboer et al., 1976; Adachi and Iyer, 1980; Zhang and Kerr, 1993; Reyes-Ramirez and Ibarra, 2008). Our attempts to isolate plasmid from the gram -ve bacteria (Agrobacterium tumefaciens, A. rhizogenes, Rhizobium sp., and gram +ve bacterium (Bacillus thuringiensis) using kits (QIAprep Spin Miniprep kit from Qiagen and PureLink Quick Plasmid Miniprep kit from Invitrogen) and the user-modified protocol of QIAprep plasmid kit (Weber et al. 1998; https://www.qiagen.com/no/resources/resourcedetail?id=95083ccb-9583-489e-b215-99bd91c0604e&lang=en) did not yield satisfactory results. Extracting and purifying plasmid DNA from these strains and diverse other bacterial strains, recalcitrant to quality plasmid isolation, warrant a simple, short, and reliable protocol. We believe that the present study’s high-yielding, high-quality plasmid DNA protocol will be useful for other bacterial strains resistant to cell lysis, especially for low-copy number plasmid strains.
Materials and reagents
Biological materials
Agrobacterium tumefaciens strain EHA105, AGL1, GV3101, and LBA4404 [all containing binary plasmid pCAMBIA 1201 harboring gusA gene under the control of Cauliflower Mosaic Virus (CaMV) 35S promoter and bacterial selection marker chloramphenicol]
A. rhizogenes strains ATCC15834 and A4, containing binary plasmid harboring green fluorescent protein (GFP) gene under the control of 35S promoter and bacterial selection marker kanamycin
Rhizobium sp. (isolated from root nodules of desert tree legume, Prosopis cineraria)
Bacillus thuringiensis (received as a gift from a colleague, collected from the United Arab Emirates)
Reagents
Luria-Bertani (LB) broth (LB Miller Modification; PhytoTech Labs, catalog number: L475)
Tryptone (PhytoTech Labs, catalog number: T832)
Yeast extract (PhytoTech Labs, catalog number: Y892)
Sodium chloride (NaCl) (Sigma, catalog number: S1679)
Potassium dihydrogen phosphate dibasic (KH2PO4) (Sigma, catalog number: P3786)
Magnesium sulfate heptahydrate (MgSO4·7H2O) (Sigma, catalog number: M2773)
Mannitol (Sigma, catalog number: M1902)
Bacto-agar (Himedia, catalog number: GRMO26)
Kanamycin sulfate (PhytoTech Labs, catalog number: K378)
Rifampicin (PhytoTech Labs, catalog number: R7382)
Formamide (Sigma, catalog number: 47671)
Chloramphenicol (PhytoTech Labs, catalog number: C1919)
QIAprep Spin Miniprep kit (Qiagen, catalog number: 27109)
PureLink Quick Plasmid Miniprep kit (Invitrogen, catalog number: K210011)
Resuspension solution P1 with RNase and Blue lysate (Qiagen, catalog number: 19051)
Resuspension buffer R3 (Invitrogen, catalog number: K2100-14)
Lysis solution P2 (Qiagen, catalog number: 19052)
Lysis buffer L7 (Invitrogen, catalog number: K2100-14)
Neutralizing solution N3 (Qiagen, catalog number: 19064)
Precipitation buffer N4 (Invitrogen, catalog number: K2100-14)
Sodium dodecyl sulfate (SDS) (Sigma, catalog number: L3771)
RNase (Thermo Scientific, catalog number: EN0531)
Ethanol absolute (Sigma, catalog number: NC1971050)
Disodium ethylene diamine tetraacetate (Na2EDTA·2H2O) (Sigma, catalog number: E6760)
Tris-base (Sigma, catalog number: T1503)
Hydrochloric acid (HCl) (Sigma, catalog number: 07102)
Sodium hydroxide (NaOH) (Sigma, catalog number: 06203)
Chloroform (Sigma-Aldrich, catalog number: C2432)
Glacial acetic acid (VWR, catalog number: VWRC20104.334)
Isoamyl alcohol (Sigma, catalog number: 19392)
Methanol ChromasolvTM (Sigma-Aldrich, catalog number: 34860-2.5L-R)
NcoI-HF (NEB, catalog number: R3193M), storage at -20 °C
BstEII-HF (NEB, catalog number: R3162M), storage at -20 °C
HotStar Taq DNA Polymerase PCR kit (Qiagen, catalog number: 203205)
PCR primers (Macrogen, South Korea)
Agarose D1 Low CE (Conda Lab, catalog number: 8010.00)
HydragreenTM Safe DNA Dye (20,000×) (ACTGene, catalog number: ACT-IDMG04)
6× DNA loading dye (NEB, catalog number: B7024A)
GeneRuler 1 kb Plus DNA ladder (Thermo Fisher Scientific, catalog number: SM1333)
Lambda DNA/EcoRI Plus HindIII marker (Thermo Fisher Scientific, catalog number: SM0191)
Solutions
1 N HCl
1 N NaOH
LB Agrobacterium medium (see Recipes)
Yeast Mannitol medium (see Recipes)
Tris-EDTA (TE; see Recipes)
Tris (see Recipes)
Na2EDTA (see Recipes)
Kanamycin (see Recipes)
Rifampicin (see Recipes)
Tris-acetate-EDTA (TAE, see Recipes)
Chloramphenicol (see Recipes)
5 M Sodium chloride (see Recipes)
20% (w/v) SDS (see Recipes)
Chloroform:isoamyl alcohol (CIA) (see Recipes)
Ethanol (70%, v/v) (see Recipes)
Media components (1 L) for the bacterial culture (see Recipes)
Antibiotics preparation (see Recipes)
Stock solutions (see Recipes)
Tris-EDTA (TE) buffer (100 mL) (see Recipes)
10× Tris-acetate-EDTA (TAE) (see Recipes)
Agarose gel (0.8%, w/v) preparation (see Recipes)
Recipes
Media components (1 L) for the bacterial culture
Components LB Agrobacterium (g/L) LB (g/L) YM (g/L)
Tryptone 10 (5 g for LBA4404 strain) 10
Yeast extract 5 5 0.1
NaCl 5 10 0.2
KH2PO4 10
MgCl2·7H2O 0.2
Mannitol 10
Dissolve the components in 800 mL of MilliQ water and adjust to 1 L using MilliQ water. Transfer into a Duran bottle. Adjust the pH to 7.0 using 1 N HCl/1 N NaOH.
For solid media, add bacto-agar 15 g/L after adjusting the pH.
Autoclave at 121 °C (15 psi) for 15 min and store at room temperature (RT) for two months.
Antibiotics preparation
Antibiotics Preparation and storage Working concentration
Kanamycin Weigh 50 mg in a 1.5 mL Eppendorf, add 1 mL of sterile water, dissolve, and store at 4 °C for one week. 50 mg/L
Chloramphenicol Weigh 25 mg in a 1.5 mL Eppendorf, add 1 mL of absolute ethanol, dissolve, and store at 4 °C for one week. 25 mg/L
Rifampicin Weigh 20 mg in a 1.5 mL Eppendorf, add 1 mL of formamide*, dissolve and warp with aluminum foil, and store at 4 °C for one week. 20 mg/L
*Methanol is an alternative solvent.
Stock solutions
Stock Preparation and storage
1 M Tris-HCl (pH 8.0) Dissolve 121.1 g of Tris-base in 800 mL of MilliQ water. Adjust the pH to 8.0 by adding concentrated HCl. Adjust to 1 L using MilliQ water. Transfer into a 1 L Duran bottle. Autoclave at 121 °C (15 psi) for 15 min. Store at RT for six months.
0.5 M EDTA (pH 8.0) Weigh 93 g of disodium EDTA·2H2O in ~400 mL of MilliQ. Stir vigorously on a magnetic stirrer with a stir bar. Adjust the pH to 8.0 with NaOH (~10 g of NaOH pellets). Adjust to 500 mL using MilliQ water. Transfer into a 500 mL Duran bottle. Autoclave at 121 °C (15 psi) for 15 min. Store at RT for six months.
5 M NaCl Dissolve 29.2 g of NaCl in 80 mL of MilliQ water and adjust to 100 mL using MilliQ water. Transfer into a 100 mL Duran bottle. Autoclave at 121 °C (15 psi) for 15 min. Store in a Duran bottle at RT for six months.
SDS (20%, w/v) Weigh 20 g of SDS, dissolve it in MilliQ water, and adjust to 100 mL using MilliQ water. Transfer into a 100 mL Duran bottle. Autoclave at 121 °C (15 psi) for 15 min. Store at RT for six months. Caution: use face masks during preparation.
Chloroform:isoamyl alcohol (24:1) (CIA) Add 2 mL of isoamyl alcohol in a 50 mL sterile Falcon tube and adjust to 50 mL by adding chloroform. Mix well and cover with aluminum foil. Store at RT for two weeks. Caution: prepare in a fume hood.
70% (v/v) ethanol Measure 35 mL of absolute ethanol in a 50 mL sterile Falcon tube, add 15 mL MilliQ water, and mix well. Store at RT for six months.
Tris-EDTA (TE) buffer (100 mL)
Components Volume (mL) Final concentration
Tris-HCl (pH 8.0) 1.0 10 mM
Na2EDTA (pH 8.0) 0.2 1 mM
MilliQ water 98.8
Autoclave in a 100 mL Duran bottle at 121 °C (15 psi) for 15 min and store at RT for six months.
10× Tris-acetate-EDTA (TAE)
Components Amount Concentration 1× 1× preparation
Tris-base 48.5 g 40 mM Mix 10 mL of 10× with 90 mL
Na2EDTA (pH 8.0) 20 mL 1 mM MilliQ water
Glacial acetic acid 11.4 mL 20 mM
Dissolve tris-base in approximately 800 mL of MilliQ water. Add acetic acid and EDTA. Adjust to 1 L. Transfer into 1 L Duran bottle. Store at RT for six months.
Agarose gel (0.8%, w/v) preparation
Weigh 0.48 g of agarose and add into 60 mL of 1× TAE buffer in a 250 mL Erlenmeyer flask. Microwave for 2 min, cool to 50 °C, add 6 μL of Hydragreen, and cast in the gel tray with 15 wells combs.
Laboratory supplies
90 mm Petri dish (Sangon Biotech, catalog number: F611001)
Safe-lock tubes (2.0 and 1.5 mL) (Eppendorf, catalog number: 0030 120.094)
Pipettes (1,000, 200, 100, 20, and 10 μL) (Rainin, catalog number: L1000XLS+, L200XLS+, L100XLS+, L20XLS+, and L10XLS+, respectively)
Duran bottle [DURAN® ORIGINAL GL 45, catalog number: 10113399 (1,000 mL) and 10108298 (500 mL)]
Beakers (0.6 and 1 L) (Nalgene, catalog number: 1201-0600 and 1201-1000, respectively)
PYREX® 250 mL narrow mouth Erlenmeyer flask (Corning, catalog number: 4980-250)
Graduated cylinders (0.5 and 1 L) (Nalgene, catalog number: 3662-0500 and 3662-1000, respectively)
Pipette tips (1,000, 250, and 20 μL) (Rainin, catalog number: 30389294, 30389301, and 30389297, respectively)
Magnetic stirrer (Corning®, model: PC-620D, catalog number: 6796-620D)
Stir bars (PhytoTech Labs, catalog number: B011)
Tube racks (Globe Scientific, catalog number: 456350B)
Kimwipes (KIMTECH, catalog number: 34120)
14 mL sterile Falcon bacterial culture tubes (Falcon, catalog number: 352057)
Disposable gloves (Kimberly-Clark, catalog number: 52816)
Inoculation loops (VWR, catalog number: 10806-354)
Wide Mini-Sub Cell GT Horizontal Electrophoresis System, 15 cm × 7 cm tray, with gel caster and casting gates (Bio-Rad Laboratories, catalog number: 1704469)
Weigh dish (Thomas Scientific, catalog number: 3846D19)
Weigh paper (Thomas Scientific, catalog number: 9885G50)
Spatula (PhytoTech Labs, KS, USA, catalog number: S798)
Face mask (Euromed, China, catalog number: EM1834)
Disposable cuvettes (BrandTech, catalog number: 759075D)
Equipment
Autoclave (Tomy, model number: SX-700)
Weighing scale (Mettler Toledo®, model: MS603S/01)
Refrigerator (4 and -20 °C) (Evermed, model: BLCRF-370W)
pH meter (Thermo Fisher, model: Fisherbrand Accumet AB150, catalog number: 13636AB150B)
Laminar air flow cabinet (Esco, Horizontal Laminar Flow Cabinet, Gen 3, model: LHG-3AG-F8, catalog number: 2120387)
Bead sterilizer (PhytotTech Labs, model: ErgoSteri VT glass bead sterilizer, catalog number: S7520)
Shaker incubator (Eppendorf, model: New Brunswick Innova® 42, catalog number: M-1335-0004)
UV-Visible spectrophotometer (Thermo Scientific, model: Evolution 201, catalog number: 840-210600)
Centrifuge (Tomy, model: MX-307)
Vortexer (Scientific industries, model: Vortex-GenieTM 2T mixer, catalog number: SI-T266)
Heating block (Labnet International, AccuBlock, Digital dry bath, model: Labnet International D1100, catalog number: D1100-230V)
Fume hood (Esco, AscentTM Max Ductless Fume Hood, model: ADC-4BI, catalog number: 2040042)
Nutating Mixer (Labnet International, model: Labnet GyroMiniTM Nutating 3-D Mixer with dimpled mat, 120 V, catalog number: S0500)
NanoDropTM (Thermo Scientific, catalog number: ND2000C)
Microwave (Samsung, model: ME733K)
MilliQ water purification system (MilliQ, model: Elix integral 10)
Gel DocTM EZ imager (Bio-Rad Laboratories, catalog number: 170-8270)
PowerPacTM Basic (Bio-Rad Laboratories, catalog number: 164-5050)
Electrophoresis unit (Bio-Rad Laboratories, catalog number: 1704405)
Desktop (Dell, model: Optoplex 9020)
Thermal cycler (Bio-Rad Laboratories, model: T100)
Software and datasets
NanoDrop 2000 (version 1.6, free, Thermo Fisher Scientific)
Image Lab 6.0 (Bio-Rad Laboratories)
Insight2 (Thermo Fisher Scientific, catalog number: 837-002700)
Microsoft Excel for Mac (v16.69.1, 2022)
Procedure
Note: All steps are carried out at room temperature (RT) unless otherwise stated.
Streak the bacterial species: EHA105, AGL1, GV3101, and LBA4404 strains on solid LB Agrobacterium media supplemented with 20 mg/L rifampicin and 25 mg/L chloramphenicol; Bacillus thuringiensis on LB solid medium; A. rhizogenes strains ATCC15834 and A4 on YM solid medium with 50 mg/L kanamycin; and Rhizobium sp. on YM solid medium (see Recipes). Incubate for two days at 28 °C in an incubator.
Transfer bacterial culture from step 1 into 5 mL of liquid media in 14 mL bacterial culture tubes with appropriate antibiotics as above specified to each strain and incubate at 28 °C overnight on an orbital shaker at 250 rpm.
Note: Transfer a single colony for A. tumefaciens, Rhizobium, and Bacillus, and 2–3 colonies of A. rhizogenes from the solid medium to 5 mL liquid medium. Check the OD of the culture using the spectrophotometer; use culture media with no bacteria as blank. Cultures of A. tumefaciens with OD 0.5–0.7 and an OD of 0.8–1.0 for other strains are used for plasmid extraction. A 10 mL culture of slow-growing or low-copy number plasmid results in a high yield.
Centrifuge the cultures at 9,000× g (8,000 rpm) for 5 min.
Decant the supernatant. Remove the liquid media completely by pipetting out or by keeping the tube upside-down on a Kimwipe for 2–5 min. (Caution: do not disturb the pellet.)
Add 250 μL of resuspension solution (P1 buffer containing RNase and Blue lysate of QIAprep Spin Miniprep kit) into each tube and vortex to resuspend the cell pellet completely.
Transfer the resuspended bacterial pellet into 1.5 mL Eppendorf tubes using a pipette.
Add 250 μL of lysis solution (P2 buffer of QIAprep Spin Miniprep kit) and mix by gently inverting the tubes 5–6 times. The solution quickly turns blue and viscous, indicating bacterial lysis.
Add 350 μL of neutralizing solution (N3 buffer of QIAprep Spin Miniprep kit) and mix by inverting the tubes 5–6 times. The blue solution turns white, and the bacterial chromosomal DNA appears as a white precipitate.
Centrifuge the tubes at 20,000× g (15,000 rpm) for 10 min.
Carefully transfer the supernatant into new 1.5 mL Eppendorf tubes using a pipette. (Caution: do not disturb the white precipitate.)
Add an equal volume of absolute ethanol into each tube and mix by inverting the tubes a few times.
Centrifuge the tubes at 20,000× g (15,000 rpm) for 5 min.
Decant the supernatant and carefully remove the liquid completely using a pipette or by leaving the tube upside-down on a Kimwipe for 1–3 min.
Add 300 μL of TE buffer (pH 8.0) (see Recipes), 0.5 vol (150 μL) of 5 M NaCl, and 0.1 vol (45 μL) of 20% SDS. Mix by inversion and incubate at 65 °C in a heating block for at least 10 min.
Note: Make sure the DNA is completely dissolved in TE. If not, increase the volume of TE. If using 10% SDS, add 0.2 vol.
Add an equal volume (500 μL) of CIA (see Recipes) and keep shaking on a Nutating Mixer for at least 5 min.
Centrifuge at 20,000× g (15,000 rpm) for 7–10 min.
Carefully transfer the upper phase (~400 μL) into new 1.5 mL Eppendorf tubes, add 16 μL (20 mg/mL) of RNase, mix well by inversion, and incubate at 37 °C in a heated block for at least 15 min. Add an equal volume of CIA and keep shaking on a Nutating Mixer for at least 5 min.
Note: Take the heat block and keep it at RT after step 14 for a few minutes. Incubation of the vial after adding RNase in the block will also work.
Centrifuge at 20,000× g (15,000 rpm) for 7–10 min.
Carefully transfer the upper phase (300 μL) into new 1.5 mL Eppendorf tubes, add an equal volume of absolute ethanol, and mix well by inversion.
Centrifuge at 20,000× g (15,000 rpm) for 7–10 min.
Carefully decant or pipette out the liquid phase and add 500 μL of 70% ethanol. Tap the tube to release the pellet.
Centrifuge at 20,000× g (15,000 rpm) for 1–2 min and repeat step 21.
Centrifuge as above, pipette out the liquid completely, and air dry by keeping the tube open at RT for 2–5 min.
Add 50 μL (if the pellet is large, add more volume) of TE. Completely dissolve the pellet by finger tapping.
Measure the quantity of the plasmid DNA using NanoDrop (Figure 1A–1D; Table 1).
Figure 1. NanoDrop quantitation of the isolated plasmids. A. A4; B. Rhizobium; C. EHA105; and D. Bacillus thuringiensis. The x-axis represents the wavelength (nM), and the y-axis represents the 10 mm absorbance.
Table 1. Purity and quantity of the plasmid isolated from different bacterial strains using Qiagen protocol for Agrobacterium (Weber et al., 1998) and the optimized protocol
Bacterial strain A260/280 ratio A260/230 ratio Yield (ng/μL)
Control (Qiagen) Optimized protocol Control
(Qiagen)
Optimized protocol
Control
(Qiagen)
Optimized protocol
A. tumefaciens EHA105 1.61 ± 0.03 1.91 ± 0.08 1.60 ± 0.04 2.10 ± 0.05 32 ± 1.8 232.4 ± 10.5
A. tumefaciens AGL1 1.40 ± 0.19 2.0 ± 0.05 1.52 ± 0.10 2.21 ± 0.1 24 ± 4.8 185 ± 8.6
A. tumefaciens GV3101 1.51 ± 0.07 1.93 ± 0.06 1.51 ± 0.07 2.09 ± 0.08 34 ± 1.4 212 ± 9.2
A. tumefaciens LBA4404 1.38 ± 0.14 2.05 ± 0.07 1.40 ± 0.10 1.98 ± 0.07 26 ± 3.4 176 ± 7.2
A. rhizogenes A4 1.49 ± 0.05 1.97 ± 0.08 1.49 ± 0.05 2.28 ± 0.1 22 ± 2.0 192.2 ± 9.8
A. rhizogenes ATCC15834 1.36 ± 0.11 1.99 ± 0.05 1.21 ± 0.08 2.15 ± 0.09 20 ± 3.1 183.1 ± 9.1
Rhizobium sp. 1.59 ± 0.03 1.89 ± 0.08 1.53 ± 0.04 2.11 ± 0.07 37 ± 4.3 372.0 ± 11.2
Bacillus thuringiensis 1.47 ± 0.12 1.91 ± 0.04 1.38 ± 0.11 2.10 ± 0.09 31 ± 3.5 211.7 ± 10.8
E. coli* 1.94 ± 0.06 - 2.09 ± 0.05 245 ± 8.1 -
Data represent the mean ± SE of five replicates. The purity and quantity of plasmid isolated using PureLink Quick Plasmid Miniprep kit, which also served as the control, are not significantly different from that of the Qiagen protocol; hence, it is not shown. The same plasmid of that in A. tumefaciens strains in E. coli served as the positive control*.
Load 100 ng of plasmid DNA mixed with 6× loading dye (1× final) on a 0.8% agarose gel and run at 100 V for 25 min (Figure 2).
Document the gel using the Gel DocTM EZ Imager with the Image Lab 6.0 program.
Figure 2. Gel electrophoresis of the isolated plasmids from different bacterial strains. EHA: EHA105, LBA: LBA4404, GV: GV3101, and AGL: AGL1 (all are A. tumefaciens); A4: A. rhizogenes A4; ATC: A. rhizogenes ATCC15834; Rhi: Rhizobium; Bt: Bacillus thuringiensis (shows two bands). PC: positive control of the same plasmid from E. coli using QIAprep Spin Miniprep kit; M: Lambda DNA/EcoRI Plus HindIII marker.
Data analysis
The quantity of the isolated plasmid DNA is determined using NanoDrop 2000 (Table 1; Figure 1A–1D). The data analyzed using Microsoft Excel represents the mean ± SE of five replicates (Table 1).
Validation of protocol
The protocol is repeated five times for each sample, once with a 10 mL culture of each bacterial strain, and control for each sample is carried out. Figure 2 shows the electrophoresis of the isolated plasmid DNA in an agarose (0.8%, w/v) gel in TAE containing Hydragreen (a noncarcinogenic nucleic acid stain), carried out in a Bio-Rad electrophoresis system, documented using the Gel DocTM EZ imager with the Image Lab 6.0 program. The quality of the plasmid DNA isolated in the present protocol is validated by gel electrophoresis (Figure 2) and restriction digestion (Figure 3): double digestion of 1 μg of plasmid DNA (pCAMBIA 1201) using NcoI-HF and BstEII-HF following the manufacturer’s double-digest protocol (NEB, USA). The digestion resulted in the gusA gene fragment of 2,060 bp (Figure 3). PCR is carried out using isolated plasmids with specific primers (Macrogen, Korea). Amplification of the gusA gene (Figure 4) of the extracted plasmid DNA from A. tumefaciens strains is carried out as per Kodackattumannil et al. (2023). PCR of the extracted Rhizobium plasmid with the primer pairs fD1 – rP2 for 16S rRNA (Weisburg et al., 1991) and R16-1 – R23-3R for ITS region (Figure 4) is performed as described by Kwon et al. (2005). Amplification of the GFP gene in the isolated plasmid DNA from A. rhizogenes strains (Figure 4) is accomplished according to Dutta et al. (2013). PCR of the Cry3a gene using the primers (forward: ATGAATCCGAACAATCGAAG and reverse: TTAATTCACTGGAATAAATTCAATTTTG) is carried out with an initial denaturation of 95 °C for 15 min, 30 cycles of denaturation at 94 °C for 30 s, annealing at 62 °C for 30 s, and extension at 72 °C for 2 min, and followed by a final extension at 72 °C for 10 min. In all cases, the PCR is carried out using HotStar Taq polymerase kit following the manual. The transient expression of GFP (plasmid isolated from ATCC15834) in indica rice IR5 callus (Figure 5), checked using particle bombardment (PDS-1000/Bio-Rad), validated the downstream application following the protocol of Sarangi et al. (2019).
Note: The isolated plasmid with the GFP gene (from ATCC15834) using the Qiagen protocol is not of good quality, and the quantity is very low (Table 1), which is insufficient to perform the particle bombardment.
Figure 3. Restriction digestion of the plasmids isolated from A. tumefaciens strains EHA105, GV3101, and E. coli (positive control) containing the binary plasmid pCAMBIA 1201 harboring gusA (2,060 bp) gene under the control of 35S promoter (digestion of gusA gene using the restriction enzymes NcoI and BstEII (NEB). 1. EHA105; 2. GV3101; 3. E. coli; 4, 5, and 6 are undigested. M—1 Kb Plus DNA ladder.
Figure 4. PCR analysis of the isolated plasmids from different bacterial strains. 1, 2, 3, and 4: gusA (353 bp), A. tumefaciens strain EHA105, GV3101, LBA4404, AGL1, and E. coli (PC1); 5 and 6: GFP (800 bp) from A. rhizogenes A4 and ATCC15834, respectively; 7 and 8: Rhizobium with R16-1–R23-3R, and fD1–rP2 primer pairs (1,500 bp); 9: amplification of cry3A gene (1,935 bp) from Bacillus thuringiensis. PC2: positive control of GFP; NC: negative control of the respective primers; M: 1 Kb Plus DNA ladder.
Figure 5. Green fluorescent protein expression on indica rice IR5 callus bombarded (using PDS-1000/Bio-Rad) as per the protocol of Sarangi et al. (2019) with the plasmid (harboring GFP gene under the control of 35S promoter) isolated from A. rhizogenes ATCC15834. Image is captured using Leica Thunder Model Organism Microscope with GFP filters (excitation 488 nm, emission 507 nm). Scale bar: 0.1 cm.
General notes and troubleshooting
Troubleshooting
Problems Troubleshooting
The quality of plasmid DNA is not good a. Completely remove the bacterial culture medium before the addition of the P1 buffer.
b. Completely remove the ethanol before the addition of TE in the second extraction.
SDS carry over Do not use more than 0.1 vol of 20% SDS.
RNA contamination Incubate at 37 °C for 30 min.
Acknowledgments
We acknowledge The Presidential Court, the United Arab Emirates, for financial support.
Author contributions statement: MK designed the experiment. PK and SS optimized the protocol. SK and GL validated the protocol. MK and PK drafted the manuscript. KA supervised the project. All the authors reviewed the manuscript.
Competing interests
The authors declare no conflict of interest by any means.
Ethical considerations
The protocol has no animal or human subjects.
References
Adachi, T. and Iyer, V. (1980). A procedure for the isolation and purification of plasmid DNA from Rhizobiummeliloti. Anal. Biochem. 101(2): 271–274.
Clewell, D. B. and Helinski, D. R. (1969). Supercoiled circular DNA-protein complex in Escherichia coli: Purification and induced conversion to an open circular DNA from. Proc. Natl. Acad. Sci. U.S.A. 62(4): 1159–1166.
Dutta, I., Kottackal, M., Tumimbang, E., Tajima, H., Zaid, A. and Blumwald, E. (2013). Sonication-assisted efficient Agrobacterium-mediated genetic transformation of the multipurpose woody desert shrub Leptadenia pyrotechnica. Plant Cell, Tissue Organ Cult. 112(3): 289–301.
Kodackattumannil, P., Whitley, K., Sasi, S., Lekshmi, G., Krishnan, S., Al Senaani, S., Kottackal, M. and Amiri, K. M. A. (2023). Novel inducible promoter DREB1G cloned from date palm exhibits high fold expression over AtRD29 to drought and salinity stress. Plant Cell, Tissue Organ Cult.: e1007/s11240-023-02460-3.
Kwon, S. W., Park, J. Y., Kim, J. S., Kang, J. W., Cho, Y. H., Lim, C. K., Parker, M. A. and Lee, G. B. (2005). Phylogenetic analysis of the genera Bradyrhizobium, Mesorhizobium, Rhizobium and Sinorhizobium on the basis of 16S rRNA gene and internally transcribed spacer region sequences. Int. J. Syst. Evol. Microbiol. 55(1): 263–270.
Ledeboer, A., Krol, A., Dons, J., Spier, F., Schilperoort, R., Zaenen, I., van Larebeke, N. and Schell, J. (1976). On the isolation of TI-plasmid from Agrobacterium tumefaciens. Nucleic Acids Res. 3(2): 449–464.
Marmur, J. (1961). A procedure for the isolation of deoxyribonucleic acid from micro-organisms. J. Mol. Biol. 3(2): 208–218, IN1.
Reyes-Ramírez, A. and Ibarra, J. E. (2008). Plasmid Patterns of Bacillus thuringiensis Type Strains. Appl. Environ. Microbiol. 74(1): 125–129.
Sarangi, S., Mandal, C., Dutta, S., Mukherjee, P., Mondal, R., Kumar, S. J., Choudhury, P. R., Singh, V. P., Tripathi, D. K., Mandal, A. B., et al. (2019). Microprojectile based particle bombardment in development of transgenic indica rice involving AmSOD gene to impart tolerance to salinity. Plant Gene 19: 100183.
Weber, S., Horn, R. and Friedt, W. (1998). Isolation of a low-copy plasmid from Agrobacterium using QIAprep technology. QIAGEN News, number: 5, 7.
Weisburg, W. G., Barns, S. M., Pelletier, D. A. and Lane, D. J. (1991). 16S ribosomal DNA amplification for phylogenetic study. J. Bacteriol. 173(2): 697–703.
Wise, A. A., Liu, Z. and Binns, A. N. (2006). Nucleic acid extraction Agrobacterium strains. In: Wang, K. (Ed.). Agrobacterium Protocols (pp. 67–76). Methods in Molecular Biology. Humana Press.
Zaenen, I., van Larebeke, N., Teuchy, H., van Montagu, M. and Schell, J. (1974). Supercoiled circular DNA in crown-gall inducing Agrobacterium strains. J. Mol. Biol. 86(1): 109–127.
Zhang, L. and Kerr, A. (1993). Rapid purification of Ti plasmids from Agrobacterium by ethidium bromide treatment and phenol extraction. Lett. Appl. Microbiol. 16(5): 265–268.
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A Method for Studying Social Signal Learning of the Waggle Dance in Honey Bees
SD Shihao Dong *
TL Tao Lin *
JN James C. Nieh
KT Ken Tan
(*contributed equally to this work)
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4789 Views: 457
Reviewed by: Komuraiah MyakalaDipesh Kumar VermaRubikah Vimonish
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Original Research Article:
The authors used this protocol in Science Mar 2023
Abstract
Honey bees use a complex form of spatial referential communication. Their waggle dance communicates to nestmates the direction, distance, and quality of a resource by encoding celestial cues, retinal optic flow, and relative food value into motion and sound within the nest. This protocol was developed to investigate the potential for social learning of this waggle dance. Using this protocol, we showed that correct waggle dancing requires social learning. Bees (Apis mellifera) that did not follow any dances before they first danced produced significantly more disordered dances, with larger waggle angle divergence errors, and encoded distance incorrectly. The former deficits improved with experience, but distance encoding was set for life. The first dances of bees that could follow other dancers had none of these impairments. Social learning, therefore, shapes honey bee signaling, as it does communication in human infants, birds, and multiple other vertebrate species. However, much remains to be learned about insects’ social learning, and this protocol will help to address knowledge gaps in the understanding of sophisticated social signal learning, particularly in understanding the molecular bases for such learning.
Key features
• It was unclear if honey bees (Apis mellifera) could improve their waggle dance by following experienced dancers before they first waggle dance.
• Honey bees perform their first waggle dances with more errors if they cannot follow experienced waggle dancers first.
• Directional and disorder errors improved over time, but distance error was maintained. Bees in experimental colonies continued to communicate longer distances than control bees.
• Dancing correctly, with less directional error and disorder, requires social learning.
• Distance encoding in the honey bee dance is largely genetic but may also include a component of cultural transmission.
Keywords: Honey bee Waggle dance Referential communication Social learning Experience
Background
Social learning, the process of learning by observing or interacting with others, is crucial for complex behaviors and adapting these to specific environmental circumstances (Leadbeater and Chittka, 2007). Honey bee workers learn resource location and quality through the waggle dance, a sophisticated form of spatial referential communication (von Frisch, 1967). However, it was unknown if dance following could improve the performances of young waggle dancers or if the dance was entirely innate. Prior research and protocols focused on describing the waggle dance in experienced or novice bees but did not deprive them of the ability to follow waggle dances before they first danced. This deprivation, in which honey bees were unable to use social learning of waggle dancing, has not previously been described as a protocol or used to investigate waggle dancing.
Our protocol, therefore, explains how to design an experiment in which bees cannot follow other waggle dancers before they begin to dance (Dong et al., 2023). These findings indicate that social signal learning can enhance the accuracy of the waggle dance. When compared to bees that were exposed to waggle dancing, naive bees that could not observe dances prior to their first dance displayed larger divergence angle errors, signaled longer distances, and demonstrated significantly more disarray in their dances. However, when these same bees were 20 days old and had gained experience with dance following and dancing, they noticeably reduced divergence angle errors and performed more coordinated dances. Despite these improvements, they were consistently unable to accurately encode distance. These corrections in the experimental older dancers, as compared to their first dances, could be attributed to increased age, more experience following dances, additional practice with flight and foraging, or a combination thereof. Control bees only managed to reduce distance errors as they aged. Having the opportunity to observe experienced dancers before their first dance was enough for the control bees to have the lower directional errors characteristic of older, experienced bees (Dong et al., 2023).
This setup can be employed by other researchers to further explore the role of social learning in eusocial insects and its impact on communication and decision-making within colonies. Such experiments can provide a foundation for designing studies that investigate various aspects of social learning, including the following:
Comparisons across species: by conducting similar experiments on different species of eusocial insects, researchers can compare the extent and mechanisms of social learning between species, as well as the influence of genetic and environmental factors on the development of complex behaviors and communication.
Impact of social learning on colony performance: researchers can investigate the relationship between the quality of social learning and the overall performance of the colony, including foraging efficiency, resource allocation, and reproductive success. This can help to identify the benefits of social learning in colony organization and decision making.
Development of individual learning abilities: by manipulating the availability of social learning opportunities, researchers can study the development of individual learning abilities in eusocial insects and determine the importance of social learning in the acquisition and refinement of specific skills.
Neural and molecular mechanisms: investigating the neural and molecular mechanisms underlying social learning in eusocial insects can help to elucidate the cognitive and physiological processes that facilitate the acquisition of complex behaviors and communication strategies.
Social learning and colony adaptation: by examining the impact of social learning on the ability of colonies to adapt to changing environments, researchers can better understand the role of social learning in the evolution and resilience of eusocial insect societies.
Artificial intelligence and robotics: insights gained from studying social learning in eusocial insects can be applied to the development of artificial intelligence and swarm robotics in which decentralized decision-making and communication strategies are crucial for efficient problem-solving and adaptation.
By using this experimental setup as a starting point, researchers can expand our understanding of social learning in eusocial insects and its broader implications for evolution, ecology, and technology.
Location and timing
The following are instructions on the location and timing of the experiments:
Experiments were carried out at the Southwest Center for Biological Diversity, Chinese Academy of Sciences (Kunming, China). These experiments can be conducted at multiple different locations that support healthy Apis mellifera colonies.
Carry out the experiments during the months of April–June when conditions are good. Note that these months were chosen because the temperature differences between day and night are moderate and facilitate the survival of young bee colonies. The correct months to conduct such experiments will vary depending on local conditions.
Materials and reagents
Biological materials
Ten Apis mellifera colonies (five control and five experimental) were studied at a bee apiary. Queens from all colonies should be obtained from the same breeder to ensure greater genetic uniformity.
Materials
Graduated cylinder, 1,000 mL (Fisher ScientificTM, catalog number: 08-559GC) or volumetric flask with seal, 1,000 mL (Fisher ScientificTM, catalog number: 10-200G) (includes the flask and seal)
ParafilmTM wax sheet (BemisTM PM999, Fisher ScientificTM, catalog number: 13-374-12)
Reagents
Sucrose
ddH2O
Sucrose solution (see Recipes)
Recipes
Sucrose solution
Reagent Final concentration Quantity
Sucrose (100%) 55% (w/v) 55 g
ddH2O n/a n/a
Total n/a 100 mL (final volume)
To make a standard 55% (w/v) sucrose solution, weigh out 55 g of reagent-grade sucrose with an electronic balance.
Add the sucrose to a graduated cylinder or volumetric flask and then double-distilled water (ddH2O) until reaching a volume of 75 mL. You may also use distilled water.
Shake the vessel to dissolve the sugar. For a graduated cylinder, cover the opening with a Parafilm wax sheet and carefully invert it multiple times to dissolve. For a volumetric flask, use the seal that comes with the flask to seal the flask and then invert the sealed flask multiple times.
Add more ddH2O until the total volume is 100 mL.
The water should be freshly obtained but does not have an expiration date if it comes from a sterile, sealed container.
Similarly, the dry sucrose does not have an expiration date, provided that the manufacturer’s expiration date is not exceeded.
It is recommended to prepare this solution freshly each day and use it at room temperature (21 °C). Over time, this sucrose solution can begin to grow mold.
If necessary, the prepared solution can be kept at 4 °C for 24 h. Discard after 24 h.
Equipment
High-definition video camera (HDR-PJ790, Sony)
ncubator (PRX-250B, Ningbo Saifu Experimental Instrument Co., Ltd., https://www.chem17.com/product/detail/37456106.html)
Aspirator to capture bees (custom-made):
A manual aspirator is best because the suction can be controlled by the user.
To make such an aspirator, take a clear plastic cylindrical container that is 14 cm in diameter with a removable lid. Drill the lid to add two holes, spaced 10 cm apart and 1 cm in diameter.
Obtain two pieces of clear vinyl tubing that are 1 cm in outer diameter. The mouthpiece tube should be 30 cm long and the bee collection tube should be 60 cm long.
To prevent aspirating the bee into the user’s mouth, take a fine stainless-steel mesh and attach it with a 1 cm diameter hose clamp to the tube once it has been inserted through the lid into the collection container.
Take another hose clamp and similarly attach it to the bee collection tube once it has been inserted inside the lid. The clamps will keep the tubes from pulling out the lid.
Software and datasets
Tracker software (V4.91) runs on Windows, MacOS, and Linux operating systems and is free. This software is built on the Open Source Physics (OSP) Java framework and is designed for educational use. The software can be downloaded at this website: https://physlets.org/tracker/
We used JMP Pro V16.1.0 to analyze our data. A free 30-day trial of JMP is available. This software can be downloaded from this website: https://www.jmp.com/en_us/home.html
Procedure
Creating colonies
You will be creating pairs of colonies (one control and one experimental) from the same source colony to ensure that each pair of colonies has the same genetic background.
Control and experimental colonies should have very similar levels of food stores.
It is important that all bees (control and experimental) experience the same incubator conditions to account for the potential effects of incubation on bee development and behavior.
Each observation hive consists of two combs (43.5 cm × 23 cm)—one frame of brood and one frame of honey and pollen—and is connected by a 2.2 cm inner diameter and 25 cm long tube through the wall to the outside.
Creating the experimental colony:
Remove combs with late-stage pupae from haphazardly selected large, healthy colonies.
Place the combs inside a nuc box (standard honey bee nucleus box) inside an incubator for 24 h.
Maintain the incubator in a dark environment with a temperature of 34 and a relative humidity of 75%.
Transfer 2,800 one-day-old bees that emerge to a two-frame observation hive with a new egg-laying queen.
Mark 200 of the one-day-old bees with a paint pen. You will track these bees in the experiment.
Ensure that the experimental colonies contain no eggs or brood but only the queen, 2,800 newly emerged bees (of which 200 are marked), and approximately a half comb of pollen and a half comb of honey.
Creating the control colony:
Take the source colony and place one comb with late-stage pupae into a nuc box in the same incubator for 24 h.
Maintain the incubator in a dark environment with a temperature of 34 and a relative humidity of 75%.
Mark 200 of the one-day-old bees that emerge with a paint pen. You will track these bees in the experiment.
At the same time, take 2,600 bees of all ages and, likewise, place them on combs in a nuc box in the same incubator for 24 h under the same conditions.
Transfer the 2,800 one-day-old bees to a two-frame observation hive with a new egg-laying queen.
Repeat as needed to create five experimental colonies and five control colonies that derive from five original source colonies.
These videos illustrate the waggle dances of control bees (Video 1) that exhibit normal levels of error and the waggle dances of experimental bees that did not have the opportunity to follow other dancers before they first danced and therefore show higher levels of error (Video 2). Video files may be downloaded or viewed at the following links. Downloading will usually enable the viewer to see the video at a higher quality.
Video 1. First waggle dance of a bee from a control colony that was able to follow other dancers before performing its first waggle dance (orange and green paint on thorax)
Video 2. First waggle dance of a bee from an experimental colony that was never able to follow another waggle dancer before performing its first waggle dance (green and purple paint on thorax)
Monitoring first waggle dances with no prior experience
Observe all colonies every day during daylight hours until the first bees fly out to forage.
Train these foragers to a 55% (w/v) sucrose feeder placed 150 m from the colony.
Choose locations such that a direct flight path from the colony to the feeder results in bees from each colony experiencing similar levels of optic flow (see this example of our field site: https://map.bmcx.com/ciba__map/).
Ensure that feeders are placed on a grass field with similar local landmarks (similar buildings and trees) at each feeder site.
Create feeders consisting of a 70 mL vial (8 cm high) inverted over a circular plastic disk with 18 feeding grooves through which the sucrose can flow.
Fill the vial with sucrose solution, then invert on the plastic disk and place over a piece of colored paper to help bees return to the feeder once they have learned its location (Figure 1).
Figure 1. Honey bees foraging for 55% (w/v) sucrose solution on a grooved plate feeder
Ensure that no more than 30 foragers visit the feeder at a time and remove bees with an aspirator as needed to reduce crowding.
To train bees, place a glass vial at the entrance of the nest to trap the bees flying out and bring them to a feeder 150 m away, where they are released and begin to imbibe sucrose.
Individually mark the bees with different paint pen colors.
Allow bees to visit the feeder a few times until they waggle dance.
Continuously observe all visits of every bee to the feeder.
Record the first five waggle dance performances of five different bees per colony using a high-definition video camera (HDR-PJ790, Sony).
Illuminate observation colonies with natural light from a window.
Measure the time it takes for foragers to fly back from the feeder to the nest immediately before they waggle dance (defined as return times, with measurements coordinated with the hive and feeder observers via two-way radios).
Revisiting waggle dancing after 20 d of experience
Twenty days later, retrain the marked foragers whose first dances were observed to the same 150 m feeder locations and record their waggle dances to determine if their dancing has changed. At this point, the workers should be, on average, 29 and 30 days old in experimental and control colonies, respectively. See Table 1 for a detailed description of bee type names.
Table 1. Definitions of the different colonies and bee types. The rationale explains the main reason for observing the dances at each stage.
Name Definition Rationale
Experimental colony A single cohort colony created from bees that are all one day old. Experimental colony
E1 naive first dances (E1FD naive) The first waggle dances performed by bees (9.0 ± 2.0 days old, mean ±1 standard deviation) in an experimental colony. They are naive because these bees could not follow other dances before performing their first dance. What do the first dances of foragers that have never danced before or followed other dances look like?
E2 older dancers (E2OD) The waggle dances of E1 bees when they are 29 days old and have more experience following other dancers and dancing. Do the dances of E1 bees change when they are 20 days older and have had experience dancing and following other dances?
Control colony A multi-cohort colony consisting of bees of all ages. Control colony
C1 first dances (C1FD)
The first waggle dances performed by bees (9.9 ± 1.0 days old) in a control colony (control for E1). What do the first dances of foragers that have never danced before but were able to follow other dancers look like?
C2 older dancers (C2OD)
The waggle dances of C1 bees when they are 30 days old (control for E2). Do the dances of C1 bees change when they are 20 days older and have had additional experience dancing and following other dances?
Video-record these waggle dancers and measure their return times (see above).
Hypothesize that E2Older Dancers dances, due to experience, have increased precision and orderliness above that of E1First Dances naive dances.
In the 20 days between E1First Dances naive and E2Older Dancers dances, the feeder is not available.
Note that E2Older Dancers can only follow other dancers of the same age with similar experience levels. C2Older Dancers can follow dancers of different ages with different experience levels.
Measuring the waggle dance
To analyze the waggle dance, use Tracker software (V4.91).
Exclude the first waggle run for each dance of every bee and analyze the subsequent six waggle runs (Couvillon et al., 2012).
Define a dance as a series of consecutive waggle runs and return phases during one visit of a forager inside the nest.
For each dance (Figure 2), measure:
The waggle run angle relative to gravity.
The waggle dance divergence angle (the maximum difference between waggle angles during six waggle runs).
The waggle run duration, defined from when the bee begins to turn and starts to waggle her abdomen to when she stops waggling her abdomen and moves into another turn.
The waggle duration error (the duration difference between longest and shortest waggle run within a dance).
Waggle duration variance (coefficient of variance of the waggle durations).
The number of waggles per waggle run (each waggle is defined as one complete movement of the abdominal tip from right to left to right).
Variance in the number of waggles per waggle run (coefficient of variance of the number of waggles per waggle run per dance).
Return phase duration.
Return phase duration variance (coefficient of variance).
Total number of waggle runs within a dance.
The number of dance followers (each follower defined as a bee following a waggle dancer for ≥ 5 s) per waggle dance performance.
To measure the start and stop of a waggle run, respectively define them by measuring the start and stop of dancer wing oscillations from the video (recorded at 60 fps).
Measure the rate of return phase non-alternations (dance disorder) and define it as the number of non-alternating return phases (Tan et al., 2012).
i. Count the pattern of consecutive return phases and calculate the disorder proportion by dividing the total number of return phase non-alternations by the total number of waggle runs within a dance.
ii. After a dancer makes a waggle phase, she can make a return phase by turning either to her left (L) or her right (R). Dancers often alternate these return phases (i.e., L-R-L-R). A non-alternation return phase occurs when she makes two consecutive return phases in the same direction. For example, the pattern L-L-L-L would be counted as three return phase non-alternations, since we compare each return phase direction with the prior one.
For additional details on how to measure waggle dances, we recommend these excellent papers: Griffin et al., 2012; Preece and Beekman, 2014; Wario et al., 2015; Schürch et al., 2019.
Please see Video 1 and Video 2 for examples of waggle dances that show different levels of error.
Figure 2. Diagram illustrating waggle dance measurements. The diagram on the left shows the location of the food source relative to the sun and the nest. This arrangement results in the waggle dance diagram shown in the center. In this diagram, the angle (α) of the waggle run is the angle between the waggle direction and the vertical dashed blue line (defined by gravity on a vertical comb). This waggle angle communicates the direction of the food source from the colony relative to the sun’s azimuth at the time of the dance. For example, if a food source is in the direction of the sun, the waggle run points straight up. If a food source is in the direction opposite the sun, the waggle run points straight down. In this example, α = 45° to the left of the vertical line, and thus the food source is located 45° to the left of the sun’s azimuth. The waggle run consists of waggling motions of the abdomen, made as the bee traverses a line crossing three points: A, B, and C. The waggle duration is measured from point A to point C. Each waggle is defined by the abdominal tip moving from points a-b-c. (one peak-to-peak cycle). The return phase duration is measured as the time it takes a bee to return from a completed waggle run, and is shown by the curve C-D-A or the curve C-E-A. In a fully ordered waggle dance, the dancer typically turns in the opposite direction on each repetition. For example, if the dancer traverses C-D-A, on the next return phase it will likely follow the path C-E-A. A photo of a waggle dancing bee (w) surrounded by dance followers (f) is shown for context on the right.
Data analysis
To analyze the data, use JMP Pro V16.1.0. You may also use different statistical software, but it is important to use repeated-measures models since you will be comparing the behavior of the same bees when they are younger and then older.
For testing the differences in age of first foraging between experimental and control colonies, use ANOVA with colony type as the independent variable and age of first foraging as the dependent variable.
For the measurements shown in Table 1, use Repeated Measures Mixed Models (REML algorithm) with colony type, bee ID, time point, the interaction of colony type × time point, and colony as random effects.
As needed, log-transform the waggle durations, waggle duration range errors, and the number of waggle runs based on the inspection of model residuals.
To test for differences between the treatment groups within a dance on variance in waggle durations, the number of waggles per waggle run, and return phase durations, calculate the coefficient of variation (CV = standard deviation/mean), and run the models with these coefficients of variation. Make all corrected pairwise comparisons using Tukey HSD tests.
To test for correlations between divergence angles and disorder proportions per dance, use linear regressions, one per bee type (E1First Dances naive, E2Older Dancers, C1First Dances, and C2Older Dancers, Table 1).
Validation of protocol
Data obtained by using this protocol can be found here: DOI: 10.5281/zenodo.7301648. The number of replicates recommended is shown in the datasheet provided.
Recommended statistical analyses are shown above.
These results were published in Science (2023), DOI: doi/10.1126/science.ade1702.
General notes and troubleshooting
Troubleshooting
Problem 1: It can be difficult to train bees to feeders some times of the year due to varying natural food availability.
Solution: Employ a classic, albeit slower, training technique by positioning the feeder in contact with the colony entrance. Note that this training method is not ideal for this experiment since we want to make sure to capture the first waggle dance of a bee. Thus, you will need to use only a low-concentration sugar solution that will still attract bees but for which they will not waggle dance.
Gradually increase the gap between the feeder and nest entrance in increments, first by moving it only 1 cm at a time and later by meters once bees commence flying to the feeder (Seeley, 1995).
Ensure all trained bees return before relocating the feeder.
If bees fail to return, revert to the feeder's previous position, and decrease the relocation distance.
Maintain a record of bees visiting the feeder using a census sheet each 15 min or at shorter regular time intervals, as determined by the rate at which you are moving the feeder. Recording bee visitation helps to ensure that your bees are continuing to visit as you move the feeder.
Once bees are at the final location, increase the sugar solution concentration to elicit waggle dancing.
Problem 2: Bees do not consistently perform waggle dances.
Solution: Select a period of relative natural food scarcity.
Increase sucrose concentration if consistent waggle dancing remains unobserved.
Utilize a saturated sucrose solution (2.5 M) if needed.
If waggle dances persistently fail to materialize, consider rescheduling experiments for a different time of the year.
Problem 3: During periods of food dearth, non-focal colonies discover the feeder and compete with trained bees.
Solution: Implement the following preventative measures:
Rigorously monitor training. Mark each bee that lands on the feeder and verify its return to the focal colony. If the marked bee is absent from the focal colony, capture it immediately once it returns to the feederwith an aspirator and freeze it at day's end upon its subsequent return to the feeder. This minimizes bee sacrifice by curbing recruitment from non-focal colonies.
To freeze bees, it is simplest to have multiple aspirators.
Place the aspirator with the captured bees inside a freezer at -20 °C for at least 12 h to sacrifice them.
Refrain from training bees at previously used locations visited by non-focal bees.
Minimize sucrose concentration to limit recruitment, as higher concentrations increase recruitment. Employing the minimum concentration necessary for trained foragers to revisit the feeder mitigates the risk of massive recruitment by non-focal colonies and regulates focal colony forager numbers. Once you have established the feeder at the correct final location, you can then use a higher sucrose concentration to elicit waggle dancing by the bees.
Recognize that the observation colony has fewer combs than standard and feral colonies, making it easier for non-focal colonies to overtake the feeder.
If feasible, conduct experiments in areas with fewer competing honey bee colonies.
Acknowledgments
This work was supported by the CAS Key Laboratory of Tropical Forest Ecology, Xishuangbanna Tropical Botanical Garden, Chinese Academy of Sciences. Additional funding was provided by the CAS 135 program (No. 2017XTBG-T01), the National Natural Science Foundation of China (No. 31770420) to K. Tan.
This protocol was derived from the original work of Dong et al. (2023).
Competing interests
The authors declare no competing interests.
Ethical considerations
We used honey bees, Apis mellifera, which are invertebrates that are not endangered or protected and do not require Institutional Animal Care and Use Committee protocol authorization.
References
Couvillon, M. J., Riddell Pearce, F. C., Harris-Jones, E. L., Kuepfer, A. M., Mackenzie-Smith, S. J., Rozario, L. A., Schürch, R. and Ratnieks, F. L. W. (2012). Intra-dance variation among waggle runs and the design of efficient protocols for honey bee dance decoding. Biol. Open 1(5): 467–472.
Dong, S., Lin, T., Nieh, J. C. and Tan, K. (2023). Social signal learning of the waggle dance in honey bees. Science 379(6636): 1015–1018.
Griffin, S. R., Smith, M. L. and Seeley, T. D. (2012). Do honeybees use the directional information in round dances to find nearby food sources? Anim. Behav. 83(6): 1319–1324.
Leadbeater, E. and Chittka, L. (2007). Social Learning in Insects — From Miniature Brains to Consensus Building. Curr. Biol. 17(16): R703–R713.
Preece, K. and Beekman, M. (2014). Honeybee waggle dance error: adaption or constraint? Unravelling the complex dance language of honeybees. Anim. Behav. 94: 19–26.
Schürch, R., Zwirner, K., Yambrick, B. J., Pirault, T., Wilson, J. M. and Couvillon, M. J. (2019). Dismantling Babel: creation of a universal calibration for honey bee waggle dance decoding. Anim. Behav. 150: 139–145.
Seeley, T. D. (1995). The Wisdom of the Hive. Harvard University Press.
Tan, K., Wang, Z., Yang, M., Fuchs, S., Luo, L., Zhang, Z., Li, H., Zhuang, D., Yang, S., Tautz, J., et al. (2012). Asian hive bees, Apis cerana, modulate dance communication in response to nectar toxicity and demand. Anim. Behav. 84(6): 1589–1594.
von Frisch, K. (1967). The dance language and orientation of bees. 2nd printing, 1993 Edition. Belknap Press.
Wario, F., Wild, B., Couvillon, M. J., Rojas, R. and Landgraf, T. (2015). Automatic methods for long-term tracking and the detection and decoding of communication dances in honeybees. Front. Ecol. Evol. 3: e00103.
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Determination of Poly(3-hydroxybutyrate) Content in Cyanobacterium Synechocystis sp. PCC 6803 Using Acid Hydrolysis Followed by High-performance Liquid Chromatography
JK Janine Kaewbai-ngam
Aran Incharoensakdi
TM Tanakarn Monshupanee
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4790 Views: 578
Reviewed by: Dennis J NürnbergAmberley D. Stephens Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Plant & Cell Physiology Sep 2022
Abstract
Various photoautotrophic cyanobacteria accumulate intracellular poly(3-hydroxybutyrate) (PHB) granules. This protocol can be used for determining the PHB contents of the cells as % PHB weight per dry cell weight using acid hydrolysis followed by high-performance liquid chromatography (HPLC). This HPLC analysis is rapid, with a running time of approximately 5 min per sample. The technique can accurately determine PHB concentrations in the range of 2–1,000 μg/mL PHB. However, this technique is not applicable for determining the contents of poly(3-hydroxybutyrate-co-3-hydroxyvalerate) in cyanobacteria.
Keywords: Cyanobacteria Synechocystis Poly(3-hydroxybutyrate) PHB HPLC Nitrogen deprivation
Background
Poly(3-hydroxybutyrate) (PHB) is a common biopolymer accumulated in various heterotrophic bacteria and autophototrophic cyanobacteria as a storage for carbon and energy (Amadu et al., 2021; Yashavanth et al., 2021). PHB can be used as biodegradable plastic and scaffold for tissue engineering (Bhati et al., 2010; Monshupanee et al., 2016; Tarawat et al., 2020). In the well-studied cyanobacterium Synechocystis sp. PCC 6803 (hereafter, Synechocystis), cells have low PHB contents 0.4%–5% under a normal growth condition but, under nitrogen or phosphorus deprivation, PHB accumulation is significantly increased, up to 5%–13% w/w dry weight (Wu et al., 2001; Panda and Mallick, 2007; Monshupanee and Incharoensakdi, 2014; Koch et al., 2019). In Synechocystis, PHB biosynthesis is catalyzed by three enzymes (Figure 1). First, two acetyl-CoA are combined into acetoacetyl-CoA by 3-ketothiolase (PhaA). Second, acetoacetyl-CoA is reduced to 3-hydroxybutyryl-CoA by acetoacetyl-CoA reductase (PhaB). Finally, PHA synthase (PhaEC) polymerizes 3-hydroxybutyryl-CoA to form PHB (Taroncher-Oldenburg et al., 2000).
Figure 1. Poly(3-hydroxybutyrate) (PHB) biosynthesis in Synechocystis according to Taroncher-Oldenburg et al. (2000)
Various cyanobacteria accumulate PHB in the absence of essential nutrients including nitrogen or phosphorus (Drosg et al., 2015; Koller and Maršálek, 2015; Kaewbai-ngam et al., 2016). The previous reports on Synechocystis suggested that PHB serves as a carbon and energy reserve for growth recovery under nitrogen repletion (Koch et al., 2019; Kaewbai-ngam et al., 2022).
The common method for quantifying PHB content in microbes is the depolymerization of PHB by methanolysis in acid, with subsequent analysis by gas chromatography (GC) (Juengert et al., 2018). Another alternative method is the alkaline lysis of PHB, followed by the enzymatic assay method using 3-hydroxybutyrate dehydrogenase (Zilliges and Damrow, 2017). However, the GC method is time consuming, and the enzymatic assay requires enzyme preparation, which is costly. Here, we describe a rapid quantification of PHB content in cyanobacterial cells using acid hydrolysis followed by high-performance liquid chromatography (HPLC). This technique takes only 5 min of HPLC operation per sample and can be used to determine PHB contents in 135 evolutionarily diverged cyanobacterial species (Kaewbai-ngam et al., 2016). Poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) is the co-polymer, which comprises 3-hydroxybutyrate (HB) and 3-hydroxyvalerate (HV) monomers (Verlinden et al., 2007; Balaji et al., 2013). PHBV changes material properties upon altering mole ratio (mol %) of the two monomers: 3-hydroxybutyrate (HB) and 3-hydroxyvalerate (HV) (Tarawat et al., 2020). We are reporting here a detailed protocol of PHB content determination that summarizes the steps for converting PHB to crotonic acid by acid hydrolysis, followed by the determination of crotonic acid contents using HPLC. We also show that this protocol cannot be used to determine PHBV content.
Materials and reagents
Organism: wildtype Synechocystis sp. PCC 6803 (Pasteur Culture Collection, France)
Pipette tips 5,000 μL (Biohit, catalog number: 780 300)
Pipette tips 1,000 μL (Labcon, catalog number: 1046-800-000-9)
Pipette tips 200 μL (Labcon, catalog number: 1165-800-000)
Pipette tips 10 μL (Labcon, catalog number: 1161-800-000)
Plastic cuvette for spectrophotometer (Brand, catalog number: 7591 50)
50 mL tubes (Axygen Scientific, catalog number: SCT-50ml-25-S)
10 mL glass tubes with screw cap (Pyrex, catalog number: 9825)
2 mL HPLC glass vials with septum cap (Thermo Scientific, catalog number: CHSV9-10P)
1.5 mL reaction tubes (Labcon, catalog number: 3016-870-000)
3 mL disposable syringes (Nipro, catalog number: 20B20/FEB2020)
Sterile syringe polypropylene filter, 0.22 μm (Filtep-bio, catalog number: 20220516002)
Polypropylene membrane, 0.45 μm (Filtrex, catalog number: FTM-PP47045-010)
Poly(3-hydroxybutyrate) (PHB) (Sigma-Aldrich, catalog number: 363502)
Poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) (Sigma-Aldrich, catalog number: 403121)
Crotonic acid (Sigma-Aldrich, catalog number: 113018)
Adipic acid (Fluka Analytical, catalog number: 09582)
Distilled water
Acetonitrile, HPLC grade (Honeywell, catalog number: AHO15-4A)
Methanol, HPLC grade (VWR chemicals, catalog number: 67-56-1)
Absolute ethanol (Merck, catalog number: 100983)
Citric acid (BDH, catalog number: 08D150022)
Sodium hydroxide (Merck, catalog number: 1310-73-2)
Potassium dihydrogen phosphate (KH2PO4) (Merck, catalog number: 104873)
Ortho-phosphoric acid (H3PO4) (Sigma-Aldrich, catalog number: 7664-38-2)
Calcium chloride dihydrate (CaCl2·2H2O) (Merck, catalog number: 102382)
Ethylenediamine tetraacetic acid disodium salt dihydrate (Na2EDTA·2H2O) (Merck, catalog number: 108418)
Iron (III) chloride (FeCl3) (Ajax Finechem, catalog number: AF608291)
Magnesium sulfate heptahydrate (MgSO4·7H2O) (Merck, catalog number: 105886)
Sodium carbonate (Na2CO3) (Carlo Erba Reagenti, catalog number: 7783-20-2)
Sodium nitrate (NaNO3) (Ajax Finechem, catalog number: 1502523945)
(4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)-NaOH (HEPES-NaOH) (OmniPur, catalog number: 7365-45-9)
Boric acid (H3BO3) (Scharlau, catalog number: 10043-35-3)
Manganese (II) chloride tetrahydrate (MnCl2·4H2O) (Kemaus, catalog number: 13446-34-9)
Zinc sulfate heptahydrate (ZnSO4·7H2O) (Ajax Finechem, catalog number: 0707330)
Sodium molybdate dihydrate (Na2MoO4·2H2O) (Kemaus, catalog number: 10102-40-6)
Copper (II) sulfate pentahydrate (CuSO4·5H2O) (BDH, catalog number: 100915R)
Cobalt (II) nitrate hexahydrate (Co(NO3)2·6H2O) (Carlo Erba Reagenti, catalog number: 10026-22-9)
Dipotassium hydrogen phosphate (K2HPO4) (Carlo Erba Reagenti, catalog number: 7758-11-4)
Sulfuric acid (H2SO4) (Merck, catalog number: 100731)
BG11 media (see Recipes)
Standard sulfuric acid solution (see Recipes)
Standard adipic acid solution (see Recipes)
Standard crotonic acid solution (see Recipes)
Premix solution used in Table 1 (see Recipes)
Mobile phase for HPLC analysis (see Recipes)
Equipment
Micro pipettes (Gilson: 20–1,000 μL)
Micro pipettes (Boeco, model: SA series ADJ 500–5,000 μL)
Glass pipettes (Qualicolor: 1–10 mL) and rubber pipette bulbs
Wide-neck 250 mL Erlenmeyer glass flasks (Duran)
Laboratory glass bottles (Duran)
Personal protective equipment (PPE)
Note: This should be worn at all times when working with concentrated acids.
Safety glasses
Lab coat
Gloves, purple nitrile (Kimtech, catalog number: SM0182ZZZ_37AX)
Autoclave (Hirayama manufacturing corporation, model: HA-30)
Laminar flow bench (Boss tech, model: HBV120)
Fume hood (HYSC LAB, model: FH-120)
Spectrophotometer (Jenway, model: UV/VIS 6400)
Culture shaker [Sac Science, model: Sling shaker (120–150 rpm), 3 levels]
Centrifuge (Hettich zentrifugen, model: Mikro 220/220 R)
Microcentrifuge (Eppendorf, model: 5417C)
Analytical balance (Mettler Toledo, model: PJ360)
Water bath (Hangzhou Bioer technology, model: N series)
Oven (Contherm, model: 2050)
Vortex shaker (Scientific industries, model: K-550-GE)
pH meter (Mettler Toledo, model: FiveEasy F20)
Filtration equipment (Pyrex, model: 33980-1L)
HPLC machine (Shimadzu, model: Shimadzu-02-1060)
HPLC UV/Vis detector (Shimadzu, model: SPD-10A)
HPLC pump (Shimadzu, model: LC-10AD)
HPLC auto-injector (Shimadzu, model: SIL-10AD VP)
HPLC column oven (Shimadzu, model: CTO-10A VP)
HPLC column (GL Sciences, model: InertSustain C18, 4.6 × 150 mm, 5 μm)
Conventional guard column (GL Sciences, model: InertSustain C18, 4.0 mm × 10 mm, 5 μm)
Software
Shimadzu HPLC LGE system, Japan
Excel, Microsoft, USA
Procedure
The procedure for determining the PHB content of Synechocystis cells is described in Figure 2. It consists of four steps, including (A) the cultivation of cyanobacteria, (B) acid hydrolysis to convert PHB into crotonic acid, (C) HPLC sample preparation, and (D) determination of PHB content using HPLC.
Two-step Synechocystis cultivation: grow cells under standard growth conditions to increase cell biomass and then transfer cells to nitrogen-deprived media to stimulate PHB accumulation
Inoculate BG11 media (Rippka et al., 1979) (see Recipes) with Synechocystis in Erlenmeyer flasks using the initial cell density of OD730nm = 0.2 and a total volume of 100 mL. Then, cultivate cells at 28 °C under continuous white light at 50 μmol m-2·s-1 with an atmospheric CO2 supply via culture shaking at 160 rpm. Cultivate cells for 4–5 days to achieve the cell density of OD730nm = 0.6–0.8.
Harvest cells by centrifugation at 4,000× g for 10 min at room temperature and wash cells twice with BG110 media (nitrogen-deprived medium) (see Recipes), followed by using the aforementioned centrifugation step. Remove supernatant medium in each washing step.
Perform the second cultivation step by inoculating the harvested cells in BG110 medium, using the initial OD730nm = 0.4 and the total volume of 100 mL. Next, cultivate cells for 7–14 days to induce PHB accumulation at 28 °C under continuous white light at 50 μmol m-2·s-1 with an atmospheric CO2 supply via culture shaking at 160 rpm.
Harvest cells from 40 mL of cell culture by centrifugation at 4,000× g for 10 min at room temperature. Next, resuspend fresh biomass in BG11 (total volume not exceeding 1.4 mL) and transfer the resuspended cell sample to the 1.5 mL tube. Then, centrifuge the tube, remove the liquid medium, and immediately dry cells at 60 °C in a hot oven until the weight of the sample stabilizes.
Note: The wet cell pellets can be stored at -20 °C or dried at 60 °C immediately before being used.
Figure 2. Overview of procedures for determining poly(3-hydroxybutyrate) (PHB) content of Synechocystis. Cells are cultivated, harvested, and dried. Next, dry cells are hydrolyzed by acid to convert PHB into crotonic acid. Then, crotonic acid content is quantified using HPLC. Details of each step are described in the procedure.
Acid hydrolysis to convert PHB into crotonic acid
Note: This step uses a strong acid, thus it must be performed under the fume hood and while wearing personal protective equipment.
Weigh approximately 5–10 mg of dry cells or 5 mg of the standard commercial PHB into 10 mL screw-capped glass tubes.
Add 1 mL of 18.4 M sulfuric acid by using a glass pipette to dry cells; boil at 100 °C for 1 h in a hot water bath to break cells and hydrolyze PHB into crotonic acid (Figure 3). The typical percentage of mass conversion of PHB to crotonic acid is in the range of 83% ± 7% (w/w) (Singhon et al., 2021; Kaewbai-ngam et al., 2022).
Figure 3. Depolymerization of poly(3-hydroxybutyrate) (PHB) by sulfuric acid to form crotonic acid. The reaction information is according to Koch et al. (2020).
Cool sample until reaching room temperature; then, dilute the obtained total hydrolysate (1 mL) by adding 4 mL of distilled water and mix thoroughly by vortexing. The total volume of sample is 5 mL.
Note: For safety reasons, assure that the hot samples after acid hydrolysis are cooled down before proceeding to the next step.
For the commercial PHB sample (5 mg) that has been hydrolyzed and diluted to the total 5 mL in the step B3, make a 1:2, 1:5, 1:10, 1:50, 1:100, and 1:500 dilution in the total 5 mL volume to generate the standard PHB samples corresponding to the total PHB of 2.5, 1, 0.5, 0.1, 0.05, 0.01 mg of PHB in the 5 mL total volume.
HPLC sample preparation
Prepare HPLC samples according to Table 1 and mix thoroughly by vortexing.
Filter HPLC sample using a syringe and the polypropylene filter (0.22 μm); then, transfer the obtained filtrate to HPLC glass vials.
Table 1. HPLC sample preparation. N.A.: no addition
HPLC sample name Component 1 Component 2 Total sample volume
1. Hydrolysate from Synechocystis cells 200 μL of cell hydrolysate from step B3 800 μL of the premix solution (see Recipes) 1,000 μL
2. Hydrolysate from commercial PHB (PHB standard) 200 μL of PHB hydrolysate and its six diluted samples from step B4 800 μL of the premix solution (see Recipes)
3. Standard sulfuric acid (negative control) 1,000 μL of standard sulfuric acid (see Recipe 2) N.A.
4. Standard adipic acid (internal standard) 1,000 μL of standard adipic acid (see Recipe 3) N.A.
5. Standard crotonic acid (positive control) 1,000 μL of standard crotonic acid (see Recipe 4) N.A.
Note: The acid hydrolysis of biomass and HPLC analysis should be completely conducted within two days. The hydrolyzed samples can be stored at 4 °C for maximum of two days.
Determination of PHB content using HPLC
Set up the HPLC operation as shown below:
• InertSustain Carbon-18 column
• UV detection at 210 nm
• 40 °C oven temperature
• 10 μL injection volume
• 1 mL/min flow rate
• 5 min running time per one sample
• Mobile phase: [30:70 (% v/v) acetonitrile:10 mM KH2PO4 buffer pH 2.3]
Note: There is no washing step between runs because no gradient of the mobile phase is used.
Data analysis
By using the mobile phase [30:70 (% v/v) acetonitrile:10 mM KH2PO4 buffer pH 2.3], the obtained retention times of the samples are shown in Table 2. Note that these retention times are specific for the HPLC column, the HPLC system, and the mobile phase used in this study.
Table 2. Obtained retention time of the HPLC standards and cell sample using 30% (v/v) acetonitrile in 10 mM KH2PO4 buffer as the mobile phase
Sample Obtained retention times (min) Chromatogram peak
1. Hydrolysate from commercial PHB 3.02 (crotonic acid) and 2.25 (adipic acid internal standard) Figure 5A
2. Hydrolysate from Synechocystis 3.09 (crotonic acid) and 2.29 (adipic acid internal standard)
3. Standard sulfuric acid (negative control) 1.59–1.65 Figure 5B
4. Standard crotonic acid (positive control) 3.00–3.10
5. Standard adipic acid (internal standard) 2.20–2.30
Calculation of PHB content
Generate a standard curve of the six amounts (0–5 mg, see step B4) of the standard PHB sample.
i. Collect HPLC peak areas of crotonic acid and adipic acid from each amount of PHB standard.
ii. Calculate the ratio between the peak area of crotonic and the peak area of adipic acid. All peak areas used in this study were defined by the HPLC software.
iii. Plot the graph: the ratio between the peak area of crotonic and the peak area of adipic acid on the y-axis against the amount of PHB (0–5 mg) on the x-axis.
iv. Determine the linear equation (y = a × x + b, where b = 0). The linear regression coefficient should be in the range of 0.99 ≤ R2 ≤ 1 (Figure 4).
Figure 4. Standard curve for the determination of poly(3-hydroxybutyrate) (PHB) weight. The linear correlation was obtained under standard PHB = 0–5 mg, but not at 8 and 10 mg in the total volume of 5 mL sample. Therefore, the maximum standard PHB used for the analysis was 5 mg in 5 mL.
Calculate the weight of PHB in the cell sample using the formula:
x = y/a
x: Weight of PHB (mg) in cell sample
y: The ratio between the peak area of crotonic: the peak area of adipic acid derived from cell sample
a: The value obtained from the formula (y = a × x) in step B1
Calculate PHB content in Synechocystis cells as % w/w dry weight (DW) using the formula:
PHB content (% w/w DW) = [x (mg)/cell weight (mg)] × 100
x: Weight of PHB (mg) in cell sample obtained in step B2
Samples from at least three independent cultures must be used.
Increased acetonitrile concentrations in the mobile phase significantly reduced the retention times of all target samples (adipic acid and crotonic acid) (Figure 5A). For example, the HPLC using the mobile phase without acetonitrile showed the retention time of crotonic acid at 34.7 min, while HPLC using the mobile phase containing 30% (v/v) acetonitrile significantly reduced the retention time of crotonic acid to only 3.0 min (Figure 5A). Therefore, the recommended percentage of acetonitrile in mobile phase is 30% (v/v), which was used throughout this experiment, and the HPLC running time per sample is only 5 min (Figure 5A).
Figure 5. HPLC chromatograms of all samples used in this study. (A) The chromatograms of the commercial poly(3-hydroxybutyrate) (PHB) and the cell sample in the mobile phase containing 0, 15, 30, and 45% (v/v) acetonitrile. (B) The chromatograms of the indicated samples in the mobile phase containing 30% (v/v) acetonitrile. PHBV: commercial poly(3-hydroxybutyrate-co-3-hydroxyvalerate).
The lower limit of the sensitivity of this protocol is a PHB content of 0.01 mg per 5 mL of total volume, corresponding to a PHB concentration of 2 μg/mL (Figure 4). The recommended maximal amount of PHB to be analyzed by this technique is 5 mg per 5 mL of total volume, corresponding to a PHB concentration of 1 mg/mL (Figure 4). If the PHB concentration is higher than 1 mg/mL, this sample must be diluted after acid hydrolysis prior to HPLC analysis.
The analysis of the acid hydrolysis followed by HPLC of the commercial poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) showed only the peak of crotonic acid (the hydrolyzed product of 3-hydroxybutyrate), and no peak of the hydrolyzed product of 3-hydroxyvalerate (Figure 5B) as detected by 210 nm UV absorption. Thus, the hydrolyzed product of 3-hydroxyvalerate may not absorb the light at 210 nm. There is no current report on the chemical identity of the hydrolyzed product of 3-hydroxyvalerate. Therefore, the acid hydrolysis followed by HPLC used in this study cannot be used to determine the content of P(HB-co-HV) in cyanobacteria. It is noted that successful analyses of P(HB-co-HV) from cyanobacterial cells using methanolysis in acid followed by gas chromatography have been reported previously (Bhati and Mallick, 2015; Taepucharoen et al., 2017; Tarawat et al., 2020).
Recipes
BG11 media (autoclaved)
The medium composition is according to Rippka et al. (1979); (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)-NaOH (HEPES-NaOH) was added to the medium.
Number
Trace element component
Final concentration [mM]
BG11 media
BG110 media
1
NaNO3
17.60
0
2
K2HPO4
0.23
0.23
3
MgSO4·7H2O
0.30
0.30
4
CaCl2·2H2O
0.24
0.24
5
FeCl3
0.021
0.021
6
Na2EDTA·2H2O
0.0027
0.0027
7
Na2CO3
0.19
0.19
8
H3BO3
46.00
46.00
9
MnCl2·4H2O
9.00
9.00
10
ZnSO4·7H2O
0.77
0.77
11
Na2MoO4·2H2O
1.60
1.60
12
CuSO4·5H2O
0.30
0.30
13
Co(NO3)2·6H2O
0.17
0.17
14
HEPES-NaOH (pH 7.5)
20.00
20.00
Standard sulfuric acid (negative control) (5 mL)
0.74 M H2SO4
Standard adipic acid (internal standard) (10 mL)
15 mg/mL adipic acid in 0.74 M H2SO4
Standard crotonic acid (10 mL)
10 mg/mL crotonic acid in distilled H2O
Premix solution used in Table 1 (100 mL per 100 samples)
0.375 g of adipic acid in 100 mL of 1.84 M H2SO4
Mobile phase for HPLC analysis (1 L per 100 samples)
1.36 g/L KH2PO4 buffer [adjust to pH 2.3 using H3PO4 and filtrate the solution using the polypropylene membrane (0.45 μm) and 30% (v/v) acetonitrile (HPLC grade)]
Note: Solutions 2–6 must be freshly prepared just before use.
Acknowledgments
This protocol was adapted from the previous protocol described by Schlebusch and Forchhammer (2010). This work was supported by The Royal Golden Jubilee Ph.D. Program (PHD/0011/2560) and National Research Council of Thailand (NRCT): NRCT5-RSA63001-21. The authors declare no conflicts of interest.
Competing interests
We have no competing interests.
References
Amadu, A. A., Qiu, S., Ge, S., Addico, G. N. D., Ameka, G. K., Yu, Z., Xia, W., Abbew, A. W., Shao, D., Champagne, P., et al. (2021). A review of biopolymer (Poly-β-hydroxybutyrate) synthesis in microbes cultivated on wastewater. Sci. Total Environ. 756: 143729.
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Kaewbai-ngam, J., Sukkasam, N., Phoraksa, O., Incharoensakdi, A. and Monshupanee, T. (2022). Production of glycogen, PHB, biohydrogen, NAD(P)H, and proteins in Synechocystis sp. PCC 6803 disrupted in metabolically linked biosynthetic pathway(s). J. Appl. Phycol. 34(4): 1983–1995.
Koch, M., Bruckmoser, J., Scholl, J., Hauf, W., Rieger, B. and Forchhammer, K. (2020). Maximizing PHB content in Synechocystis sp. PCC 6803: a new metabolic engineering strategy based on the regulator PirC. Microb. Cell Fact. 19(1): e1186/s12934-020-01491-1.
Koch, M., Doello, S., Gutekunst, K. and Forchhammer, K. (2019). PHB is Produced from Glycogen Turn-over during Nitrogen Starvation in Synechocystis sp. PCC 6803. Int. J. Mol. Sci. 20(8): 1942.
Koller, M. and Maršálek, L. (2015). Cyanobacterial polyhydroxyalkanoate production: status quo and quo vadis? Curr. Biol. 4(3): 464–480.
Monshupanee, T. and Incharoensakdi, A. (2014). Enhanced accumulation of glycogen, lipids and polyhydroxybutyrate under optimal nutrients and light intensities in the cyanobacterium Synechocystis sp. PCC 6803. J. Appl. Microbiol. 116(4): 830–838.
Monshupanee, T., Nimdach, P. and Incharoensakdi, A. (2016). Two-stage (photoautotrophy and heterotrophy) cultivation enables efficient production of bioplastic poly-3-hydroxybutyrate in auto-sedimenting cyanobacterium. Sci. Rep. 6(1): e1038/srep37121.
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An In-depth Guide to the Ultrastructural Expansion Microscopy (U-ExM) of Chlamydomonas reinhardtii
NK Nikolai Klena
GM Giovanni Maltinti
UB Umut Batman
GP Gaia Pigino
PG Paul Guichard
VH Virginie Hamel
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4792 Views: 2028
Reviewed by: Xin Xu Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Science Jul 2022
Abstract
Expansion microscopy is an innovative method that enables super-resolution imaging of biological materials using a simple confocal microscope. The principle of this method relies on the physical isotropic expansion of a biological specimen cross-linked to a swellable polymer, stained with antibodies, and imaged. Since its first development, several improved versions of expansion microscopy and adaptations for different types of samples have been produced. Here, we show the application of ultrastructure expansion microscopy (U-ExM) to investigate the 3D organization of the green algae Chlamydomonas reinhardtii cellular ultrastructure, with a particular emphasis on the different types of sample fixation that can be used, as well as compatible staining procedures including membranes.
Graphical overview
Keywords: Expansion microscopy Chlamydomonas Flagella Basal body Super-resolution microscopy Chloroplast Membranes Mitochondria Microtubules Cell architecture
Background
Chlamydomonas reinhardtii, a single-celled green algae, is an ideal model organism in the fields of carbon fixation and ciliary-based motility, owing to its large chloroplast and two flagella. As such, electron microscopists have used Chlamydomonas for decades to unravel cellular questions about photosynthetic processes (Salomé and Merchant, 2019), basal body biogenesis and function (Dutcher, 2003), and flagellar motility (Klena and Pigino, 2022). However, the cellular localization of specific proteins of interest is difficult to address by electron microscopy (EM) and, when possible, it requires complex correlative light and EM approaches. Light microscopy and immunofluorescence microscopy are often used to visualize specific protein targets, but the resolution is often insufficient, and the cellular context is indeterminable. Furthermore, utilizing super-resolution techniques such as stochastic optical reconstruction microscopy and stimulated emission depletion requires highly specialized setups and complicated antibody labeling. To circumvent this, we recently utilized super-resolution ultrastructural expansion microscopy (U-ExM) on Chlamydomonas to assess how intraflagellar transport trains (IFT) assemble at the ciliary base (van den Hoek et al., 2022). U-ExM is an approach of expansion microscopy derived from the magnified analysis of the proteome protocol that allows for a physical, volumetric expansion of the entire proteome by a factor of 4 (Ku et al., 2016; Gambarotto et al., 2019). By coupling U-ExM with cryo-fixation (Laporte et al., 2022b), NHS-ester (a reactive dye interacting with amines)-mediated pan-ExM experiments (M’Saad and Bewersdorf, 2020), and BODIPYTM (conjugated dye to label lipids) membrane staining (Liffner and Absalon, 2021), the native ultrastructural environment of the entire organism can be assessed through light microscopy, and the localization of specific proteins of interest can be determined by immunofluorescence. The protocol presented here is dedicated to the expansion of Chlamydomonas cells but it can also be used for other organisms such as human cells (Laporte et al., 2022a), mouse tissue (Mercey et al., 2022), or yeast (Hinterndorfer et al., 2022).
Materials and reagents
Reagents
Chlamydomonas growth media
Tris(hydroxymethyl)aminomethane (TRIS) (Sigma, catalog number: 252859), store at room temperature (RT)
Hutner’s trace elements (Chlamydomonas Resource Center)
Potassium phosphate monobasic (KH2PO4) (Sigma, catalog number: P5655), store at RT
Potassium phosphate dibasic (K2HPO4) (Sigma, catalog number: P3786), store at RT
Acetic acid (Sigma, catalog number: A6283)
TRIS-acetate-phosphate (TAP) medium (see Recipes)
Phosphate buffer II (see Recipes)
Solution A (40×) (see Recipes)
Regular U-ExM protocol
Formaldehyde, 36.5%–38% (FA) (Sigma, catalog number: F8775), store at RT
Acrylamide, 40% (AA) (Sigma, catalog number: A4058), store at 4 °C
Sodium dodecyl sulfate (SDS) (Carl Roth, article number: CN30.3), store at RT
N,N’-methylenebisacrylamide, 2% (BIS) (Sigma, catalog number: M1533), store at 4 °C
Sodium acrylate, 97%–99% (SA) (Sigma, catalog number: 408220), store at -20 °C
Ammonium persulfate (APS) (Thermo Fisher, catalog number: 17874), store at RT
Tetramethylethylenediamine (TEMED) (Thermo Fisher, catalog number: 171919), store at RT
TWEEN 20 (Sigma, catalog number: P1379), store at RT
Bovine serum albumin (BSA) (Sigma, catalog number: A3059), store at 4 °C
Poly-D-lysine, 0.1 mg/mL (PDL) (Gibco, catalog number: A3890401), store at 4 °C
PBS (1× and 10×)
ddH2O
Nuclease-free water (Thermo Fisher Scientific, catalog number: AM9937)
Antibodies (primaries and secondaries)
NHS-ester Atto488 (Sigma, catalog number: BCCJ6663)
NHS-ester 405 (Sigma, catalog number: 250966)
NHS-ester Atto 594 (Sigma, catalog number: 08AA12)
NHS-ester Atto 647 (Sigma, catalog number: BCCD9924)
BODIPY 558/568 (Red) (Thermo Fisher Scientific, catalog number: D3835)
Paraformaldehyde (PFA), 16% (EMS, catalog number: 15710)
Glutaraldehyde (GA), 25% (Sigma, catalog number: G5882)
Methanol (MeOH) (Sigma, catalog number: 34860)
α-tubulin antibody (ABCD Antibodies, catalog number: AA345)
β-tubulin antibody (ABCD Antibodies, catalog number: AA344)
SNAP-tag antibody (New England Biolabs, catalog number: P9310S)
PsbA antibody (Agrisera, AS06 143)
Centrin antibody clone 20H5 (Millipore, catalog number: 04-1624)
Formaldehyde/acrylamide mixture (1.4% FA, 2% AA) (see Recipes)
Sodium acrylate solution (38% w/w) (see Recipes)
Monomer solution, 10 aliquots of 90 μL (see Recipes)
10% TEMED, 10 aliquots of 100 μL (see Recipes)
10% APS, 10 aliquots of 100 μL (see Recipes)
Denaturation buffer, pH 9, 100 mL (see Recipes)
PBS-Tween (0.1% w/v) (1×, 1 L) (see Recipes)
PBS-BSA (2%) (1×, 100 mL) (see Recipes)
Additional reagents for Cryo-ExM protocol
Acetone (99.8% AcroSeal) (Acros Organics, catalog number: 67-64-1)
Ethanol (EtOH, absolute) (Thermo Fisher Scientific, catalog number: 397691000)
Dry ice
Ethane gas bottle (PanGas). It is also possible to use a 37%:63% mixture of ethane:propane (PanGas), which avoids the freezing of the gas during the preparation
Materials
Regular U-ExM protocol
Erlenmeyer flasks
Tweezer Dumont Style 5 with super thin tips (0203-L5-PO) or Negative-Action tweezer Style no. 5 (0203-N5-PO)
12 mm coverslips, high precision no. 1.5H (Marienfeld Laboratory Glassware, catalog number: 47442)
24 mm coverslips, high precision no. 1.5H (Marienfeld Laboratory Glassware, catalog number: 48639)
12-well plates (Thermo Fisher Scientific, catalog number: 150200)
6-well plates (Thermo Fisher Scientific, catalog number: 150239)
P1000 pipette tips
P200 pipette tips
P20 pipette tips
Spatula (Bochem, catalog number: 3101)
Spoon (Bochem, catalog number: 3421)
Razor blade (Carl-Roth, catalog number: CK07.2)
250 mL beaker (VWR, catalog number: 213-1124)
500 mL beaker (VWR, catalog number: 213-1126)
Caliper
Fine-mesh net (such as mosquito netting or insect netting for a garden)
Parafilm (BemisTM, catalog number: PM996)
Scissors
Pasteur pipettes (Alpha Laboratories, catalog number: LW4111)
1.5 mL Eppendorf tubes
35 mm imaging chamber (metallic O-ring 35 mm) (Okolab, catalog number: RA-35-18 2000–06)
Coverslip rack (Thermo Fisher Scientific, catalog number: C14784)
Additional material for Cryo-ExM protocol
Polystyrene box (23 cm × 23 cm × 21 cm)
5 mL Eppendorf tubes (Eppendorf, catalog number: 0030119401)
Tweezer Dumont Style L5 Inox 8 with clamping ring (Electron Microscopy Sciences)
Whatman filter paper, 55 mm (GE Healthcare, catalog number: 1001-055)
Equipment
Regular U-ExM protocol
37 °C incubator or climate-controlled room
Heat block (Cole-ParmerTM, catalog number: SBH130DC)
Inverted fluorescent microscope, such as widefield Leica Thunder, SP8, or Zeiss LSM 980, equipped with a 63× oil objective.
Additional equipment for Cryo-ExM protocol
Cryo-EM manual plunger [both devices used in the preparation of this manuscript were homemade plunging devices; however, commercial manual cryo-plunging devices are usable (https://www.mitegen.com/product/manual-plunge-cooler/), and homemade manual plunging devices can be produced by groups (Comolli et al., 2012)].
Software
Fiji (ImageJ, https://imagej.net/software/fiji/) (Schindelin et al., 2012)
Depending on the brand of microscope used for imaging, dedicated software is required, such as:
LAS X (Leica, https://www.leica-microsystems.com/products/microscope-software/p/leica-las-x-ls/)
Zen Blue (Zeiss, https://www.zeiss.com/microscopy/en/products/software/zeiss-zen.html)
Procedure
Chlamydomonas cell culture and strains
Inoculate Chlamydomonas strain in at least 10 mL of 1× TAP medium and incubate at 23 °C while shaking (Hoober, 1989). Only a few hundred microliters of culture are required for seeding coverslips.
Grow cells until they reach the logarithmic growth phase and the culture turns bright green. This time corresponds to approximately three days.
Note: Unlike yeast (Hinterndorfer et al., 2022), the cell wall of Chlamydomonas does not appear to prevent expansion or induce expansion artifacts (Figure 1). We tested two strains, CC124- (cell wall–positive) and CW15- (cell wall–negative), in U-ExM without fixation, and in both cases the cells expand properly by a factor of 4.
Figure 1. The cell wall of Chlamydomonas reinhardtii does not affect the four-fold expansion factor in ultrastructure expansion microscopy (U-ExM). (A) Representative brightfield image of Chlamydomonas CC124- (cell wall–positive) strain before expansion, displayed as single z plane. Scale bar: 25 μm. (B) Representative widefield image of Chlamydomonas CC124- strain after expansion, stained with pan NHS-ester (far red). The cells are not fixed prior to expansion. Scale bar: 25 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor. (C) Measurements of the average cell area of the entire Chlamydomonas CC124- and CW15- (cell wall–negative) strains, before and after the expansion. The area of expanded cells is divided by the expansion factor for comparison to the non-expanded cell area. (n = 329, 126, 256, and 111 cells, respectively, from two independent experiments. CC124- non-expanded = 40.50 ± 2.21 μm2, expanded = 42.12 ± 0.06 μm2, CW15- non-expanded = 50.10 ± 0.36 μm2 and expanded = 50.69 ± 3.07 μm2; mean ± S.D.) (ns, p > 0.05) (D) Schematic representation of cell area calculation of Chlamydomonas.
Media, stock solution, and expansion microscopy reagent preparation
In the days awaiting Chlamydomonas growth, prepare stock solutions of sodium acrylate, APS and TEMED aliquots, denaturation buffer, PBS-Tween, and PBS-BSA.
Prepare 12 and 24 mm PDL-coated coverslips by pipetting stock PDL onto coverslips until the surface is covered, which corresponds to roughly 50 μL for 12 mm coverslips and 100 μL for 24 mm coverslips. Place coverslips at 37 °C at ambient humidity, for 30 min, and wash 3× in sterile water. Remove excess water and let air dry. Store coverslips at 4 °C until use.
Note: Plasma-cleaning the surface of 24 mm coverslips prior to PDL-coating greatly reduces sample drift during the acquisition. See Procedure section E and Notes.
Chlamydomonas coverslip seeding, fixation, and gel formation (day 0)
When Chlamydomonas cells reach logarithmic growth, remove PDL-coated 12 mm coverslips from 4 °C and bring to RT for approximately 10 min.
Using a P1000 with a cut tip, add 100 μL of Chlamydomonas culture to the center of the PDL-coated 12 mm coverslip. After 5–10 min, the cells will adhere to the bottom of the coverslip (Figure 2).
Figure 2. Tools and visual aids for Chlamydomonas ultrastructure expansion microscopy (U-ExM). A. Spatula, spoon, tweezers, plastic blade, and imaging chamber used in U-ExM. B. Chlamydomonas seeded on a 12 mm coverslip prior to fixation. The tweezer used to hold the coverslip here is a Tweezer Negative-Action Style no. 5. C. Gelation chamber on ice used to form gels. The blue ring is a dampened paper towel. Inset: 45° drop of coverslip onto gel droplet. D. Representative gel after the first round of expansion. E. Representative gel prior to expansion.
Proceed to fixation steps.
Multiple options for fixation are possible and should be chosen depending on the cellular compartment to be imaged. Fixation choices include cold methanol, paraformaldehyde + glutaraldehyde (PFA + GA), cryo-fixing by plunge freezing (Laporte et al., 2022b), and our recently optimized cryo-fixation protocol for ideal membrane preservation, by using 0.1% PFA + 0.02% GA in acetone during the substitution procedure (Louvel et al., 2022). Methanol fixation is well known to preserve the cytoskeletal network but generates a large amount of cytoplasmic and membrane clearance of other cellular components due to protein precipitation and extraction. On the other hand, PFA + GA is a gold standard fixation method to preserve membrane-bound organelles, such as mitochondria and the Golgi apparatus, but results in a poor expansion of some elements of the cytoskeleton. In particular, we observed that the centriole does not expand well if the fixation contains too much GA, whereas PFA or FA do not seem to interfere with the expansion factor (Gambarotto et al., 2019). Finally, cryo-fixation followed by freeze-substitution, recently adapted for immunofluorescence and expansion microscopy techniques, is the method of choice to best preserve cells in their native state (Laporte et al., 2022b). This method coupled with U-ExM has been shown to preserve the cytoskeleton as well as membranous structures. To further improve membrane preservation, addition of PFA + GA in the freeze-substitution step was recently adapted (Louvel et al., 2022) (Figure 3). It is to be noted that cells can also remain unfixed, especially to visualize the basal bodies and flagella, which remain intact under this condition, as well as some fibrous structures as visualized by Centrin staining (Figure 3A–3C) (van den Hoek et al., 2022). However, some cellular structures might be affected, such as dynamic microtubules (Gambarotto et al., 2021), membranous organelles, or the pyrenoid (Figure 3B). To best preserve the organization of these organelles, cryo-fixation is then the approach of choice (Laporte et al., 2022b; Louvel et al., 2022) (Figure 3D).
Figure 3. Comparison of ultrastructure expansion microscopy (U-ExM) without fixation and cryo-ExM. A. Chlamydomonas CW15- cells were expanded following the U-ExM protocol. No fixation was performed prior to expansion. Gels were stained with Centrin (cyan), Tubulin (magenta), and NHS-ester (gray). Top panel shows the whole cell. Note that the Centrin signal was overexposed to unveil the Centrin localization around the nucleus. Middle panel shows the inner basal body and striated fiber localization of Centrin. Bottom panel shows Centrin localization at NBBC (nucleus-basal body connector). Images were acquired with a Leica TCS SP8. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor. B. Grouped maximum intensity projection of 10 z-stacks from the same cell as in (A). Nine grouped projects were montaged together using ImageJ montage stacks tool. Numbers left to the cells indicate the physical distance from z = 0. Images were acquired with Leica TCS SP8 with a z-step size of 0.22 μm. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor. C. Chlamydomonas CW15- cells were expanded following the cryo-ExM protocol. Gels were stained with Centrin (cyan), Tubulin (magenta), and NHS-ester (gray). Top panel shows the whole cell. Note that the Centrin signal was overexposed to unveil the Centrin localization around the nucleus. Middle panel illustrates the inner basal body and striated fiber localization of Centrin. Bottom panel highlights Centrin localization at NBBC. Images were acquired with Leica TCS SP8. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on expansion factor. D. Grouped maximum intensity projection of 10 z-stacks from the same cell as in (C). Nine grouped projects were montaged together using ImageJ montage stacks tool. Numbers left to the cells indicate the physical distance from z = 0. Images were acquired with Leica TCS SP8 with a z-step size of 0.18 μm. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor.
The different fixation procedures are as follows:
Methanol
Chill MeOH at -20 °C for at least 1 h. We usually store it at -20 °C in a little plastic box in which we immerse a small ceramic coverslip rack adapted for 12 mm coverslips, as described here (Guichard et al., 2015). Add 100 μL of Chlamydomonas culture to 12 mm PDL-coated coverslips. Incubate for 5 min, gently blot away liquid, and quickly place in the coverslip rack immersed in the cold MeOH. Place the container at -20 °C for 10 min. Remove coverslips and place in PBS. Store at 4 °C or process for U-ExM cross-linking prevention.
Cryo-ExM
To perform cryo-fixation, first, fill a polystyrene box with dry ice and prepare 5 mL Eppendorf tubes with 2.5 mL of acetone (one tube per condition/coverslip). Snap-freeze the 5 mL tubes in liquid nitrogen. Next, fill a cryo-plunging station (Figure 4A) with liquid nitrogen to cool to -180 °C (approximately 5 min) and add ethane or an ethane-propane mixture to the cryogen container. When the plunging station is prepared, add 100 μL of Chlamydomonas culture to 12 mm PDL-coated coverslips (Figure 2B). Incubate for 5 min and use the plunging tweezers to grab the coverslip at approximately halfway through the coverslip (Figure 4B). Blot the coverslip by applying a piece of filter paper to its bottom for approximately 3 s. There will be an initial excess of media absorbed by the filter paper but allow for a thin layer of media to remain on the coverslip. Plunging stations contain a button or foot pedal to initiate the rapid plunging of the coverslip into the cryogen. Press the button or foot pedal to plunge the coverslip immediately after blotting (Figure 4C). The coverslip is then transferred to solid acetone to proceed to the freeze-substitution step. Note that this transfer must be done in the ethane gas area above the liquid nitrogen to prevent the coverslip from heating up and losing the vitreous state.
To do so, rapidly transfer the plunge frozen coverslip into the 5 mL tube containing frozen acetone (Figure 4D). Place the 5 mL tubes at a 45° angle, immersed in dry ice. Agitate gently overnight using a tabletop shaker (50 rpm), allowing the temperature to rise to -80 °C. The following morning, remove the dry ice to allow the samples to reach 0 °C, which takes approximately 1.5 h. After reaching 0 °C, measured using a thermometer, use a 12-well plate to perform successive 5 min ethanol:water baths in order to rehydrate. The order is as follows: 100% EtOH, 100% EtOH, 95% EtOH, 95% EtOH, 70% EtOH, 50% EtOH, and lastly, PBS. The coverslips can now be processed for U-ExM cross-linking prevention.
Figure 4. The setup and methodology of 12 mm coverslip cryo-fixation. A. Manual plunger for rapid vitrification in liquid ethane. B. Plunging tweezers holding cell-containing coverslip while blotting with filter paper. Note the position of the tweezers halfway down the coverslip. P denotes PDL and cell containing side. C. Plunged coverslip immersed in the liquid ethane of the cryogenic chamber. D. Representative transfer of the coverslip from the cryogenic chamber to the 5 mL tube containing frozen acetone. The tube is opened to facilitate the rapid transfer of the coverslip, and transfer occurs in the ethane gas area, just above the liquid nitrogen.
Membrane optimized cryo-ExM
To better preserve the membranes and retain the lipids, the freeze-substitution step can be done as follows (Louvel et al., 2022). Prior to plunge freezing, make a mixture of 0.1% PFA + 0.02% GA in acetone in a 5 mL Eppendorf tube and put the tube in the liquid nitrogen to cool it, resulting in solid acetone. The tube should stay vertical during this step. Then, proceed for cryo-fixation using plunge-freezing as explained above. For the substitution step, coverslips are transferred into the 5 mL Eppendorf tube containing solid acetone supplemented with PFA and GA. As for the initial cryo-ExM protocol, place the 5 mL tubes at a 45° angle, emersed in dry ice. Agitate gently overnight, allowing the temperature to rise to -80 °C. The next morning, remove the remaining dry ice to bring the temperature up to approximately 0–4 °C. Next, rehydrate the samples by incubating the coverslips in successive 4 °C ethanol baths also containing 0.1% PFA + 0.02% GA. The order is as follows: ethanol 100% + 0.1% PFA + 0.02% GA (5 min), ethanol 100% + 0.1% PFA + 0.02% GA (5 min), ethanol 95% + 0.1% PFA + 0.02% GA (3 min), ethanol 95% + 0.1% PFA + 0.02% GA (3 min), ethanol 70% + 0.1% PFA + 0.02% GA (3 min), ethanol 50% + 0.1% PFA + 0.02% GA (3 min), ethanol 25% + 0.1% PFA + 0.02% GA (3 min), water (3 min), and PBS.
Note: We have also observed that the 70%, 50%, and 25% ethanol steps can be performed without the addition of PFA + GA, and membranes remain preserved, as shown in Figure 5.
Unfixed
Add 100 μL of Chlamydomonas culture to 12 mm PDL-coated coverslips and proceed directly to U-ExM cross-linking prevention step.
After fixation, the protocol proceeds as follows. For each fixed coverslip, make a crosslinking prevention solution containing 2.0% AA/1.4% FA in PBS in a total volume of 1 mL per sample. Add individual coverslips to wells of a 12-well plate and add 1 mL of crosslinking prevention solution. Fill unused wells with ddH2O to prevent evaporation, wrap the edge of the plate with parafilm, and incubate at 37 °C for 3 h.
When the coverslips have been incubating for approximately 3 h, place a gelation chamber (Figure 2) at 4 °C to chill. Place the required number of monomer solution (MS) aliquots (one aliquot per two coverslips), as well as an aliquot of 10% APS and 10% TEMED, on ice.
Note: Aliquots of APS and TEMED can be stored again for future use, while the MS is for one-time use.
The gelation chamber is created by simply placing a damp paper towel to create humidity into a polystyrene box or dish on top of a piece of parafilm (Figure 2C).
Retrieve the samples from crosslinking prevention and remove coverslips from the well. Gently blot away excess liquid and place the chilled humidity chamber into an ice bucket as leveled as possible (Figure 2). Pre-set a P200 pipette to 35 μL and a P20 pipette to 5 μL.
Once the gelation chamber is prepared, make the activated MS by adding 5 μL of APS and then 5 μL of TEMED to the 90 μL MS aliquot. Rapidly, yet calmly, vortex the activated solution and add two 35 μL droplets to the parafilm on the bottom of the humidity chamber (Figure 2). Quickly place the two coverslips, one by one, cell-side facing down, on the droplet with the coverslip taking a 45° angle (Figure 2, inset).
Let the gelation chamber sit for 5 min on ice and then transfer it to a 37 °C environment for 1 h.
Retrieve the gelation chamber from 37 °C and remove the now formed gel associated with the coverslip with a spatula. Place the gel/coverslip associated–side face down, in the well of a 6-well plate. Use a Pasteur pipette to add approximately 1 mL of denaturation buffer to each sample. Place the 6-well plate on a shaker for 15 min at RT. At this stage, the gel will start to expand approximately 2× and dissociate from the coverslip.
Prewarm a heat block to 95 °C and add 1 mL of denaturation solution to 1.5 mL Eppendorf tubes, one per sample. After gels have dissociated from the coverslips, use a spoon to collect the gels and place them in Eppendorf tubes filled with denaturation buffer (gels are slightly rolled to enter in the tube). Make sure that the gels are completely submerged in denaturation buffer and add Eppendorf tubes to the 95 °C heat block for 1.5 h. Place a weight on top of the tubes to prevent them from popping.
After the 1.5 h of denaturation, remove the denaturation solution into a waste solution using a spatula to trap the gel at the edge of Eppendorf tube. Immediately pour the gel into a 500 mL beaker containing approximately 100 mL of ddH2O. Let the gel wash for 30 min and remove the ddH2O. Perform one more wash as just described, then refill with ddH2O and let the gel sit overnight for the first round of expansion. Note: When removing the water during gel washes, a fine-mesh net can be used to cover the waste beaker to catch the gel if it falls.
Antibody staining of gels (post-expansion labeling) (day 1)
Prepare stocks of PBS-0.1% Tween and PBS-2% BSA and store at 4 °C.
Remove the gel from the 500 mL flask and place it on a clean, flat surface. Ensure that the gel does not completely dry. Use a caliper to measure the diameter of the fully expanded gel to determine the expansion factor. From a 12 mm coverslip, a measured expanded gel diameter of 54 mm corresponds to an expansion factor of 4.5 (Figure 2).
Shrink the gels by placing them into 500 mL beakers with 100 mL of PBS and let sit for 15 min. Repeat 1×.
Place the shrunken gels on a flat surface, again ensuring that the gel remains slightly hydrated. Cut the gel into four quarter pieces with a razor blade and place each piece in the well of a 12-well plate.
Note: It is possible to keep the entire gel for immunostaining, but a 6-well plate is then used with twice the volume of antibody (500 μL of total volume vs. 1 mL; next step of this section). Cutting the gel further enables more antibody staining conditions on the same sample. If not all gel pieces are needed for immediate staining, the pieces can be stored in PBS at 4 °C for a few weeks or in a 50% glycerol solution at -20 °C for longer (see Notes section).
Add the desired primary antibody combinations to a final volume of 500 μL of PBS-2% BSA and add this mixture to the desired gels. (Note that antibody concentration requires optimization, but a 1:300 dilution is a good place to start.)
Seal the 12-well plate with parafilm and place it on a shaker at 37 °C for 3 h.
Retrieve the 12-well plate and remove primary antibody solution from wells with a Pasteur pipette. Add approximately 1 mL of PBS-0.1% Tween to gels and gently shake at RT for 10 min. Repeat 2× to completely wash unbound antibodies.
After completion of the PBS-0.1% Tween washes, prepare desired secondary antibody combinations in 500 μL of PBS 2% BSA. Add the secondary antibody solutions to the corresponding gel pieces and wrap the 12-well plate in aluminum foil, as the fluorophore of the secondaries may be photosensitive.
Note: 1:400 dilution is a reasonable concentration for most secondary antibodies.
Place the wrapped 12-well plate on a shaker at 37 °C for 3 h.
Retrieve the 12-well plate and repeat the PBS-0.1% Tween washes described in step D7. Take care to keep the 12-well plate wrapped during the washes.
After completion of the final wash, proceed to either NHS-ester staining, if desired (section E), or re-expansion. For re-expansion, use a spoon or spatula to remove the gel piece from the well, and place it in a 250 mL beaker containing approximately 50 mL of ddH2O. Incubate at RT for 30 min and remove the water. Repeat one more time and then incubate gels in ddH2O until imaging.
Note: NHS-ester staining can also be performed the next day. Keep gel pieces in a 12-well plate, add PBS to the well, and store at 4 °C until NHS-ester staining is performed.
Table 1. Antibodies against tagged proteins
Tag Species Manufacturer Ref U-ExM Dilution
GFP rabbit Torrey Pines TP401 1:200
YFP rabbit Torrey Pines TP401 1:200
mNeongreen* mouse Chromotek 32F6 Non compatible
HA mouse Sigma-Aldrich H6908 1:100
HRV3C rabbit Thermo Fisher PA1-118 1:150
SNAP mouse NEB P9310S 1:100
Antibodies against tagged proteins. List of antibodies against tags, the manufacturer, and dilutions tested in the U-ExM of tagged Chlamydomonas strains.
NHS-ester staining
NHS-ester staining in expansion microscopy has been introduced by M’Saad and Bewersdorf (M’Saad and Bewersdorf, 2020) as well as Mao et al. (2020) and Yu et al. (2020). NHS-ester is an ester-dye conjugate that reacts with primary amines on proteins. With this labeling, it is possible to reveal the cellular context, comparable to electron microscopy images (M’Saad and Bewersdorf, 2020).
Prior to the experiments, make 20 μL working aliquots of NHS-ester dye (1 mg/mL) by suspension in PBS and store at -20 °C. NHS-ester dye is photosensitive. Store in a lightproof box. Note: NHS-ester can be conjugated against many different fluorophores, resulting in preferential labeling of different cellular compartments based on hydrophobicity, pH, etc. (Sim et al., 2021). This protocol describes the concentrations to use in NHS-ester conjugated against Atto 488, 405, 594, and 647.
The day of the experiment, remove the desired number of working aliquots from the freezer and thaw in the dark.
Add the thawed working aliquot to a final volume of 1 mL of PBS to generate a final concentration of 20 μg/mL NHS-ester. Add 500 μL of NHS-ester solution per well containing gels in PBS. Note: one working aliquot of 20 μL of NHS-ester at 1 mg/mL corresponds to staining two wells of a 12-well plate. Diluting further is also possible.
Cover the 12-well plate in aluminum foil and incubate for 1.5 h at RT on a shaker.
Remove the NHS-ester staining solution from wells with a Pasteur pipette and replace it with approximately 1 mL of PBS for washing. Place on the shaker for 10 min. Repeat the washes 4×.
After the last wash, proceed to re-expansion in ddH2O as described in step D11.
BODIPYTM membrane staining
In order to preserve the lipid membranes in U-ExM, cryo-fixation followed by freeze-substitution containing PFA and GA should be used to obtain an optimal result. To visualize the membranes, it is then possible to use BODIPYTM in the same way as with NHS-ester (Figure 5).
Figure 5. BODIPYTM lipid staining of Chlamydomonas using membrane-optimized cryo-ExM protocol. A. Chlamydomonas CC124- strain was cryo-fixed and freeze-substituted in acetone supplemented with 0.1% PFA and 0.02% GA prior to expansion (see cryo-fixation step for detail). The gel was stained with BODIPYTM. The image shows a single z-plane. Observed structures are labeled with a star. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor. B. Grouped maximum intensity projection of 12 z-stacks from the same cell as in (A). Nine grouped projects were montaged together using ImageJ montage stacks tool. Note that the flagellar membrane and pyrenoid structures are shown here, which were not visible in the single z-plane shown in (A). Numbers under the cells indicate the physical distance from z = 0. Images were acquired with Leica TCS SP8 with a z-step size of 0.18 μm. Scale bar: 10 μm. The scale bar shows the measured physical length that was not rescaled based on the expansion factor. C. The plot profile is drawn on the flagellar membrane using ImageJ line scan tool [shown as 1 in (B)]. L shows the measured peak-to-peak distance. The measured length is corrected by the expansion factor. The corrected distance shows the diameter of the flagellar membrane based on BODIPYTM staining. D. The plot profile is drawn on the cell wall using ImageJ line scan tool [shown as 2 in (B)]. W shows the distance between 50% intensity of the peak.
Make 10 μL aliquots of BODIPYTM at 2 mM by diluting in DMSO. Unused aliquots should be stored at -20 °C, away from the light.
To stain for BODIPYTM, perform a 1:200 dilution with PBS 2% BSA and add to wells containing the shrunken gels in PBS. One aliquot of 10 μL of BODIPYTM should be enough to stain two pieces of gel in the 12-well plate.
Cover the 12-well plate in aluminum foil and incubate for 1.5–2 h at RT on a shaker.
Remove the BODIPYTM staining solution from wells with a Pasteur pipette and replace with approximately 1 mL of PBS for washing. Place on the shaker for 10 min. Repeat the washes 3×.
After the last wash, proceed to re-expansion in ddH2O as described in step D11.
Imaging of gels (days 2–7+)
Remove the expanded gel piece from the 250 mL beaker by pouring out the water while trapping the piece (with a spatula or manually by wearing a glove). Place it on a clean and flat surface and cut a square piece for imaging that is approximately 1.5 cm × 1.5 cm in size. Return the remaining gel pieces to the beaker containing ddH2O for potential future use (see Note 5).
Place the 1.5 cm × 1.5 cm gel piece in the center of a 24 mm coverslip and place the coverslip inside of the imaging chamber (Imaging chamber example, Figure 2A).
Note: The gel piece should not touch the edges of the imaging chamber to prevent drifting during acquisition.
Make sure an appropriate stage mount is present on the microscope stage and insert the imaging chamber.
Change the objective to medium magnification (20× or similar) to determine the side of the gel containing the cells.
Note: This may require removing the imaging chamber and flipping the gel piece with a spatula. If the gel is sided inversely, heavy background fluorescence can be observed in the imaging plane.
When the correct sidedness of the gel is determined, gently blot the four edges of the gel as well as the reverse side to remove excess water. Retrieve a PDL-coated 24 mm coverslip and place the cell-containing side face down in the center. Use a small paintbrush or the side of a spatula to gently mount the gel against the PDL coating, until the gel is completely adherent to the surface. Place the PDL-coated coverslip containing the gel back into the imaging disk. Gently add approximately 5 μL of ddH2O to the top of the gel and place a 24 mm coverslip on top to create a mini humid chamber.
Note: Failure to remove excess liquid of the gel or poor PDL coating results in substantial sample drift during acquisition and failure to collect images. See Notes section for assistance with sample drift.
Return the imaging chamber to the microscope stage and perform acquisition, adding 5 μL of ddH2O approximately every hour to prevent gel shrinkage. Add a non-coated 24 mm coverslip to the top of the chamber to prevent water evaporation.
Chlamydomonas cells are positioned in different orientations within the gel. Examine the gel to find a sample in the desired orientation.
When imaging with a Leica microscope, image in Lightning mode, with the strategy as “Adaptive” and “Water” selected as the mounting medium for the best results. If using a Leica confocal such as an SP8, set the pixel size to 35 nm for maximum resolution. If imaging with a Zeiss microscope such as an LSM980, image in Airyscan mode with “Resolution” selected. (Representative z-stack, Video 1.)
Note: Acquiring a pixel size of 35 nm is the maximum boundary when Nyquist sampling, owing to the expected lateral resolution size of 70 nm as previously published (Chen et al., 2015). However, given that the optical properties of the microscope are unchanged, sampling with a larger pixel size will decrease acquisition time with minimal impact on the resolution. See data analysis for a more in-depth explanation of the resolution.
Video 1. Representative z-stack of a top view of an expanded Chlamydomonas cell. Cryo-ExM of a transgenic cell expressing IFT46-mNeonGreen:D1bLIC-mCherry imaged with a LSM980 confocal microscope using Airyscan mode with a step size of 0.15 μm. The staining was performed with NHS-ester (gray), anti-Tubulin antibody (magenta), and anti-Acetylated Tubulin antibody (green). Scale bar corrected by the expansion factor: 5 μm.
Data analysis
Traditional application of confocal microscopy is conservatively diffraction-limited to approximately 200 nm in the lateral dimensions (MacDonald et al., 2015). A normal expansion factor of 4.2–4.5× yields an effective lateral resolution of approximately 70 nm as measured in the original expansion microscopy protocol (Chen et al., 2015). With this resolution, it is possible to resolve the membranes, labeled with BODIPY, on either side of the flagella with a distance of 188 nm (Figure 5C). Furthermore, when measuring the thickness of the cell wall, which is also stained by BODIPY, the 40 nm thickness of the cell wall (Horne et al., 1971) appears as an 80 nm thick layer. This is consistent with a resolution of approximately 70 nm.
To measure the isotropy of an expanded sample, features observable in non-expanded Chlamydomonas can be measured post-fixation, but prior to starting cross-linking prevention of U-ExM. For example, the nucleus size, pyrenoid area, or flagellar length can be measured by brightfield microscopy and then re-measured after U-ExM to confirm isotropy.
When analyzing collected images, it is necessary to know the expansion factor to properly scale distances. As mentioned in Section D, the expansion factor can be used to scale images. In our laboratories, we also routinely use a well described benchmark, such as the width of a basal body, to scale images. For example, a basal body with an expanded proximal width of 900 nm corresponds to an expansion factor of 4, given the 225 actual width of the basal body. Otherwise, we found in U-ExM that measuring the gel before and after expansion is quite reliable to determine the expansion factor, with values similar to those obtained by measuring the basal body, so this simpler approach can be used if you do not have an internal molecular ruler.
A broad range of cellular compartments can be visualized through antibody labeling, for example IFT train localization at the basal body and along the flagella by antibody labeling against an IFT component (Figure 6A), contrasted against the labeling of the chloroplast in the cell body (Figure 6C, Video 2).
To perform analyses, any regular image analysis software can be used. We routinely use Fiji, in particular the plot profile function to assess colocalization and measure the distances and periodicities along a substrate, using peak-to-peak or full-width half-maximum. For instance, we stained for the Dynein regulatory complex subunit 3 (DRC3), a flagellar protein important for the regulation of the flagellar waveform (Huang et al., 1982), and found a colocalization signal with the microtubule doublets (Figure 6B).
Figure 6. Representative examples of Chlamydomonas imaging in ultrastructure expansion microscopy (U-ExM). A. Methanol-fixed Chlamydomonas IFT46-YFP cell stained for tubulin and IFT46. Note the accumulation of intraflagellar transport trains (IFT) particles at the basal body and flagellar tip, as well as train-like staining on the flagellar axoneme. Scale bar corrected by the expansion factor: 2 μm. B. High-magnification image of tubulin staining with SNAP-tagged DRC3, a component of the dynein regulatory complex. Bottom right panel is a plot profile generated from the dotted line in the DRC3 staining. Scale bar corrected by the expansion factor: 50 nm. C. Cryo-ExM-fixed Chlamydomonas cell stained for the chloroplast (PsbA staining), tubulin, and NHS-ester. Scale bar corrected by the expansion factor: 2 μm. Images were acquired with a Leica Thunder (A) and LSM980 Airyscan2 (B, C).
Video 2. Representative z-stack of an expanded Chlamydomonas cell with chloroplast staining.Cryo-ExM of a transgenic cell expressing IFT46-mNeonGreen:D1bLIC-mCherry imaged with a LSM980-NLO confocal microscope using Airyscan mode with a step size of 0.15 μm. The staining was performed with NHS-ester (gray), anti-acetylated Tubulin antibody (magenta), and anti-PsbA antibody (green). Scale bar corrected by the expansion factor: 10 μm.
Notes
Sodium acrylate
As with all expansion microscopy protocols, the quality of sodium acrylate is a frequent cause of expansion microscopy issues. Dissolving the sodium acrylate powder gradually into stirring water facilitates dissolution. Take care to dissolve the sodium acrylate on ice at 4 °C, or immediately transfer to 4 °C. Different sodium acrylate batches are variable in purity. Only solutions that are clear or faintly yellow should be used in experiments.
Fixation methods
The desired fixation method should be considered prior to starting the experiment. Each method has its benefits and detractions, but cryo-fixation offers the best preservation of samples in their native state (Dubochet et al., 1988). If cryo-fixation is unavailable, consider the cellular compartment to be stained, and proceed accordingly with either unfixed, methanol, or PFA/GA fixations.
When fixing, especially if using cryo-fixation, it is important to make sure that the coverslip is not overly blotted. Completely drying the coverslip will result in dehydrated cells where the cytoplasmic volume shrinks but the cell wall remains at the same volume, resulting in a large gap between the cytoplasmic volume and cell wall, creating the appearance of a “halo” in expansion microscopy. Avoiding over-blotting of the coverslip circumvents this artifact.
Broken flagella while imaging
Chlamydomonas often adheres to surfaces via their flagella. In these cases, the flagella may fragment during expansion, and cracks or pieces can be observed on the coverslip. This is common, and the gel can be scanned to find intact flagella.
Microscopic drift
As stated in the Procedure section, sample drift at the microscope is a common problem, resulting in failure to acquire usable images. We found that plasma-cleaning the surface of 24 mm coverslips prior to the addition of PDL greatly enhances the adhesion between gel and coverslip. The gel should remain completely adherent even after the addition of the water to prevent gel shrinkage. Avoid imaging at the edges of the gels and stay towards the center for data collection. If the gels are noticeably drifting at the initiation of imaging, remove the imaging chamber from the microscope, gently blot the edges of the gel, and return to the microscope for imaging. Freshly PDL-coated coverslips result in less drift.
Gel storage
The imaging quality of a gel decreases over time due to progressive fluorescence degradation or antibody loss. We recommend imaging the gels as close to the time of staining as possible. Imaging up to seven days post-staining is generally permissible once they have been expanded in ddH2O. Longer storage of both labeled and unlabeled gels can be attained by immersing expanded gels in ddH2O containing 50% glycerol in a 10 cm dish, shaking for 3 h, wrapping the dish in parafilm, and placing at -20 °C. To thaw the frozen gels, wash once with ddH2O to remove the excess glycerol and then wash at least three times in PBS (1×) for 1 h. (The gel will shrink in PBS and force the glycerol out.) Make sure to wash off the glycerol completely or a signal background might be induced in the staining.
Recipes
All stocks are assumed to be the same as those listed in reagents.
TAP, 0.5 L
TRIS, 1.21 g
Phosphate buffer II, 0.5 mL
Solution A, 5 mL
Hutner’s trace elements, 0.5 mL
Acetic acid, 0.5 mL
Mix the ingredients and add water to bring to 0.5 L. Adjust pH to 7.0. Autoclave to sterilize.
Phosphate buffer II
1 M K2HPO4, 250 mL
1 M KH2PO4, 170 mL
Adjust the pH to 7
Solution A (40×)
Tris, 96.8 g
Phosphate buffer (pH 7), 40 mL
Acetic acid, 40 mL
Adjust the solution to 1L with distilled water.
Formaldehyde/acrylamide mixture (1.4% FA, 2% AA), 1 mL/1 coverslip
FA (38%), 38 μL
AA (40%), 50 μL
PBS 1×, 912 μL
Mix ingredients and add 1 mL to a well of a 12-well plate. The table below displays the volume of each reagent when scaling up for number of coverslips.
Coverslips 1 2 3 4 5 6 7 8
FA (1.4%) 38 76 114 152 190 228 266 304
AA (2.0%) 50 100 150 200 250 300 350 400
PBS (1×) 912 1824 2736 3648 4560 5472 6384 7296
Sodium acrylate solution (38% w/w)
Sodium acrylate, 19 g
Nuclease-free water, 31 mL*!
Add the sodium acrylate incrementally in stirring water to mix. Attempt to work at 4 °C and store at 4 °C immediately after sodium acrylate is dissolved.
*31 mL of nuclease-free water = 31 g of nuclease-free water. 31 g of nuclease-free water + 19 g of sodium acrylate = 50 g total. 19 g of sodium acrylate/50 g total = 38% w/w solution.
!38% w/w solution of sodium acrylate corresponds to a 46% w/v solution owing to the high density of sodium acrylate in solution.
Monomer solution, 10 aliquots of 90 μL*
Sodium acrylate solution (38% w/w), 500 μL
Acrylamide (40%), 250 μL
BIS (2%), 50 μL
10× PBS, 100 μL
Mix the ingredients on ice and vortex for 30 s. Make 90 μL aliquots and store at -20 °C for three weeks. One aliquot is used for two coverslips.
10% TEMED, 10 aliquots of 100 μL
TEMED, 100 μL
Nuclease-free water, 900 μL
Mix TEMED and nuclease-free water. Make 100 μL aliquots and store at -20 °C.
10% APS, 10 aliquots of 100 μL
APS, 0.1 g
Nuclease-free water, 1,000 μL
Mix APS powder and nuclease-free water. Make 100 μL aliquots and store at -20 °C.
Denaturation buffer, pH 9, 100 mL
SDS solution (350 mM stock solution, 10 g in ddH2O, 100 mL final volume), 57.14 mL
NaCl (5 M), 4 mL
Tris-BASE, 0.6 g
ddH2O, fill to 100 mL
Dissolve 0.6 g of TRIS-BASE in 10 mL of ddH2O. Add NaCl and SDS. Adjust to pH 9 with HCl. Fill to 100 mL with ddH2O and store at RT.
PBS-Tween (0.1% w/v) (1×, 1 L)
Prepare 1× PBS:
NaCl, 8 g
KCL, 0.2 g
Na2HPO4,1.44 g
KH2PO4, 0.24 g
Adjust the solution to 1 L with distilled water.
Add 100 µL Tween 20 detergent.
PBS-BSA (2%) (1×, 100 mL)
Dissolve 2 g of BSA in 100 mL of 1× PBS.
Acknowledgments
We thank the Light Imaging Facility of Human Technopole and the Photonic Bioimaging Center of the University of Geneva. We thank Vincent Louvel for critical reading of the manuscript. We acknowledge EMBO fellowship ALTF 537-2021 to N.K. We would like to acknowledge funding from Human Technopole, the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (grant agreement no. 819826) to G.P. This work was also supported by the Swiss National foundation 310030_205087 attributed to P.G. and V.H. as well as by the University of Geneva. This protocol was derived from the original work of van den Hoek et al. (2022).
Competing interests
The authors declare no competing interests.
References
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Stereotactic Delivery of Helper-dependent Adenoviral Viral Vectors at Distinct Developmental Time Points to Perform Age-dependent Molecular Manipulations of the Mouse Calyx of Held
CK Christian Keine
MA Mohammed Al-Yaari
TR Tamara Radulovic
SJ Samuel M. Young Jr.
Published: Vol 13, Iss 16, Aug 20, 2023
DOI: 10.21769/BioProtoc.4793 Views: 609
Reviewed by: David PaulOlga Kopach Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Oct 2022
Abstract
Synapses are specialized structures that enable neuronal communication, which is essential for brain function and development. Alterations in synaptic proteins have been linked to various neurological and neuropsychiatric disorders. Therefore, manipulating synaptic proteins in vivo can provide insight into the molecular mechanisms underlying these disorders and aid in developing new therapeutic strategies. Previous methods such as constitutive knock-out animals are limited by developmental compensation and off-target effects. The current approach outlines procedures for age-dependent molecular manipulations in mice using helper-dependent adenovirus viral vectors (HdAd) at distinct developmental time points. Using stereotactic injection of HdAds in both newborn and juvenile mice, we demonstrate the versatility of this method to express Cre recombinase in globular bushy cells of juvenile Rac1fl/fl mice to ablate presynaptic Rac1 and study its role in synaptic transmission. Separately, we overexpress CaV2 α1 subunits at two distinct developmental time points to elucidate the mechanisms that determine presynaptic CaV2 channel abundance and preference. This method presents a reliable, cost-effective, and minimally invasive approach for controlling gene expression in specific regions of the mouse brain and will be a powerful tool to decipher brain function in health and disease.
Key features
• Virus-mediated genetic perturbation in neonatal and young adult mice.
• Stereotaxic injection allows targeting of brain structures at different developmental stages to study the impact of genetic perturbation throughout the development.
Keywords: Calyx of Held Synaptic transmission Rac1 Presynaptic mechanisms Helper-dependent adenovirus Stereotactic injection
Background
The ability to manipulate specific neuronal populations at defined time points is critical for understanding the regulation of neural circuit function and for the creation of animal models of neurological disorders. Neurons interact via synapses, specialized structures that serve as points of neuronal communication. Synaptic dysfunction due to alterations or mutations in synaptic proteins has been linked to various neurological and neuropsychiatric disorders (Lepeta et al., 2016; Taoufik et al., 2018; Wang et al., 2018; Koopmans et al., 2019). Therefore, the ability to manipulate synaptic proteins in vivo can provide insight into the molecular mechanisms underlying these disorders and aid in the development of new therapeutic strategies. While constitutive knock-out animals have served as powerful tools in deciphering brain function, they are limited by potential developmental compensation, off-target effects, and embryonic or postnatal lethality (Sigmund, 2000; Eisener-Dorman et al., 2009; Navabpour et al., 2020; Suzuki et al., 2020; Sztretye et al., 2020). Furthermore, creating and establishing new genetic mouse lines can be time- and cost-prohibitive. Recombinant viral vector technology has emerged as an indispensable tool in neuroscience for manipulating neuronal function, in vivo and in vitro, at both the cellular and molecular levels (Lentz et al., 2012; S. H. Chen et al., 2019; Haery et al., 2019). The current approach outlined in this protocol uses helper-dependent adenoviral vectors (HdAd) for age-dependent protein ablation in mice. HdAds are third-generation Ad vectors that contain only the Ad viral inverted terminal repeats (ITRs) and packaging sequence. Due to their large packaging capacity of 37 kb compared to other viral vectors (adeno-associated virus: 5 kb, lentivirus: 9 kb), rapid and stable long-term gene expression, low toxicity, and the ability to enable long-term phenotypic correction in preclinical animal models (Vetrini and Ng, 2010; Brunetti-Pierri and Ng, 2017; Watanabe et al., 2021), they have emerged as a useful tool in studying neuronal circuit function (Z. Chen et al., 2013; Montesinos et al., 2015 and 2016; C. Chen et al., 2017a and 2017b; Lübbert et al., 2017 and 2019; Dong et al., 2018; Ostrovskaya et al., 2018; Sutton et al., 2018; Radulovic et al., 2020; Keine et al., 2022). The high packaging capacity of HdAds allows for the packaging of large promoters and large genes of interest such as calcium channels (e.g., 7 kb for CaV2.1 α1 subunit cDNA) (C. Chen et al., 2017a and 2017b; Lübbert et al., 2017; Ostrovskaya et al., 2018; Sutton et al., 2018; Lübbert et al., 2019) in combination with regulatory elements to enhance long-term gene expression and markers for fluorescence and electron microscopy imaging (Thomas et al., 2019). Using Cre-expressing viral vectors in combination with commercially available conditional knock-out mouse lines and the stereotactic injection procedure outlined here allows for targeted ablation of various proteins at distinct developmental time points to study synaptic function and development.
In this protocol, we demonstrate the versatility of this approach. Specifically, we expressed Cre recombinase in globular bushy cells of juvenile Rac1fl/fl mice, thereby specifically ablating Rac1 in the presynaptic calyx of Held to study its role in synaptic transmission (Keine et al., 2022). In another study, we used HdAd’s large packaging capacity and fast onset of gene expression to overexpress CaV2 α1 subunits at two distinct developmental time points [postnatal day 1 (P1) and P14] to elucidate the mechanisms that determine presynaptic CaV2 channel abundance and preference (Lübbert et al., 2019). While the combination of large packaging capacity and fast onset of gene expression is specific to HdAds, the many aspects of the protocol are applicable to adeno-associated (AAV) and lentiviral vectors (LVV) within the limitations of expression timelines and packaging of large constructs.
In summary, this method presents a reliable, cost-effective, and minimally invasive approach for manipulating gene and protein expression in specific mouse brain regions. Finally, this protocol can be adapted to study other genes and brain regions of interest, making it a valuable and broadly applicable tool for neuroscience research.
Materials and reagents
Biological materials
Helper-dependent adenovirus prepared as in Montesinos et al. (2016) or from the University of Iowa Viral Vector Core (catalog number: VVC-Young-6896)
Reagents
HEPES (Sigma-Aldrich, catalog number: H3375)
Magnesium chloride solution, 1 M (MgCl2) (Sigma-Aldrich, catalog number: M1028)
Saccharose/sucrose (Sigma-Aldrich, catalog number: S7903)
HCl 1 N (Carl-Roth, catalog number: K025.1)
NaOH 1 N (Carl-Roth, catalog number: K021.1)
D-Mannitol (Sigma-Aldrich, catalog number: M4125)
Povidone-iodine solution, 10% (Betadine)
Ethanol 70% solution (Fisher Scientific, catalog number: BP82031GAL)
Lidocaine hydrochloride solution, 0.5%
Bupivacaine solution 0.25%
Meloxicam-SR solution 2 mg/mL (ZooPharm, IZ-800000-191903)
Hydrogen peroxide solution 3% (CVS)
Fluorescent dye of high molecular weight (e.g., Tetramethylrhodamine, Thermo Fisher, catalog number: D1817)
Fluorescent microspheres (e.g., Fluoresbrite, Polysciences, catalog number: 16662-10)
Solutions
Storage buffer (see Recipes)
Mannitol solution (20%) (see Recipes)
Virus injection solution (see Recipes)
Recipes
Storage buffer
Reagent Final concentration Amount
Sucrose 250 mM 428 mg
HEPES 10 mM 12 mg
MgCl2 (1 M) 1 mM 5 μL
H2O n/a 4.995 mL
Total n/a 5 mL
Dissolve sucrose and HEPES in 4 mL of ultrapure water while stirring and add MgCl2.
Adjust pH to 7.4 with 1 N HCl and 1 N NaOH.
Bring up to a final volume of 5 mL.
Filter and aliquot in 50 μL aliquots and store at -20 °C.
Mannitol solution (20%)
Reagent Final concentration Amount
Mannitol 20% 2 g
H2O n/a 10 mL
Total n/a 10 mL
Dissolve mannitol in 9 mL of ultrapure water while stirring.
Bring up to a final volume of 10 mL.
Filter and aliquot in 10 μL aliquots and store at -20 °C.
Virus injection solution
Reagent Final concentration Amount
Mannitol solution (20%) 6.7% 10 μL
Storage buffer n/a 17 μL
Virus stock solution
(3 × 1012 vp/mL)
3 × 1011 vp/mL 3 μL
Total n/a 30 μL
Laboratory supplies
Eppendorf safe-lock tubes 0.5 mL (Eppendorf, catalog number: 0030121023)
Glass beaker 400 mL (Fisher Scientific, DWK Life Science, catalog number: 09-841-102)
Glass capillaries 3.5” (Drummond)
Micropipettes (Blaubrand, intraMark, catalog number: 708707)
Microliter syringe (Hamilton, 700 series, catalog number: 80600)
Autoclaved cotton swabs (Q-tips)
Parafilm wrapping film (Fisher Scientific, catalog number: S37441)
Ceramic scoring wafer (Restek, catalog number: 20116)
Absorption spears (Fine Science Tools, catalog number: 18105-01)
Ophthalmic ointment (Altaire Pharmaceuticals, Puralube Vet Ointment, NDC: 17033-211-38)
Hair remover lotion (Church & Dwight, Nair)
Mineral oil (Sigma-Aldrich, catalog number: M5904)
Reverse-cutting needle and suture thread (Surgical Specialties, C17, 5-0, catalog number: 1013B)
Equipment
Foot-pedal drill (Foredom, MH-170)
Micro drill burrs 0.7 mm (Fine Science Tools, catalog number: 19008-07)
Nanoliter injector (Neurostar, NanoW)
Infusion pump (Chemyx, Fusion 100)
Mechanical pipette 0.5–10 μL (Eppendorf, Research Plus, catalog number: 3123000071)
Mechanical pipette 10–100 μL (Eppendorf, Research Plus, catalog number: 3123000047)
Robot stereotaxic instrument (Neurostar)
Stereotaxic frame (David Kopf Instruments, model: 940)
Horizontal micropipette puller (Sutter Instrument, P-1000 Micropipette Puller)
Angled probe (Fine Science Tools, catalog number: 10140-02)
Homeothermic monitoring system (Harvard Apparatus, catalog number: 55-7020)
Software
StereoDrive (Neurostar, v3.1.5, https://neurostar.de/)
Procedure
Manufacture of injection pipettes
NOTE: In the following protocol, injection pipettes are manufactured from Blaubrand or Drummond glass capillaries for P1 and P14 injections, respectively. While for P1 injections a syringe-pump is used, P14 injections are performed using a computer-controlled nano-injector, which offers more control of injection speed and volume. However, both types of injections can be performed with either injection method.
For injections at P1, pull glass pipettes from Blaubrand intraMark glass micropipettes using a P-1000 horizontal micropipette puller (2.5 mm box filament) with the following settings:
Sutter Pipette Cookbook Type B protocol (microinjection or sharp electrode):
Heat: ramp; pull: 55–60; velocity: 70–75; time: 120; pressure: 500
For injections at P14 and older, pull the injection pipettes from 3.5” Drummond glass capillaries using a P-1000 horizontal micropipette puller (2.5 mm box filament) with the following settings:
Sutter pipette Cookbook Type A Protocol:
Heat: ramp; pull: 18–20; velocity: 32–35; time: 250; pressure: 500
Carefully cut the tip of the pipette using a ceramic scoring wafer, ceramic razor blade, or another glass pipette. Hold the pipette in one hand and carefully slide the ceramic scoring wafer perpendicular to the pipette axis to cut the pipette to a tip opening of 25–50 μm. Using fine forceps to cut the pipette tip is possible but might result in broken and uneven edges, which could impair viral delivery results and cause tissue damage.
Confirm that the pipette tip is cut smoothly and not broken using a standard laboratory microscope (Figure 1).
Figure 1. Tip of injection pipette after cutting with a ceramic scoring wafer. A. The tip should be cut to an approximate diameter of 25–50 μm and appear smooth and without major irregularities. B. A broken tip might result in poor injection results and tissue damage. Images were acquired using a 10× objective. Scale bars: 50 μm.
Virus injection in P1 mice
Slowly thaw the virus solution on ice.
Add mannitol and storage buffer to the desired concentration of viral particles and keep the solution on ice. Ensure that the final number of injected virus particles is < 1 × 109:
If the stock concentration of the virus is 3 × 1012 virus particles per milliliter (vp/mL), use the dilution as in Recipe 3.
Carefully mix the solution and keep it on ice for the rest of the experiment. When injecting 1 μL of this solution, a total number of 3 × 108 virus particles will be delivered at the injection site. The total injection volume might be adjusted depending on the size of the target structure. Adjust the dilution in accordance with the virus particle concentration of the stock solution.
Fill the Hamilton syringe with mineral oil and place it into the infusion pump. Ensure to remove any air bubbles from the solution. Attach a tightly fitting plastic tube to the Hamilton syringe and fill it with mineral oil.
To fill the injection pipette, first fill 1–2 μL of virus solution into a Blaumark glass capillary. Using a small piece of silicon tubing, connect the glass capillary containing the virus solution to the back of the injection pipette so that the virus solution can flow between the glass capillary and injection pipette. Connect the other side of the glass capillary to the oil-filled tubing of the Hamilton syringe.
Use the syringe pump to push the virus solution from the glass capillary into the tip of the injection pipette, trying to minimize the amount of air in the system.
Anesthetize the mouse pups with cryoanesthesia (deep hypothermia): place one pup at a time, tail first, into the cut-off finger of a laboratory glove. Fill a 400 mL glass beaker with 70% crushed ice and 30% ice-cold water. Attach the glove finger to the inside wall of the glass beaker using a binder clip or clothespin so that the pup is immersed in ice water. Make sure that the glove opening is above the water level so that water does not enter the glove and the pup is not in direct contact with ice or water. Remove pups from the ice bath after a maximum period of 5 min to avoid any damage by hypothermia. The following steps should be performed swiftly, as the duration of cryoanesthesia is limited to 10–15 min.
Check the mouse pup for pedal reflex indicating proper anesthesia. Place the mouse pup in the stereotactic frame in a prone position and use soft ear bars to hold the head in place. Position the mouse as straight as possible and tighten the ear bars sufficiently to prevent head movement when slight pressure is applied.
Clean the skin on the head with alternating swaps of Betadine and 70% ethanol (three repetitions each).
Locate the superior sagittal sinus and identify the position of the bregma and lambda points beneath the skin. A light source next to the head might aid in identifying the sagittal sinus.
Position the injection pipette on top of the bregma point and set stereotaxic coordinates to zero. This point will be used as the zero reference for all future coordinates.
Lift the pipette and move 5.2 mm caudally and 1.5 mm laterally to mark the injection site. These coordinates might vary depending on the animal’s size.
Carefully lift the skin with forceps and make a small incision using sharp scissors at the previously marked injection site.
Penetrate the soft skull with the injection pipette and lower it to a depth of 2.9 mm below the surface of the skull. Move the pipette quickly to pinch through the skull but slowly once the pipette enters the brain.
Slowly inject the virus solution (< 1 μL) at a speed of 500 nL/min.
After the injection, leave the injection pipette in place for one minute to allow any pressure to dissipate.
Slowly remove the injection pipette.
Place the pup under a heating lamp or a heat-controlled cage maintained at 33–36 °C and allow it to recover. Remove any leftover blood from the skin with water.
When pups are awake and actively mobile, move them back to their home cage with their mother. Cover the pups with bedding material from the cage so they acquire a familiar smell, which helps to minimize cannibalism.
Virus injection in P14 mice
Slowly thaw the virus solution on ice.
Add mannitol and storage buffer to the desired concentration of viral particles and keep the solution on ice. Ensure that the final number of injected virus particles is < 1 × 109.
If the stock concentration of the virus is 3 × 1012 vp/mL, use the dilution as in Recipe 3.
Carefully mix the solution by slowly pipetting up and down; then, keep it on ice for the rest of the experiment. When injecting 1 μL of this solution, a total number of 3 × 108 virus particles will be delivered at the injection site. The total injection volume might be adjusted depending on the size of the target structure. Adjust the dilution in accordance with the virus particle concentration of the stock solution.
Anesthetize the animal using 5% isoflurane in oxygen for induction; then, maintain at 1%–2% isoflurane and ensure proper anesthesia throughout the procedure by testing for the absence of the pedal reflex.
Inject Meloxicam subcutaneously (6 mg/kg body weight).
Apply ophthalmic ointment to the animal’s eyes.
Carefully remove hair from the animal’s head using hair remover lotion. Remove all the remaining lotion with water as the smell might disturb the mother when returning the animals after the procedure.
Place the animal in the stereotaxic frame and fixate with ear bars. Gently pull out the mouse’s tongue to facilitate breathing.
Clean the skin with alternating swipes of betadine and ethanol (70%), three times each.
Subcutaneously inject lidocaine/bupivacaine solution at the prospective incision site on top of the skull.
Cut the skin on the head at the midline along the anterior-posterior axis and expose the skull so that lambda and bregma marks are visible. Bregma is located at the intersection of the rostral-caudal fissure and the second major medial-lateral fissure. One drop of 3% hydrogen peroxide applied to the skull may aid in the visualization of sutures. After one minute, rinse thoroughly with water.
Using the Neurostar Stereotaxic software, determine the head size, orientation, and tilt. If necessary, re-adjust the head to minimize lateral tilt and improve reproducibility. When injecting into the cochlear nucleus, the head should be slightly tilted rostrally, with lambda approximately 1 mm higher than bregma. This prevents the injection pipette from damaging the superior sagittal sinus when vertically penetrating the brain.
Select the injection site in the Neurostar Stereotaxic software (e.g., cochlear nucleus) and determine the entry site. If needed, gently push the occipital muscle further caudal.
Using the foot-pedal-controlled drill equipped with a micro drill burr, drill a hole (1 mm diameter) into the skull, taking care not to damage underlying blood vessels and brain tissue. Remove any remaining debris from the drill site and pinch through the meninges with a pointy angled hook.
Mount the glass injection pipette into the nano-inject device and lower the plunger to the lowest possible position without damaging the glass pipette.
Pipette a drop of mineral oil onto parafilm. With the pipette tip immersed in the mineral oil, retract the plunger to tip-fill the injection pipette with 50–100 nL of mineral oil. This mineral oil layer will help to keep the plunger separated from the virus solution.
Tip-fill the rest of the pipette by pipetting a drop of virus solution onto parafilm. With the pipette tip in the solution, retract the plunger of the nanoliter injector until the plunger is completely retracted and the pipette is filled with virus solution.
Place the injection pipette above the drill hole and make sure the hole is clear of any debris and the pipette tip is intact.
Slowly lower the injection pipette into the target region.
Inject pipette solution (< 1 μL) at slow speed (100 nL/min); then, leave the pipette in place for another minute.
Slowly retract the pipette and repeat steps C17–C19 to proceed to another injection site. Note that the injection pipette must be removed completely from the brain before another injection area can be targeted.
After completing all injections, clean the drill hole and suture the skin above the skull with a sterile reverse-cutting needle and suture thread (5-0).
Place the animal under a heating lamp or a heat-controlled cage maintained at 33–36 °C to recover until the anesthesia wears off and the animal is fully mobile, alert, and shows no signs of pain.
Return the animal to its home cage with its mother and provide liquid gel for additional support.
Data analysis
The virus injected into the mouse brain expressed the target protein (e.g., Cre recombinase) together with a fluorescent reporter (e.g., EGFP) to verify viral transfection and protein expression (Figure 2). During patch-clamp experiments, the EGFP expression was used to visualize target neurons, and data were acquired from EGFP-positive neurons and EGFP-negative controls. For new preparations, it is important to validate that virus injection alone does not influence the parameter of interest. To test this, a separate control group should be used in which only the EGFP-expressing virus is injected and compared to non-injected control animals.
For statistical analysis, individual calyx terminals were considered independent samples. Statistical analysis was performed in MATLAB (RRID:SCR_001622) using a two-tailed unpaired Student’s t-test with Welch’s correction or a two-tailed Mann-Whitney U test, depending on the data distribution, but other statistical software (e.g., GraphPad Prism, SPSS) may be used. Effect sizes were calculated using the MES toolbox in MATLAB (Hentschke and Stüttgen, 2011). Calyx terminals in treatment and control mice should be sampled from the same region of the medial nucleus of the trapezoid body (MNTB) to minimize location bias.
Figure 2. Injection of EGFP-expressing helper-dependent adenovirus viral vectors (HdAds) into the cochlear nucleus of a juvenile mouse. A. Confocal tile scan of a P28 mouse brain slice. EGFP-expressing HdAd was injected in the cochlear nucleus (CN) at P14. Note the strong fluorescence signal at the injection site and the fibers forming the ventral acoustic stria (VAS), terminating in the contralateral superior olivary complex including the medial nucleus of the trapezoid body (MNTB) (rectangle). The ipsilateral MNTB and the contralateral CN show no fluorescence signal, indicating targeted expression in the contralateral calyx of Held. For better orientation, the borders of the brain slice are indicated with a dotted line. For display purposes, brain slices were cut at a 30° dorsoventral angle to preserve the nerve connections between the CN and MNTB. Scale bar: 500 μm. B. Confocal maximum projection image of the MNTB region from the rectangle in A, showing passing fluorescent fibers and calyx of Held presynaptic terminals. Scale bar: 100 μm.
Validation of protocol
This protocol has been validated for viral injections at both P0/P1 (Montesinos et al., 2015 and 2016; Lübbert et al., 2017; Dong et al., 2018; Thomas et al., 2019; Radulovic et al., 2020) and P14 (Lübbert et al., 2019; Keine et al., 2022), and results have been reported in peer-review publications. To verify the coordinates for injection, the virus solution can be replaced with a fluorescent dye of high molecular weight or fluorescent microspheres. Brain slices can then be imaged using a standard fluorescent microscope and injection coordinates adjusted if needed.
Successful virus transfection can be verified by co-expression of a fluorescent marker (e.g., EGFP). Expression of Cre-recombinase can be validated by injecting the virus into a suitable Cre-reporter mouse line (e.g., B6.Cg-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J, RRID:IMSR_JAX:007909) with Cre-dependent expression of a fluorescent protein. To quantify the effectivity and time course of Cre-mediated protein ablation, mRNA or protein levels should be measured with qRT-PCR, Western plot, or immunohistochemistry in either cell cultures or brain slices.
General notes and troubleshooting
General notes
All surgical procedures should be carried out in accordance with a protocol approved by the Institutional Animal Care and Use Committee (IACUC). Variations in anesthesia, surgical procedures, pain management, and post-operative care and follow-up may be necessary. Consult with IACUC before performing any procedure and make sure to use only methods that minimize the animal’s pain and suffering. All stereotaxic surgery instruments should be sterilized before use. All reagents are molecular biology grade and should be stored in accordance with the manufacturer's instructions. Virus solutions should be kept in aliquots (10 μL) at -80 °C for long-term storage and multiple freeze-thaw cycles should be avoided. All solutions are prepared using ultrapure water (> 18 MΩ·cm at 25 °C) unless noted otherwise.
Troubleshooting (see Table 1)
Table 1. Troubleshooting
Problem observed
Possible reason
Solution
Virus solution does not flow out of the pipette
Pipette tip clogged
Make sure mannitol is fully dissolved, remove air bubbles from injection system
Target region damaged or destroyed after injection
Too high injection pressure or injection volume
• Reduce injection speed to 100 nL/min
• Reduce injection volume to 1 μL per injection site
• After injection, leave the pipette in place for one minute to dissipate the pressure
Pipette movement to fast
Reduce pipette movement speed to 1 mm/min
Little to no EGFP expression after injection
Virus injected in ventricle or liquor
Correct injection coordinates
Virus not functional/suitable
• Validate viral expression in cell cultures
• Verify that virus/serotype can transfect target cells
Site of EGFP expression not reproducible
Animal head size varies between animals
• Use animals with similar size/weight
• Correct injection coordinates for differences in animal head size
Animals not positioned correctly in stereotaxic frame
verify correct positioning of the animal’s head; small deviations between animals can have huge effect on injection coordinates
Acknowledgments
We thank the members of the Young lab for their comments on the manuscript. This protocol was derived from original research papers (Lübbert et al., 2019; Keine et al., 2022). The work has been supported by grants from NIDCD (R01 DC014093), NINDS (R01 NS110742) and NCATS R03TR004161-0) to S.M.Y. and a postdoctoral fellowship from the German Research Foundation (DFG, 420075000) to C.K.
Competing interests
The authors declare no competing interests. The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
Ethics considerations
All experiments were performed following animal welfare laws and approved by the Institutional Committee for Care and Use of Animals at the University of Iowa PHS Assurance No. D16- 00009 (A3021- 01) (Animal Protocol 0021952) and complied with accepted ethical best practices.
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The Effects of Whole-body Cold-water Immersion on Brain Connectivity Related to the Affective State in Adults Using fMRI: A Protocol of a Pre-post Experimental Design
AY Ala Yankouskaya *
HM Heather Massey *
JT John James Totman *
LL Lin Hui Lai
RW Ruth Williamson *
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4794 Views: 545
Reviewed by: Oneil Girish Bhalala Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Biology (Basel) Jan 2023
Abstract
An emerging body of behavioural studies indicates that regular swimming in cold water has positive effects on mental health and wellbeing, such as reducing fatigue, improving mood, and lessening depressive symptoms. Moreover, some studies reported immediate effects of cold-water immersion (CWI) on elevating mood and increasing a positive emotional state. However, the neural mechanisms underlying these effects remain largely unknown. The lack of studies using neuroimaging techniques to investigate how a whole-body CWI affects neural processes has partly resulted from the lack of a tested experimental protocol. Previous protocols administered tonic limb cooling (1–10 °C) while recording functional magnetic resonance (fMRI) signals. However, using very low water temperature constitutes points of contrast to painful experiences that are different from what we experience after a whole-body head-out CWI. In our protocol, healthy adults unhabituated to cold water were scanned twice: immediately before (pre-CWI) and after (post-CWI) immersion in cold water (water temperature 20 °C) for 5 min. We recorded cardiac and ventilatory responses to CWI and assessed self-reported changes in positive and negative affects. Our protocol showed reliable changes in brain connectivity after a short exposure to cold water, thus enabling its use as a tested experimental framework in future studies.
Graphical overview
Keywords: Whole-body cold-water immersion Resting state fMRI Functional connectivity Short exposure to cold water
Background
Cold-water swimming has become increasingly popular (Griffiths and Turner, 2022). With much evidence suggesting this activity has risks (Tipton, 1989; Tipton et al., 2017; Knechtle et al., 2020), the beneficial effects of cold-water immersion (CWI) have been linked to enhancing a positive mood state (Massey et al., 2022), higher perceived psycho-physical wellbeing (Huttunen, et al., 2004; Demori et al., 2021), and ameliorating mood disorders (Lindeman et al., 2002). It has been suggested that whole-body exposure to cold triggers a release of neurotransmitters such as serotonin, cortisol, dopamine, norepinephrine, and β-endorphin (Hirvonen et al., 2002), which play a crucial role in emotion (Gu et al., 2016; Canli and Lesch, 2007) and stress regulation (Godoy et al., 2018). Furthermore, emerging evidence indicates that the therapeutic benefits of cold water, such as increasing positive and decreasing negative affects, may be gained even from a single exposure (Massey et al., 2020; Kelly and Bird, 2022). This evidence was derived from behavioural studies using a questionnaire design and, as such, the neural mechanisms of the effects of CWI on affective processes remain largely unknown.
Only a few studies investigating how CWI changes brain activity have been conducted in humans (Jarrahi et al., 2017; Bitar et al., 2020; Grouper et al., 2022). However, these studies focused on assessing brain activity and its association to cognitive-affective aspects of pain, by using an experimental protocol of administered tonic limb cooling. It must be noted that physiological responses to limb cooling using very low water temperatures (1–10 °C) constitute points of contrast that result in painful experiences, which are different from what we experience after taking a cold shower or swimming in higher temperature open waters.
We developed and tested a novel protocol to examine the effects of short-term, head-out, whole-body CWI on changes in brain functional connectivity. This protocol can be adapted for other research questions focusing on brain-behaviour interactions [e.g., linking personality traits, cognitive functions such as decision making, memory, and perception to the brain's functional architecture (Cole et al., 2014)]. A tight control of experimental factors in our protocol makes it easy to replicate and minimise confounding effects.
Materials and reagents
Non-effervescent Haz-Tab (4.5 g NADDC per tablet) (Guest Medical, catalog number: H8801) made up to 1,000 ppm (one tablet per 2.5 L)
The Positive and Negative Affect Schedule (PANAS) (Watson et al., 1988)
Appropriately sized hospital scrubs for each participant
Blankets
Equipment
MRI machine (3-Tesla Siemens Magnetom Lumina, 32-channels head-coil)
Water heater (Laz-y-spa, UK)
Water pump (Laz-y-spa, UK)
Water chiller unit (Grant Instruments, UK)
Three-lead electrocardiogram (ECG, Fukuda Denshi, UK)
ECG electrodes (Ambu® Blue Sensor P, Malaysia)
Electrodes covered with Tegaderm dressings (3M, Germany)
35 mm respiratory hose (Cranlea, UK)
Reusable nose clip (Cranlea, UK)
Two-way T-shape non-rebreathing valve (Hans Rudolf, US)
Respiratory turbine (K.L. Engineering, US)
Thermistor pod (ML309, AD Instruments, Australia)
Skin temperature thermistor (MLT422/A, Australia)
Digital recorder (Powerlab, AD Instruments, Australia)
Laptop connected to the Powerlab and programmed to a sampling rate of 400 Hz (Chart software, AD Instruments Australia)
Scanning parameters
Anatomical scan: T1-weighted MPRAGE images with the following parameters: repetition time (TR) = 1,900 ms, echo time (TE) = 2.74 ms, flip angle = 8°, field of view (FOV) = 256 × 256 mm2, voxel size = 1.0 × 1.0 × 1.0 mm3, and 192 axial slices.
Functional scan: blood oxygenation level–dependent (BOLD) contrast whole-brain functional images were acquired using a T2-weighted gradient-echo Echo Planar Imaging (EPI) sequence with a 32-channel head coil. Sequence duration 13.12 min. Acquisition parameters: TR = 2,680 ms; TE = 30 ms; matrix size = 64 × 64 mm2; slice thickness = 2.5 mm, spacing between slices = 2.5 mm, flip angle = 80°; 292 volumes with 48 axial slices were measured in interleaved slice order and positioned along a line to the anterior-posterior commissure (AC-PC orientation). An automated high-order shimming technique was used to maximise magnetic field homogeneity.
Software
MATLAB 2022a (MATLAB and Statistics Toolbox Release 2012b, The MathWorks, Natick, Massachusetts, United States)
CONN (v.2021a): A functional connectivity toolbox for correlated and anticorrelated brain networks (Whitfield-Gabrieli and Nieto-Castanon, 2012, http://www.nitrc.org/projects/conn)
JASP (JASP Team, 2022, version 0.16.3) (Computer software)
Procedure
Step 1. Participant screening
Eligibility screening
Provide interested volunteers with a participant information sheet containing sections explaining eligibility criteria, procedures, and risks.
Eligibility criteria: (i) age is between 18 and 45 years old; (ii) no history of medical, neurological, psychiatric, or substance use disorders; (iii) naïve to CWI (not exposed to cold water including cold shower, unheated swimming pool, or the sea) in the last 12–18 months, and (iv) free from chronic pain of any type (Figure 1).
Figure 1. Participants selection consort diagram. The diagram provides information about the selection steps (on the left) and the number of participants excluded in each selection step.
If required, screen participants by telephone interview and undertake any additional pre-screening tests required by the medical officer to rule out any significant history of medical, neurological, psychiatric, or substance use disorders.
Claustrophobia screening
Siemens Lumina scanner has a relatively large tunnel, is fully lit, and is open at both ends. This dramatically lowers the experience of claustrophobia. Ask participants whether they have ever experienced claustrophobia (or fear of being in a closed space) in the past.
Ask participants whether they have previous experience with MRI scans.
Ask participants whether exposure to a feared situation have led to immediate anxiety or panic attacks in the past.
If the answer is yes to a. or c., the participant does not pass claustrophobia screening.
MRI compatibility screening
MRI compatibility screening should comply with the standard operation procedures approved for scanner use. The procedures can be slightly different across universities. Discuss the screening procedures with your local scanner manager.
Ask participants to answer questions assessing their eligibility for fMRI study: (i) Do you have a heart pacemaker or pacing wires? (ii) Have you had any recent surgery? (iii) Have you had any surgery to your head (including eyes/ears/brain)? (iv) Do you have any implanted devices (e.g., programmable hydrocephalus shunt, nerve stimulator, cochlea implant, aneurysm clip? (v) Have you ever sustained any injuries involving metal to your eyes or any parts of your body? (vi) Do you have any of the following: dentures with metal, body piercings, hearing aid, nitro patch, artificial limb or prothesis, tattoos? (vii) For women: do you have an IUD (coil)? Could you be pregnant?
If the answer to all questions is no, the participant will be taken to the next screening step. If the answer to one or several questions is yes, take this case to a radiographer.
Participants who passed Step 1 were scheduled to visit the Institute of Medical Imaging and Visualisation (IMIV) at Bournemouth University where the study took place. For the scheduled visit, ask participants to (i) refrain from eating food 2 h prior to the scanning session; (ii) bring a swimsuit/trunks or shorts (+vest top for women), a towel, a woollen jumper or sweatshirt with no metal, and slippers or flip flops.
Step 2. Pre-CWI assessments
MRI safety assessment
If the participant passes the MRI safety assessment, explain the experimental procedures and familiarise participants with the setup, including cold-water tub and monitoring equipment.
Answer any questions the participant may have and ask them to complete a written informed consent form.
Pre-CWI mood assessment
Provide a printed version of the PANAS questionnaire and instruct them to evaluate all items of the questionnaire, reflecting on how they feel as they complete it (in the moment).
Ask participants if they have any questions. Ensure that their clothes do not have any metal parts (or provide a set of scrubs and blankets).
Step 3. Experimental procedures
Pre-CWI scan
Position a participant in the scanner (Figure 2). Use MRI-compatible headphones to reduce scanner noise. Use pads to fixate the participant’s head to reduce head movements.
Figure 2. Participant’s position in the scanner
Provide an alarm button (by pressing the button, participants are able to stop the scanning at any time).
Ask participants to keep awake and keep their eyes open.
Start anatomical (structural) scan (4.5 min). Immediately after, start the functional scan (resting state) (13.12 min).
After completing the pre-CWI scanning procedures, take the participant to a changing room to change into swimwear.
CWI (cold-water immersion procedure)
Record participant’s height and mass.
Instrument with a three-lead ECG connected via ECG electrodes and waterproof the electrodes using Tegaderm dressings to improve the signal quality during immersion.
Provide the participants with a nose clip and mouthpiece and ask them to breathe freely throughout.
A minimum of 2 min of baseline data are collected on the Chart software with the participant seated next to the cold-water tank. The Chart program also collects water temperature, ECG, and ventilatory data throughout the immersion, and comments should be added to the chart software to highlight pre immersion, immersion, and post immersion phases.
Participant will stand, immediately enter the tank, and be seated in the water within a 10–15 s period. They will immerse to the depth of the axilla (see image in Graphical overview). If the immersion is too deep, ask them to sit at the shallow part; if it is too shallow, ask them to sit in a deeper part. This is done by putting weights on the bottom, which the participants could sit on.
Participants are encouraged to breathe freely through the mouthpiece and do not attempt to hold their breath.
Monitor the ECG during immersion for any ECG abnormalities that would necessitate withdrawal from the cold water.
Upon the completion of 5 min of immersion or if the participant requires earlier withdrawal, they stand up, exit the water tank onto a skid resistance surface, and the ECG and mouthpiece are removed.
Participants remove their swimwear and dry off and dress in private, as quickly as possible.
Record the duration of time from exiting the cold-water tub to the MRI chamber door closing, following the positioning of the participant in the scanner. The average time between exiting the cold-water tub and entering the scanner was M = 5.82 min, SD = 2.08 min (range: 2 min 25 s–10 min 30 s). It is not anticipated that the transition time between the tub and the scanner dramatically affects changes in the brain, as previous research reported the effects of CWI to last over a few days (Massey et al., 2022). However, we encouraged participants to dry off and dress quickly after the CWI. It must be noted that the cold-water tub should be set up as close to the MRI scanner as possible, to ensure the shortest transition time. In our study, the cold-water tub was next door to the scanner room.
Post-CWI scan
Position the participant in the scanner (Figure 2).
Use a lightweight cotton blanket to cover the participant’s body (to minimise shivering after CWI).
Switch off the built-in scanner ventilator (to minimise participant cooling and body movement due to cold air).
Immediately after, start the functional scan (resting state) (13.12 min).
Table 1 summarises the timeline for the experimental steps to provide a detailed estimation of the complete study per participant at the MRU suite.
Table 1. Detailed timeline for experimental steps B and C at the MRI suite
Steps Sub steps Additional time between sub steps Time
B MRI safety assessment (including explanation of CWI procedure and signing consent form) 5 min
Pre-CWI mood assessment 2–3 min
Changing into MRI-compatible clothes (if required) 1–3 min
C Position participant in the scanner (pre-CWI) 3 min
Pre-CWI scanning 4.5 min + 13.12 min = 17.62 min
Localise scan to position the brain 1 min
Get participant from the scanner, change into swimwear 3–4 min
Instrument with ECG electrodes, nose clip, and mouthpiece 2–3 min
Taking baseline measurements 2–3 min
Cold-water immersion 5 min
Remove ECG, nose clip, and mouthpiece 1 min
Drying and changing into MRI-compatible clothes M = 5.83, SD = 2.09*
Position participant in the scanner (post-CWI) 2–3 min
Post-CWI scanning 13.12 min
Localise scan to position the brain 1 min
Total time: ~62 min**
*This parameter ranged between 2 min 25 s and 11 min 15 s.
**Total time can be reduced if participants arrive in MRI-suitable clothes.
Data analysis
The original paper provides detailed data analysis for this study. The analytical steps are summarised in Figure 3.
Figure 3. Analytical steps in the present study
Tips
To disinfect the mouthpieces, nose clips, and respiratory hose, two non-effervescent Haz-Tab were dissolved into 5 L of cold tap water. The tablet was allowed to dissolve completely before the equipment was dismantled and submerged in the liquid for 20 min.
The CWI tub does not need to be drained or disinfected between participants. It was checked twice a day for pH and free and combined chlorine and was dosed with chlorine daily in accordance with the manufacturer’s guidelines. The tub also had filters for trapping hair and detritus, which were cleaned daily.
Validation of protocol
Yankouskaya et al. (2023). Short-term head-out whole-body cold-water immersion facilitates positive affect and increases interaction between large-scale brain networks. Biology, 12(2): 211.
Acknowledgments
We wish to thank the participants and radiography team who are not authors (Carlo Vitale and Theophilus Akudjedu). This work was supported by the Institute of Medical Imaging and Visualisation (Bournemouth University, UK).
Competing interests
None of the authors has any conflicts of interest.
Ethics
The protocol requires approval from a local ethics committee. The study was reviewed and approved by the Departmental Research Board and Ethics Committee at Bournemouth University (Ethics ID 34976 17.02.2021) and conducted in accordance with the Declaration of Helsinki. Written informed consent from each participant was obtained before the study.
References
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Demori, I., Piccinno, T., Saverino, D., Luzzo, E., Ottoboni, S., Serpico, D., Chiera, M. and Giuria, R. (2021). Effects of winter sea bathing on psychoneuroendocrinoimmunological parameters. Explore 17(2): 122–126.
Godoy, L. D., Rossignoli, M. T., Delfino-Pereira, P., Garcia-Cairasco, N. and de Lima Umeoka, E. H. (2018). A Comprehensive Overview on Stress Neurobiology: Basic Concepts and Clinical Implications. Front. Behav. Neurosci. 12: e00127.
Griffiths, S. and Turner, Y. (2022). Trends in Outdoor Swimming. Second Edition. 2022. Available online: https://www.outdoorswimmershop.com/products/outdoor-swimmer-trends-report-22 (accessed on 18 December 2022).
Grouper, H., Löffler, M., Flor, H., Eisenberg, E. and Pud, D. (2022). Increased functional connectivity between limbic brain areas in healthy individuals with high versus low sensitivity to cold pain: A resting state fMRI study. PLoS One 17(4): e0267170.
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Hirvonen, J., Lindeman, S., Joukamaa, M. and Huttunen, P. (2002). Plasma catecholamines, serotonin and their metabolites and beta-endorphin of winter swimmers during one winter. Possible correlations to psychological traits. Int. J. Circumpolar Health 61(4): 363–372.
Huttunen, P., Kokko, L. and Ylijukuri, V. (2004). Winter swimming improves general well-being. Int. J. Circumpolar Health 63(2): 140–144.
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Kelly, J. S. and Bird, E. (2022). Improved mood following a single immersion in cold water. Lifestyle Medicine 3(1): e53.
Knechtle, B., Waśkiewicz, Z., Sousa, C. V., Hill, L. and Nikolaidis, P. T. (2020). Cold Water Swimming—Benefits and Risks: A Narrative Review. Int. J. Environ. Res. Public Health 17(23): 8984.
Lindeman, S., Hirvonen, J. and Joukamaa, M. (2002). Neurotic psychopathology and alexithymia among winter swimmers and controls--a prospective study. Int J Circumpolar Health.61(2): 123–130.
Massey, H., Gorczynski, P., Harper, C. M., Sansom, L., McEwan, K., Yankouskaya, A. and Denton, H. (2022). Perceived Impact of Outdoor Swimming on Health: Web-Based Survey. Interact. J. Med. Res. 11(1): e25589.
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4,795 | https://bio-protocol.org/en/bpdetail?id=4795&type=0 | # Bio-Protocol Content
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Peer-reviewed
Use of Open Surface Plasmon Resonance (OpenSPR) to Characterize the Binding Affinity of Protein–Protein Interactions
CZ Cassie Shu Zhu
JL Jianhua Li
HW Haichao Wang
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4795 Views: 1038
Reviewed by: Emilia KrypotouGregory CravenSrajan KapoorQingliang Shen
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Original Research Article:
The authors used this protocol in Science Advances Feb 2023
Abstract
Surface Plasmon Resonance (SPR) is a label-free optical technique to assess protein–protein interaction kinetics and affinities in a real-time setting. Traditionally, Biacore SPR employs a continuous film of gold to detect any change in the angle of re-emitted light when the refractive index of a ligand conjugated to the flat gold surface is altered by its interaction with a local analyte. In contrast, the Nicoya Lifesciences’ OpenSPR technology uses gold nanoparticles to detect small changes in the absorbance peak wavelength of a conjugated ligand after its engagement by an analyte. Specifically, when broadband white light is shone onto the gold nanoparticles, it produces a strong resonance absorbance peak corresponding to the refractive index of a ligand conjugated to the surface of gold nanoparticles. Upon its interaction with an analyte, however, the absorbance wavelength peak of the conjugated ligand will be changed and timely recorded as sensorgrams of dynamic ligand–analyte interactions. Thus, the improvement in the detection method (from traditional detection of changes in the angle of re-emitted light to the contemporary detection of changes in the wavelength of the absorbance peak) features OpenSPR as a cost-effective and user-friendly technique for in-depth characterization of protein–protein interactions. Here, we describe the detailed method that we used to characterize procathepsin L (pCTS-L) interactions with two putative pattern recognition receptors (TLR4 and RAGE) using the 1st generation of Nicoya Lifesciences’ OpenSPR instrument with a 1-channel detection.
Key features
• Nicoya OpenSPR is a benchtop small-size equipment that provides in-depth label-free binding kinetics and affinity measurement for protein–protein interactions in real-time fashion.
• This technology is relatively intuitive and user-friendly for scientists at any skill level.
• OpenSPR sensors employ nanotechnology to reduce the cost of manufacturing complex optical hardware and Sensor Chips, and similarly reduce the consumption of precious analyte samples.
• The manufacturer provides online training for OpenSPR (Catalog: TRAIN-REMOTE) and TraceDrawer (Catalog: TRAIN-TD) to customer scientists.
Keywords: Procathepsin L (pCTS-L) Toll-like Receptor 4 (TLR4) Receptor for Advanced Glycation End Products (RAGE) OpenSPR NTA Sensor Chip
Background
In order to understand complex mechanisms underlying the regulation of various biological processes, it is often essential to characterize protein–protein interactions under many physiological or pathological conditions. For many years, surface plasmon resonance (SPR) has been widely used to analyze all types of interactions between protein–protein or protein and other molecules such as nucleic acids, lipids, and carbohydrates (Willander and Al-Hilli, 2009; Drescher et al., 2018). Traditionally, Biacore SPR uses a continuous film of gold to detect changes in the angle of re-emitted light when the refractive index of a ligand conjugated to the flat gold surface is altered by its interaction with a local analyte (Willander and Al-Hilli, 2009; Drescher et al., 2018). In contrast, the newly developed Nicoya Lifesciences’ OpenSPR employs gold nanoparticles to detect small changes in the absorbance peak wavelength of a conjugated ligand after its binding to an analyte. Nevertheless, both traditional SPR and contemporary OpenSPR can give real-time information about the binding kinetics and affinities of dynamic interactions between various molecules.
During the last few years, we have used Nicoya Lifesciences’ first generation 1-channel OpenSPR to determine the binding affinities between antigens and antibodies (Chen et al., 2020) or cytokines and putative receptors (Qiang et al., 2022; Zhu et al., 2023). Briefly, a ligand protein is immobilized onto nitrilotriacetic acid (NTA)-conjugated gold nanoparticles in a specific orientation, and an analyte solution is subsequently injected at several increasing concentrations into the microflow system. When broadband white light is shone onto the gold nanoparticles, it produces a strong resonance absorbance peak that is specific to the refractive index of a local ligand conjugated to the gold nanoparticles. If an analyte binds the ligand, it will induce a change in the wavelength of the absorbance peak of the ligand, which can be recorded in the sensorgram as an increase in the SPR signal. After a desired association time, a solution without the analyte is injected to dissociate the complex between ligand and analyte. If the analyte dissociates from the ligand, a decrease in SPR signal would be observed. By dividing the dissociation rate (koff) by the association rate (kon), the equilibrium dissociation constant (KD) can be calculated as a measure of the affinity between these specific ligand–analyte interactants. Thus, the improvement in the detection method (from traditional detection of changes in the angle of re-emitted light to the contemporary detection of changes in the wavelength of the absorbance peak) features OpenSPR as a cost-effective and user-friendly technique for in-depth characterization of protein–protein interactions. Here, we describe the detailed method that we used to characterize procathepsin L (pCTS-L) interaction with two putative pattern recognition receptors (TLR4 and RAGE) using Nicoya Lifesciences’ 1st generation OpenSPR instrument with a 1-channel detection (Zhu et al., 2023).
Materials and reagents
Reagents
Extracellular domain of human TLR4 (residue 1-631, 70.5 kDa) (Sino Biological, catalog number: 10146-H08B)
Extracellular domain of human RAGE (residue 1-344, 36.0 kDa) (Sino Biological, catalog number: 11629-H08H)
Human pCTS-L corresponding to residue 17-333 of respective procathepsin L carrying a N-terminal 6× His tag was expressed in E. coli BL21 (DE3) pLysS cells and purified as previously described (Zhu et al., 2023).
Solutions
Phosphate buffered saline (PBS) containing 10 mM Na2HPO4, 10 mM NaH2PO4, 140 mM NaCl, and 3 mM KCl at pH 7.4 (Nicoya Lifesciences, Kitchener, catalog number: NI-PBS)
HBS-T running buffer containing 0.01 M HEPES, 0.15 M NaCl, and 0.005% Tween-20 at pH 7.4 (Nicoya Lifesciences, catalog number: HBS-T)
NiCl2 solution (40 mM, store at 2–8 °C) and imidazole solution (200 mM, store at 2–8 °C) prepared using the NTA Reagent Kit (Nicoya Lifesciences, catalog number: NTA-RK)
Deionized water filtered through 0.2 μm vacuum filter bottle system (Corning, catalog number: 431097)
80% isopropanol (v/v in deionized water, prepared from isopropanol) (Sigma-Aldrich, catalog number: 563935)
Laboratory supplies
Buffer bottles (Nicoya Lifesciences, catalog number: BTL-SQ-250)
Tweezers (included in the standard supplies of OpenSPR equipment)
Lint-free wipes (Fisher Scientific, catalog number: S47299)
Disposable syringes (Nicoya Lifesciences, catalog number: SYR-PL-50)
Gastight glass syringes (Nicoya Lifesciences, catalog number: SYR-G)
Blunt-end injection tips (Nicoya Lifesciences, catalog number: TIP-BLUNT-50)
OpenSPR Nitrilotriacetic Acid (NTA) Sensor Chip (Nicoya Lifesciences, catalog number: SEN-Au-100-10-NTA)
NTA Reagent Kit for making 40 mM NiCl2 solution and 200 mM imidazole solution (Nicoya Lifesciences, catalog number: NTA-RK)
Nitrile or Latex gloves
Declogging Kit (Nicoya Lifesciences, catalog number: DECLOG OpenSPR)
Equipment
Nicoya Lifesciences’ 1st generation of gold-nanoparticle-based Open Surface Plasmon Resonance (OpenSPR) 1-Channel Instrument (Kitchener, catalog number: REV 3.0, Figure 1).
Figure 1. Photos of an OpenSPR 1-Channel Instrument and a laptop computer with installed Nicoya Lifesciences software
Note: The OpenSPR is operated by the user manually, using a laptop computer with an Experiment Data Interface that allows the user to control various aspects of the OpenSPR instrument, such as changing running buffer, fluid flow rate, as well as sample injection. For instance, when a sample is manually injected into the Sample Loop via the Injection Port, the user then needs to switch the Injection Valve clockwise to the Inject position to initiate the delivery of the injected samples from the Sample Loop to the sensor.
Laptop computer with installed Nicoya Lifesciences software (Experiment Data Interface) for control of the instrument and fast real-time data acquisition (Figure 1).
Software and datasets
TraceDrawer Kinetic Data Analysis Software V.1.6.1 (Nicoya Lifesciences) for data processing and analysis of kinetic and affinity constants.
Procedure
Set up the OpenSPR instrument and software according to the manufacturer’s detailed instructions in the OpenSPR manual, which includes the connection of the OpenSPR instrument to the computer with a USB cable and the fluidic setup of three bottles (for buffers and waste).
Open the Experiment Data Interface on the computer, and click Start to proceed to the preparation of the instrument by:
Indicating your running buffer and priming the fluidics of the OpenSPR by filling the pumps and tubing with the indicated running buffers (HBS-T, pH 7.4) at a flow rate of 150 μL/min for 2.5 min to clear out any contamination or bubbles out of the fluidic lines prior to your test.
Taking a new optical reference spectrum without a Sensor Chip in the instrument to calculate the sensor absorbance signal before loading a new Sensor Chip to start a new test.
Loading a new NTA Sensor Chip prior to the start of a test:
i. Clean the face of the Flow Cell with a lint-free wipe soaked in 80% isopropanol and allow it to air dry completely before installing a new Sensor Chip.
ii. Rinse the Sensor Chip thoroughly with distilled water and allow it to air dry completely to prevent leaking and bubble formation in the Flow Cell.
iii. Carefully slide the Sensor Chip into the Sensor Holder in the correct orientation and place the Sensor Holder into the instrument by first aligning it with the Fluidics Block, then slowly bring it into contact.
Prepare the test by:
Switching the Injection Valve to the Load position and preparing for sample injection by:
i. Slowly rinsing the Injection Port and Sample Loop with at least 1.0 mL of running buffer (HBS-T, pH 7.4) to flush any previous sample/solution out of the Sample Loop fluidic lines.
ii. Purging the Sample Loop using the same buffer syringe filled with at least 1.0 mL of air to push the air through the Sample Loop, to remove excess buffer in the loop via the outlet line.
Removing bubbles within the fluidic lines.
i. Set the pump speed at 150 μL/min and turn the Injection Valve counterclockwise to the Load position.
ii. Use a syringe with a blunt-end tip to inject 300 μL of 80% isopropanol via the Inject Port into the Sample Loop, and then turn the Inject Valve clockwise to the Inject position to deliver isopropanol to the sensor.
iii. Turn the Injection Valve counterclockwise back to the Load position.
Immobilize the 6× His Tag-containing pCTS-L ligand onto the NTA sensor according to instructions.
Surface conditioning: set the pump speed to 150 μL/mL and use disposable syringes to fill the 100 μL Sample Loop with 200 μL of imidazole solution (200 mM), and then turn the Injection Valve to the Inject position to initiate delivery of the injected imidazole solution from the Sample Loop to the sensor to clean the sensor surface. After 5 min, turn the Injection Valve counterclockwise back to the Load position.
Rinse the Inject Port and Sample Loop with at least 1.0 mL of running buffer (HBS-T, pH 7.4) and purge with at least 1.0 mL of air as described in step 3a before the next injection.
Surface activation: set the pump speed to 20 μL/mL and use disposable syringes to fill the Sample Loop with 200 μL of NiCl2 solution (40 mM), and then switch the Injection Valve to the Inject position to initiate delivery of the injected NiCl2 solution from the Sample Loop to the sensor to activate the NTA sensor surface. Leave the Inject Valve in the Inject position for the entire duration of the activation time until a stable baseline is obtained, and then turn the Injection Valve counterclockwise back to the Load position.
Rinse the Inject Port and Sample Loop with at least 1.0 mL of running buffer (HBS-T, pH 7.4), and purge with at least 1.0 mL of air as described in step 3a.
Switch the running buffer from HBS-T to 1× PBS by:
i. Clicking the Stop button in the Pump Menu of the Experiment Data Interface to stop the pump.
ii. Switching the inlet tubing from the bottle containing HBS-T to the bottle containing 1× PBS (pH 7.4).
iii. Resuming the pump by clicking the Start button in the Pump Menu.
Set the pump speed to 20 μL/mL and use disposable syringes with gastight blunt-end tips (TIP-BLUNT-50) to inject 200 μL of pCTS-L solution diluted in the running buffer (1× PBS, pH 7.4) to a final concentration of 50 μg/mL. Quickly turn the Injection Valve clockwise to the Inject position to initiate the delivery of the injected pCTS-L ligand solution from the Sample Loop to the NTA sensor. Leave the Injection Valve on the Inject position for the entire duration of the ligand interaction with the sensor surface. A representative sensorgram for immobilizing pCTS-L ligand onto an NTA Sensor Chip is shown below (Figure 2).
Figure 2. Sensorgram of pCTS-L Ligand Immobilization
Note: For most protein ligands, the recommended ligand concentrations for immobilization are typically 10–50 μg/mL. We chose a higher concentration (50 μg/mL) of pCTS-L ligand to ensure its maximal immobilization to most active sites of the NTA sensor. The amount of ligand immobilized onto the NTA Sensor Chip can be determined by the subsequent increase in the response signal in corresponding sensorgrams. Once the immobilization signal has reached plateau levels, it indicates a state of maximal ligand immobilization. However, the user can repeat the immobilization procedure one more time to ensure there will be no additional further increase in the response signal (i.e., the “net signal shift”) corresponding to additional ligand immobilization, so that most (if not all) NTA open sites are occupied by the ligand to prevent non-specific interaction with analytes containing exposed histidine residues.
When an injection is made, an Injection Details box will appear for the user to input the ligand name (pCTS-L) as well as the concentration (50 μg/mL), which will be used to identify the injection in the data analysis.
Note: For all injections, other than the Bubble Removal Injection and Analyte Injections, leave the Injection Valve in the Inject position for at least the entire duration of the interaction time of your sample with the sensor surface. When ready to perform the next injection, turn the Injection Valve back to the Load position, and rinse the Injection Port and Sample Loop with running buffer and purge the Sample Loop with air as described in step 3a.
Once the immobilization signal has reached plateau levels, it indicates a state of maximal ligand immobilization. Allow the instrument to stabilize with buffer running until the slope of the baseline is < 5 pM/min, which may take anywhere from 5 min to 1 h.
Rinse the Inject Port and Sample Loop with running buffer (1× PBS, pH 7.4), and purge with air as described in step 3a.
Prepare TLR4 analyte solution in the analysis running buffer (1× PBS, pH 7.4) at 3-fold serial dilutions (i.e., 283.7 nM, 94.6 nM, and 31.5 nM), and inject at least 200 μL of analyte solution at a flow rate of 20 μL/min in the order of increasing concentrations with an association time of 240 s and a dissociation time of 480 s.
Load syringe with 200 μL of TLR4 analyte solution at a concentration of 31.5 nM, insert it fully into the Injection Port, and then slowly depress the plunger to introduce the analyte into the instrument. Wait at least 5 s to stabilize the syringe fluidic pressure in the Sample Loop before removing the syringe from the Injection Port.
Turn the Injection Valve clockwise to the Inject position to initiate the delivery of the injected TLR4 analyte solution from the Sample Loop to the NTA sensor and wait for 240 s to ensure maximal interaction between ligand and analyte.
When the Injection Remaining counter reaches 00:00:00, quickly turn the Injection Valve back to the Load position, and wait for 480 s to allow maximal dissociation between ligand and analyte.
When the injection is complete and the baseline has settled, the user can move on to the next round of injection of TLR4 analyte at 94.6 nM and 283.7 nM, respectively, by repeating steps 6 and 7.
Note: When an injection is made, an Injection Details box will appear to let the user input the analyte sample name (e.g., TLR4) as well as the concentration (e.g., 31.5 nM), so the software can identify this injection in subsequent data analysis. Before each analyte injection, the Injection Port and Sample Loop must be rinsed with analysis running buffer (e.g., 1× PBS, pH 7.4) and purged with air as described in step 3a to prevent cross-contamination between injections.
To complete an experiment, click Finish to stop tracking the absorbance peak and create the final data files.
For the characterization of the pCTS-L-RAGE interaction, load a new NTA Sensor Chip and prepare the solution of RAGE analyte in the analysis running buffer (1× PBS, pH 7.4) at 3-fold serial dilutions (i.e., 250 nM, 83.3 nM, and 27.7 nM). Respectively, inject the Sample Loop with 200 μL of RAGE analyte solution at a flow rate of 20 μL/min in the order of increasing concentrations with an association time of 240 s and a dissociation time of 480 s following similar procedures as described in steps 2–8.
Data Analysis: Once the user finishes the test, the software will create several types of data files that will always be saved under Documents\OpenSPR\TestResults\ along with the name given by the user at the beginning of the test.
Open TraceDrawer Software and click Add run to select and import saved files and relevant curves (i.e., TLR4 or RAGE) for analysis.
Create an overlay by clicking New Overlay and then left click and drag a run to an overlay to perform data modifications in the Overlay windows.
On the left side of the screen, click New Evaluation to set the Evaluation Type to Kinetics Evaluation.
Define two timepoints where a concentration change occurs in the data, i.e., the beginning and end of the association phase.
In the Kinetics Evaluation table that displays a row with the selected timepoint, define the concentrations of each curve, and press Next.
In Fit model dropdown list, choose OneToOne model to initiate a 1:1 diffusion-corrected binding model with global fitting, then press Fit.
For more comprehensive instructions on the TraceDrawer Software, please refer to the manual.
Note: The user can zero the graph by clearing all the data points from the graph and normalizing the response tracking to zero on the y-axis at any stage in the experiment where a new baseline is established. For instance, the user can zero the graph to make a relative comparison for subsequent injections at any stage after the ligand immobilization step and before injection of the analyte for analysis. The user can also open advanced settings to manually adjust the axes scale by clicking the gear (settings) button in the graph menu and setting custom axes before saving an image of the graph.
Shutdown Procedure
If the user does not plan to use the instrument in the near future, it is recommended to perform a full shutdown before rinsing all fluidic lines thoroughly with at least 20 mL of deionized water and 20 mL of 80% isopropanol at the maximum speed (150 μL/min), followed by purging the lines with a sufficient amount of air to dry out the tubing lines. For information regarding other rinsing procedures and maintenance, please refer to the manual of the instrument.
Data analysis
For the OpenSPR analysis of pCTS-L interactions with TLR4 or RAGE, pCTS-L is immobilized as a ligand on a NTA sensor, and TLR4 or RAGE are applied as an analyte at increasing concentrations. The dynamic changes of signal response are recorded in the sensorgrams, which include a baseline phase of the buffer response, an association phase during which the analyte is passed over the immobilized ligand, and a dissociation phase where buffer flow resumes, and the bound analyte is removed from the sensor surface (Figures 3 and 4).
Figure 3. Sensorgrams of pCTS-L–TLR4 interactions. TLR4 was injected for a contact time of 240 s at increasing concentrations of 31.5 nM, 94.6 nM, and 283 nM, and dissociation was monitored for 480 s. The 1:1 models fit to the raw data are shown as solid black lines.
Figure 4. Sensorgrams of pCTS-L–RAGE interactions. RAGE was injected for a contact time of 240 s at increasing concentrations of 27.7 nM, 83.3 nM, and 250 nM, and dissociation was monitored for 480 s. The 1:1 models fit to the raw data are shown as solid black lines.
During the association phase, the sensorgrams reveal dynamic changes in the absorbance peak wavelength between the running buffer and the analyte in proportion to the analyte’s concentrations, and always exhibit some type of curvature as an indicator of exponential increase (Figures 3 and 4). When the sensorgram curve levels out, it indicates a state of equilibrium when the number of associations equals the number of dissociations. However, when the analyte is injected at a higher concentration, a new equilibrium is reached with a higher response (Figures 3 and 4), until all the ligand binding sites are occupied, and the maximal response is reached. Therefore, the concentration range of the injected analyte is important, because higher concentrations of analyte tend to produce curves in the upper part of the sensorgram, whereas lower concentrations will give low responses and little curvature. It is recommended to inject an analyte at 3–5 concentrations at a serial dilution in the range of 0.1–10-fold of the “equilibrium dissociation constant” (KD, i.e., 0.1 × KD to 10 × KD), which will space the curves evenly over the sensorgram with both low and high responses. When the analyte injection ends, it will enter a dissociation phase, during which the curve of dissociation follows a single exponential decrease (Figure 3) but can sometimes be almost horizontal in the case of a strong interaction (Figure 4). To analyze a slow dissociation, the dissociation period should be long enough to have at least 5% signal decrease compared with the initial response.
OpenSPR experiments can be repeated multiple times to get an average kon, koff, and KD to increase reliability and reproducibility. Representative sensorgrams for kinetic interactions between pCTS-L and TLR4 or RAGE are shown in Figure 3 and Figure 4, respectively. The overall shape of the sensorgram curves is determined by the analyte concentration, association rate (kon), and dissociation rate (koff) constants between a particular pair of ligand and analyte. Within a particular ligand–analyte pair, the overall shape of the sensorgram curves was extremely similar between repeating experiments (Figures 3 and 4), indicating some degree of reproducibility and reliability of the OpenSPR technique in assessing the kinetics and affinities of protein–protein interactions. Between different ligand–analyte pairs (Figure 3 vs. Figure 4), however, the overall shapes of corresponding sensorgram curves appeared to be markedly different, suggesting distinct and analyte-specific interactions with a particular ligand even under identical experimental conditions. Analysis of the sensorgrams using the 1:1 interaction algorithm for a single exponential interaction gave rise to an estimated kon, koff, as well as KD for each representative experiment (Figures 3 and 4). The almost 100-fold difference between the off rate (koff) of TLR4 and RAGE analyte may underlie the distinct sensorgram curves for pCTS-L interaction with TLR4 (Figure 3) and RAGE (Figure 4), respectively.
Validation of protocol
Because of our limited access to the Biacore equipment, we did not perform head-to-head comparison between OpenSPR and Biacore for pCTS-L interaction with TLR4 or RAGE. However, we did use both Biacore and OpenSPR to parallelly measure the KD between two different interactants (human dermcidin and the extracellular domain of human EGFR) (Qiang et al., 2022). For the traditional Biacore, dermcidin was immobilized on a CM5 chip as a ligand, and recombinant EGFR was applied at five different concentrations using a Biacore T200 instrument (GE Healthcare). The KD was determined using the Biacore evaluation software 2.0 supposing a 1:1 binding ratio. For the Nicoya OpenSPR, recombinant dermcidin with a 6× His tag was similarly immobilized on NTA Sensor Chips as a ligand, and recombinant EGFR was applied at three different concentrations to estimate the KD using the Trace Drawer Kinetic Data Analysis v.1.6.1. Impressively, the OpenSPR estimated a KD of 58.1 ± 29.6 nM (mean ± SEM, n = 3 technical replicates) for dermcidin–EGFR interaction, which was almost identical to the KD value (58.8 nM) estimated from using the traditional Biacore SPR technique (Qiang et al., 2022). These findings validate the reliability of using OpenSPR in assessing binding affinities between two different proteins.
When highly purified human pCTS-L was immobilized onto a NTA Sensor Chip and human TLR4 or RAGE were respectively applied as analytes at various concentrations, OpenSPR revealed similarly high affinities of pCTS-L to both TLR4 and RAGE with an estimated KD of 20.2 ± 3.5 nM and 3.5 ± 2.6 nM, respectively (Zhu et al., 2023). The calculated KD for eight independent experiments is summarized in Table 1, along with the mean and standard error of mean (SEM), which is indicative of the precision of OpenSPR for estimating a mean KD. When the SEM was expressed as a fraction of the estimated mean, the resultant Relative Standard Error (RSE) was relatively small (< 20%) particularly for pCTS-L/TLR4 interaction, validating the reliability of using OpenSPR to estimate some protein–protein binding affinities.
Table 1. Mean KD of eight OpenSPR experiments for pCTS-L interaction with TLR4 or RAGE
Experiment
#
KD for pCTS-L/TLR4 (nM) KD for pCTS-L/RAGE (nM)
1 24.1 2.89
2 20.6 2.66
3 24.9 0.59
4 19.7 0.0023
5 7.7 0.222
6 10.3 0.014
7 15.4 0.019
8 39.0 21.5
Mean ± SEM 20.2 ± 3.5 3.5 ± 2.6
Relative Standard Error (RSE) 17.3%
Given the 1:1 stoichiometry of pCTS-L–TLR4 interaction in the ClusPro protein docking (Zhu et al., 2023), we used the 1:1 fitting model for our data analysis. The somewhat imperfect fit between the 1:1 binding model and corresponding data for pCTS-L–TLR4 interaction (Figure 3) did slightly damper our confidence about the KD calculated from the koff/kon of the kinetic data. After all, the 1:1 fitting model cannot handle many artifact responses such as: 1) the bulk shift (bulk refractive index change) resulting from refractive index difference between analyte and running buffer; and 2) the mass transport effect resulting from the transport of analyte from its bulk solution to the Sensor Chip surface. However, we did use identical analyte and running buffer (1× PBS) to eliminate any possible bulk shift or bulk refractive index changes. Meanwhile, the observed curvature of the sensorgram during the association phase of pCTS-L–TLR4 interaction indicated a typical exponential increase of signal that argued against potential mass transport effect (Figure 3). Nevertheless, it remains elusive whether TLR4 analyte heterogeneity (e.g., oligomerization) or minute amounts of contaminant-associated non-specific binding somewhat contribute to the irregularity of pCTS-L–TLR4 sensorgrams (Figure 3).
In the absence of a perfect fit to the simplest 1:1 binding model, the equilibrium constant (KD) should also be estimated independently by equilibrium analysis if the steady-state binding data covers a wide range of analyte concentrations. Accordingly, we estimated the KD for pCTS-L–TLR4 interaction by evaluating the extrapolated steady-state signals as a function of analyte concentrations, assuming that a 10%, 50%, and 90% saturation is obtained at analyte concentrations of 0.1-fold, 1.0-fold, and 10.0-fold of the KD, respectively. Our estimated KD value (< 28.3 nM) from the dose-dependent signal saturation closely agreed with the KD (20.2 ± 3.5, Table 1) calculated from the koff/kon of the kinetic data. In addition, we performed additional OpenSPR experiments by reversely conjugating TLR4 onto different Sensor Chips before applying pCTS-L as the analyte at serial dilutions (Zhu et al., 2023). Consequently, we obtained a slightly higher but similar KD (64.6 nM vs. 20.2 ± 3.5 nM) for pCTS-L–TLR4 interaction from improved sensorgrams that exhibited an almost perfect fit to the 1:1 binding model (Zhu et al., 2023), supporting the usefulness of OpenSPR in estimating the KD for protein–protein interactions.
In contrast to typical kinetic analysis performed with alternating cycles of analyte injections and surface regenerations, we employ sequential injections of increasing concentrations of the analyte over the ligand without regeneration between each sample concentration, because harsh regeneration procedures may disrupt ligand capture by the NTA Sensor Chips. However, this single-cycle kinetics OpenSPR gives us an opportunity to assess whether pretreatment with other ligand-binding proteins (e.g., pCTS-L–neutralizing antibodies) affects pCTS-L ligand binding to analytes (e.g., TLR4 or RAGE) subsequently introduced at identical and increasing concentrations. For instance, when an irrelevant control monoclonal antibody (“c-mAb”) was injected onto the pCTS-L–conjugated NTA sensor at an extremely high concentration (e.g.,1200 nM), it did not affect subsequent pCTS-L ligand binding to either TLR4 or RAGE (Figure 5) (Zhu et al., 2023). In sharp contrast, when a pCTS-L–neutralizing IgG (mAb20) was first injected onto the pCTS-L–conjugated NTA sensor, it markedly reduced pCTS-L ligand’s interaction with TLR4 or RAGE analyte subsequently introduced at identical and increasing concentrations, as manifested by an almost 55-fold (from 20.3 ± 2.3 nM to 1144.3 ± 173.6 nM) and 10-fold (from 3.1 ± 0.4 nM to 30.4 ± 9.8 nM) increase in the KD for TLR4 and RAGE, respectively (Figure 5) (Zhu et al., 2023). The specific impairment of pCTS-L interaction with TLR4 or RAGE by a pCTS-L ligand-specific mAb supports specific interactions between pCTS-L ligand and TLR4 or RAGE analyte.
Figure 5. Effect of a pCTS-L-neutralizing monoclonal antibody (mAb20) on pCTS-L ligand interaction with TLR4 or RAGE. Recombinant pCTS-L was immobilized on the NTA Sensor Chip, and an irrelevant control monoclonal antibody (c-mAb) or a pCTS-L–neutralizing mAb (mAb20) was pre-exposed to pCTS-L–conjugated NTA Sensor Chip before subsequent injection of TLR4 (Left Panels) or RAGE (Right Panels) analyte at increasing concentrations to estimate the KD.
General notes and troubleshooting
As the OpenSPR uses microfluidic tubing with relatively narrow diameters to save precious analyte samples, all solutions (e.g., running buffers and distilled water) must be filtered through a 0.2 μm filter to prevent any large particulates from entering and clogging the fluidic lines of the OpenSPR. Similarly, analyte solutions should also be carefully inspected for any large particulates before injection into the OpenSPR.
In case of tubing clog, the user can purchase a Declogging Kit from the Nicoya Lifesciences to resolve the problem.
Injections of different samples should be performed using separate disposable syringes to avoid cross-contamination. If using the glass Hamilton syringe, ensure it is thoroughly washed with running buffer prior to using it for different samples.
The minimum sample volume required is the volume of the Sample Loop plus 50 μL of excess volume to ensure a uniform sample concentration profile as well as absence of remaining air within the Sample Loop.
The OpenSPR NTA Sensors provide a convenient capture-coupling technique to immobilize recombinant proteins with 6× His-tags, because its functional NTA groups can capture the protein ligand via its His-tag in the presence of NiCl2. These gold nanoparticles conjugated with NTA capture the His-tag ligand in a specific orientation, but the ionic interaction between the His-tag and the Ni-NTA surface is weaker than covalent direct coupling. This may result in some loss of the ligand sample from the surface over time particularly when measuring stronger (low dissociation rate) kinetic interactions with OpenSPR. Because of potential inherent dissociation of the captured ligand, this immobilization method is not recommended for analysis of kinetic systems with slow dissociation.
The overall shape of the curve is determined by the analyte concentrations, association rate (kon), and dissociation rate (koff) constants, which are independent of the concentrations of analyte and ligand. However, these association and dissociation constants are highly sensitive to the pH and salt concentrations of the solution, reinforcing the importance of keeping experimental conditions constant throughout the studies. Of course, running buffers should not contain chelating or reducing agents (e.g., EGTA) that will remove the necessary Ni2+ ions or alter the Ni2+ redox state, both of which may compromise the NTA surface activation.
We do not know whether there is a size limit for the ligands that can be used with the OpenSPR, as we have not tried any smaller peptides in the OpenSPR analysis.
If the active sites of NTA Sensor Chips were not fully occupied by the ligand, there is a possibility for false positive results if analytes contain multiple surface-exposed histidine residues. Therefore, control experiments using His-tag negative control proteins (with similar molecular weights but unable to interact with the ligand) should be performed to address these potential problems.
Acknowledgments
The research in Dr. Wang’s laboratory was supported by the National Institutes of Health (NIH) grants R01AT005076, R01GM063075, and R35GM145331. The OpenSPR method used to characterize pCTS-L interaction with two putative receptors was first described in the research article by Zhu et al. (2023).
Competing interests
None of the authors has any conflicts of interest to declare.
References
Chen, W., Qiang, X., Wang, Y., Zhu, S., Li, J., Babaev, A., Yang, H., Gong, J., Becker, L., Wang, P., et al. (2020). Identification of tetranectin-targeting monoclonal antibodies to treat potentially lethal sepsis. Sci. Transl. Med. 12(539): eaaz3833.
Drescher, D. G., Selvakumar, D. and Drescher, M. J. (2018). Analysis of Protein Interactions by Surface Plasmon Resonance. Adv. Protein Chem. Struct. Biol. 110: 1–30.
Qiang, X., Li, J., Zhu, S., He, M., Chen, W., Al-Abed, Y., Brenner, M., Tracey, K. J., Wang, P., Wang, H., et al. (2022). Human Dermcidin Protects Mice Against Hepatic Ischemia-Reperfusion–Induced Local and Remote Inflammatory Injury. Front. Immunol. 12: e821154.
Willander, M. and Al-Hilli, S. (2009). Analysis of biomolecules using surface plasmons. Methods Mol. Biol. 544: 201–229.
Zhu, C. S., Qiang, X., Chen, W., Li, J., Lan, X., Yang, H., Gong, J., Becker, L., Wang, P., Tracey, K. J., et al. (2023). Identification of procathepsin L (pCTS-L)–neutralizing monoclonal antibodies to treat potentially lethal sepsis. Sci. Adv. 9(5): eadf4313.
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4,796 | https://bio-protocol.org/en/bpdetail?id=4796&type=0 | # Bio-Protocol Content
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Fast and Sustainable Thermo-osmotic DNA Extraction Protocol for Trans-spectrum Contingency and Field Use
SG Stavroula Goudoudaki
MK Manousos E. Kambouris
MM Marianna Manoussopoulou
GP George P. Patrinos
AV Aristea Velegraki
YM Yiannis Manoussopoulos
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4796 Views: 501
Reviewed by: Shyam SolankiOlga Sin Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in OMICS: A Journal of Integrative Biology Feb 2023
Abstract
In the field of molecular genetics, DNA extraction protocols and kits are sample-specific and proprietary, preventing lateral distribution among similar facilities from different sectors to alleviate supply shortages during a crisis. Expanding upon previous fast extraction protocols such as alkaline- and detergent-based ones, the use of boiling-hot water to rupture cells, virions, and nuclei, as proposed during the COVID-19 pandemic, might alleviate shortages and costs. Different soft, relatively abundant (highly enriched), and uncomplicated (genomically homogenous and with few inhibitors) biosamples are collected in 1.5 mL tubes, mixed with boiling-hot water, and stirred vigorously, so as to have membranes lysed and proteins deactivated; mechanical disruption may be used as well if necessary. Incubation in boiling water bath for 20–30 min follows. Depending on sample type and quantity, which affects the total extraction volume, 2–5 μL are pipetted off for direct PCR and the same volume for two decimal serial dilutions. The latter are intended to optimize the crude extract to a workable DNA/inhibitor concentration balance for direct PCR. Uncomplicated, highly enriched samples such as mycelial growth in fruits and human swab samples can be processed, contrary to complicated samples such as blood and physically unyielding samples such as plant tissue. The extract can be used for immediate PCR in both benchtop and portable thermocyclers, thus allowing nucleic acid amplification tests (NAAT) being performed in resource-limited settings with low cost and waste footprint or during prolonged crises, where supply chain failures may occur.
Key features
• DNA extraction from different sample types using only boiling water and occasional mechanical assistance.
• Crude extract serially diluted twice, 10- and 100-fold, to bypass purification and quantification steps.
• Direct PCR for 2–10 μL of crude lysate and dilutions (conditional to sample type and quantity) to enhance probability of workable DNA-inhibitors’ concentrations.
• Lowers the cost and curtails the overall footprint of testing to increase sustainability in field operations and in standard lab environments under supply chain derailment.
Keywords: Thermo-osmotic DNA extraction COVID-19 Bio surveillance Ecology Minimal resource analysis One Health Pandemic fungal infections Plant pathogen detection Resource-limited settings
Background
A fast and massive DNA extraction is quintessential for nucleic acid amplification tests (NAAT), especially if performed under duress; the latter implies either field conditions in resource-limited settings (RLS), in routine or expedient setups, or in abnormal conditions with samples surging and/or supply pipelines malfunctioning due to corporate concerns or supply chain overextension/collapse. Thus, although the technology for DNA extraction systems achieves significant yields from complicated and/or miniscule samples, their use is expensive even if purchased in bulk to achieve economies of scale. Alkaline approaches were suggested for contingency (Goudoudaki et al., 2021) and field conditions (Priye et al., 2016), adaptable to the intended use (Goudoudaki et al., 2021). Still, specific chemicals were used, which may not be available in proper quantities once a health or ecological crisis prompts massive spatial dispersion and increased processing output of incoming samples, which may overload standard public health or agronomic facilities and infrastructures. Thus, ease of procurement and use by personnel moderately trained in this specific discipline but experienced in other sectors of biosciences may increase the processing rate of initial screening and offload work from dedicated infrastructures, restricting their involvement to tackle only unresolved or suspect samples. Last but not least, this almost in-situ processing approach may limit the need for dispatching samples to distant facilities, which is expensive, time consuming, creates biosafety and biosecurity risks, and may degrade the samples, thus de-valuing the actual diagnostic procedure.
By using only distilled water and heat, both easily available, there is no chemical waste and no need for special reagents, and thus for diverse stockpiles. The latter may be inexpensive and long-lead or long-expiration date, but their storage, even in normal conditions, takes space and there is a need for correct management of the stock. Also, the skills necessary are basic, found in everyone with the most elementary training in Biosciences. Thus, the overall footprint of the analysis, both before- and after-use, is kept minimal. This protocol is actually an expansion of a previous one, developed during the COVID-19 pandemic (Fomsgaard and Rosenstierne, 2020) as a result of kits supply restrictions (Benda et al., 2021), to include different extraction substrates and target genomes. It can be used in the field, given that consumables, heat, and electricity sources for portable thermocyclers (Kambouris et al., 2020) can be provided. Compared to the alkaline protocol (Goudoudaki et al., 2021), the thermo-osmotic protocol described herein requires even less reagents and produces no chemical waste, nor does it require stocking reagents that may expire and require specific storage. The protocol is named after the thermal deterioration of the membrane bonds allowing osmosis to perform the cell lysis and has been used already in one original publication (Goudoudaki et al., 2023).
Additionally, the present protocol may be used for improving the turnaround time and volume throughput in existing, indoor facilities. This comes at the expense of other performance metrics including sensitivity, which are markedly inferior compared to the ones achieved by dedicated DNA extraction protocols, even those of relatively low cost (Velegraki et al., 1999a and 1999b). Furthermore, facilities commandeered during crisis management may be among the beneficiaries. The method may be used for public health and environmental and agronomic emergencies (Kambouris et al., 2018), thus qualifying for compatibility with the One-Health framework and any other that may refer to genetic/genomic biomarkers that may be typed by basic PCR protocols, with or without follow ups such as, but not restricted to, restriction digestion (Velegraki et al., 1999a and 1999b; Arabatzis et al., 2004) and single-locus sequencing (Irinyi et al., 2015). The method, being plain and simple, shows moderate performance and should be used for highly enriched (containing high numbers of target genomes) and uncomplicated (without similar loads of pseudo-targets and inhibitors) samples.
Materials and reagents
The provisions below do not include expendables and instrumentation necessary for PCR and downstream procedures, such as gel electrophoresis.
Biological materials
Swab/scrap samples from human oral mucosa (tongue or inner cheek/gum)
Fungal mycelia grown on fruits and vegetables
Reagents
Commercially available distilled water for house use (i.e., steam ironing) from any convenience store and manufacturer, upon availability
Absolute ethanol, 99.8% denatured with IPA, MEK, and Bitrex pure (Panreac/Applichem, catalog number: 147194) or any other of comparable concentration (high purity is not essential)
Solutions
70% ethanol (see Recipes)
Recipes
70% ethanol
Reagent Final concentration Volume
Ethanol (absolute) 70% 700 mL
H2O 30% 300 mL
Total n/a 1,000 mL
Laboratory supplies
Falcon-shaped plastic tube, 50 mL (KIMA Vacutest, catalog number: KIMA-17102-50)
Beaker, 1,000 mL (Hamed, catalog number: 303173)
Polypropylene floating rack, 8 positions (Flinn Scientific, catalog number: FB1671)
Pasteur pipettes, glass (Hamed, catalog number: 303107)
Rubber bulb for Pasteur pipettes (Isolab Laborgerate, catalog number: 084-03-001)
Pipette P20, 0–20 μL (Gilson PIPETMAN®, catalog number: F123600)
Pipette P200, 20–200 μL (Gilson PIPETMAN®, catalog number: F123601)
Pipette P1000, 200–1,000 mL (Gilson PIPETMAN®, catalog number: F123602)
Pipette tips, 0–200 μL (KIMA Vacutest, catalog number: KIMA-18260)
Pipette tips, 1,000 μL (KIMA Vacutest, catalog number: KIMA-18172)
Plastic microtubes, 1.5 mL (PierceTM, catalog number: 69715)
Plastic microtubes for PCR, 0.2 mL (Thermo ScientificTM, catalog number: AB0620)
Scalpel blades (no. 22 stainless disposable; FEATHER, lot 03067430) or box cutter (e.g., Q-Connect, catalog number: 233471)
Rack (50-position for 18 mm diameter microtubes) (Fruugo)
Hydrophilic cotton (e.g., pharmacy package)
Plastic spoons (e.g., any convenience store, 10 pc package), spoons (e.g., stainless steel 18/0 convenience store), glass slides (76 mm × 26 mm, Knittel Glass, catalog number: 303157), or wood tongue depressor (disposable, Heine, catalog number: 508424)
Plastic pellet pestles, blue polypropylene autoclavable (Sigma-Aldrich, catalog number: Z359947-100EA)
Cotton (any pharmacy/convenience store) or paper tissue/napkins (any convenience store)
Equipment
Camping gas stove (e.g., KEMPER, 12.3 cm × 12.3 cm × 21.7 cm, model: 06450208) or electric kitchen stove burner (e.g., Severin, model: 3519368)
Lighter (e.g., KEMPER, model: 10422)
Procedure
Sample collection (see General note 1) and preparation
If swab/tissue collector item is not sterile, rub it with 70% alcohol on a piece of cotton or paper tissue/napkin and light it with the lighter.
Allow the flame to die out and let it cool down in the air.
Sample gathering; either:
Collect a swab from the upper surface of the tongue or the inner surface of the cheek by mildly scrapping with any of the proposed or other similar objects (see General note 2);
Cut off a piece (3–5 mm in diameter) of the mycelium grown on the plant tissue. Alternatively, scrap it from animal tissue or scrap bacterial or yeast growth from any such surface by using disposable sterile scalpel bladed or flame-sterilized scalpel, razor, or box cutter (see General note 3).
Store sample in a 1.5 mL microtube (see Troubleshooting 1) until step B1. If needed, use some guidance/assistance to put it in (see General note 4).
Fill a beaker to half capacity with tap water; this will serve as a water bath.
Fill a 50 mL Falcon tube to half capacity with commercially available distilled water and place it into the beaker.
Heat the beaker on the stove (camping gas or electrical) at full intensity until the water bath starts boiling; then, reduce to half intensity to keep the water boiling during the entire procedure.
Cell lysis
Add boiling hot water to the extraction tube
If mechanical assistance in lysis is required (due to the nature, quantity, or physical qualities of the sample), add 250 μL of boiling-hot distilled water or half the volume you intend to use (precision is of no particular importance) to the extraction microtube containing the sample from step A4 (see General note 5).
If no mechanical assistance is required, add the sum of the extraction volume of boiling water and go straight to step B4. Caution: Steam may come out of the Falcon tube upon unscrewing and the tube may be hot. Take precautions during touching and handling and keep it safely placed on the respective rack or any similar support (e.g., a small beaker or glass).
Crush with a spiral motion of the pestle or other similar instrument for 2–3 min.
Use an equal volume of boiling-hot distilled water (e.g., 250 μL, as suggested in step B1a) to wash cellular debris of the pestle within the microtube.
Close the lid of the microtube and vortex or shake vigorously by hand for 1 min (see General note 6)
Incubation
Place the microtubes in the boiling water bath for 20–30 min or more, according to the extraction volume and rigor of extraction (see General note 7; see Troubleshooting 2). Use the incubation time to prepare the molecular testing reactions, if applicable (step E1).
Upon completion, carefully open microtubes and pipette 10 μL from the center of the extraction volume to a PCR microtube.
Caution: Steam pressure will have built up in the microtube; thus, handle with care and away from the face. Preferably, place in a rack, either custom made or improvised, before opening and further handling.
Dilutions
Transfer 5 μL of the 10 μL to the first dilution tube and mix with 45 μL of distilled water by softly pipetting in and out 5–10 times, so as to have a 1/10 dilution. Use a 20 μL pipette for the whole procedure.
Remove 5 μL of the 1/10 dilution to a second PCR tube (of 200 μL volume or as used per standard practice).
Remove another 5 μL of the 1/10 dilution to the second dilution tube and mix with 45 μL of distilled water by softly pipetting in and out 5–10 times, so as to have a 1/100 dilution.
Remove 5 μL of the 1/100 dilution to a third PCR tube (see General note 8).
PCR amplification
Prepare the mastermix, according to the desired protocol, while incubating the extract in the water bath and keep on ice until use (see General note 8). This step should be implemented during step C1.
Dispense the required volumes of mastermix to the three PCR tubes with the serial dilutions for each sample and proceed to amplification in thermocycler as per standard procedure/protocol. This step should be implemented after step D4.
Data analysis
Each extraction was performed in triplicates independently, with no more than a week between two successive efforts. Any of the three dilutions producing amplicon was deemed successful. Here, we report that the protocol can be used in cases where at least two of the three independent extraction efforts produced amplicon in at least one out of three serial dilutions (the undiluted crude extract being understood as the first in the series of the three dilutions).
Validation of protocol
There is no way to verify the performance of the present protocol directly: it is an extraction protocol and since its product is not purified, it cannot be tested properly—neither by spectrophotometry, as the cell debris would absorb or scatter the emitted UV, nor by gel electrophoresis, as said debris would obstruct the motion of the DNA, mar the electric field, and also absorb the fluorescent dyes, creating noise and smear. The only applicable validation approach is indirect, by the success or failure of some downstream nucleic acid amplification test (NAAT). To secure validation, negative controls with distilled water-only added in equal volume to the template were used, to ensure that amplicon production was not due to contamination of the NAAT reagents, expendables, or equipment.
Furthermore, to ensure that the NAAT (here, PCR) does work in terms of protocol, cycler, and reagents, positive controls were used. These were semi-simultaneous extractions of mycelial tissue, by other established methods such as alkaline extraction and/or CTAB (Velegraki et al., 1999a; Goudoudaki et al., 2021), and also of stock DNA extracts kept in the lab from previous projects, extracted with conventional protocols and occasionally quantified by mini-spectrophotometer (Psarias et al., 2020). The full procedure for validation and controls may be found in the source publication (Goudoudaki et al., 2023). In short, from a number of tried applications, positive PCR results were returned for two cases of samples: (a) swab samples tested either by consensus sequence PCR targeted at the local bacteriome (product band of size compatible with the expected target range) or for one specific endogenous (human) locus (product band of the expected size); (b) consensus sequence PCR on extracts of fungal mycelia grown on the surface of fruits held in refrigeration (product band of size compatible with the expected target range).
General notes and troubleshooting
General notes
The method is valid with relatively soft and uncomplicated samples in which the target genome is abundant (highly enriched). Mouth swabs produced specific human genome amplicons and consensus sequences amplification product for bacteria but not for fungi, which are much less populous in the oral mucosa.
Glass slides for microscopy, plastic or metallic spoon—especially disposable ones for deserts—or the handle of all-metallic cutlery or tongue depressors may be used.
During sampling, samples as pure as possible of target cells must be collected (pure is understood as “containing as few admixtures and as homogenous a cellular population as possible”). The quantity of the sample must be limited, as there is no purification step to dispose of cell debris and consequent background noise and possible inhibitions.
If needed, help/guide the sample within the microtube with a similarly sterilized microbiology needle or a similar piece (i.e., a second glass slide) or a plastic tip.
If mechanical assistance is required, take caution that the volume of sample + boiled water is approximately 500 μL. Additional boiling water will be added to wash off the pestle from attached debris, but it is better that the final volume does not exceed 1 mL.
If the extraction microtube is too full (> 1 mL), punch holes using a needle, thin nail, or pin in the microtube lid before incubation so the pressure will not blow the lid open, throwing the content out of the tube.
Use purpose-built floating rack or punch/cut holes to felizol (EPC–Expanded PolyStyrene) sheets of less than 50 mm thickness recovered from packaging boxes.
Functional concentration during the NAAT may be adjusted to other values: either by i) increasing the template volume in the reaction tube while subtracting an equal volume of sterile distilled H2O from the mastermix or ii) if no mastermix is prepared, by changing the volumes in the reaction microtube. Occasionally, the opposite may also be applicable (diminishing the template volume while increasing that of the sterile distilled H2O). Similar manipulations are applicable at the serial dilutions step.
Troubleshooting
If the sample size within the microtube is excessively large (operative’s call, depending on the difficulty to properly handle the extraction mix, but see General note 6 for an example), it may be divided in two. If it is solid, this can be done by cutting with a sterilized surgical disposable blade; if it is liquid, the excess may be pipetted off to another tube, creating a technical replicate.
Longer incubation time is advised (a) when treating larger samples in terms of volume/mass and (b) if the cell lysis phase is less rigorous, as in the absence of mechanical assistance. Refer to step C1.
Acknowledgments
This project was in part funded by the European Commission, Horizon 2020 (H2020-668353; Ubiquitous Pharmacogenomics).
This protocol is based on the research paper (Goudoudaki et al. 2023).
Competing interests
The authors declare no conflicts of interest or competing interests.
Ethical considerations
The study was approved by the Institutional Research Ethics Committee of the University of Patras. Swabs were collected from the operatives performing the development of the protocol and the subsequent analysis as explicitly stated in the paper (Goudoudaki et al., 2023).
References
Arabatzis, M., Kollia, K., Menounos, P., Logotheti, M. and Velegraki, A. (2004). Delineation of Clavispora lusitaniae clinical isolates by polymerase chain reaction-single strand conformation polymorphism analysis of the ITS1 region: a retrospective study comparing five typing methods. Med. Mycol. 42(1): 27–34.
Benda, A., Zerajic, L., Ankita, A., Cleary, E., Park, Y. and Pandey, S. (2021). COVID-19 Testing and Diagnostics: A Review of Commercialized Technologies for Cost, Convenience and Quality of Tests. Sensors 21(19): 6581.
Fomsgaard, A. S. and Rosenstierne, M. W. (2020). An alternative workflow for molecular detection of SARS-CoV-2 – escape from the NA extraction kit-shortage, Copenhagen, Denmark, March 2020. Eurosurveillance 25(14): e2000398.
Goudoudaki, S., Kambouris, M. E., Siamoglou, S., Gioula, G., Kantzanou, M., Manoussopoulou, M., Patrinos, G. P. and Manoussopoulos, Y. (2023). Can Water-Only DNA Extraction Reduce the Logistical Footprint of Biosurveillance and Planetary Health Diagnostics? Toward a New Method. OMICS: J. Integr. Biol. 27(3): 116–126.
Goudoudaki, S., Milioni, A., Kritikou, S., Velegraki, A., Patrinos, G. P., Gioula, G., Manoussopoulos, Y. and Kambouris, M. E. (2021). Fast, Scalable, and Practical: An Alkaline DNA Extraction Pipeline for Emergency Microbiomics Biosurveillance. OMICS: J. Integr. Biol. 25(8): 484–494.
Irinyi, L., Serena, C., Garcia-Hermoso, D., Arabatzis, M., Desnos-Ollivier, M., Vu, D., Cardinali, G., Arthur, I., Normand, A. C., Giraldo, A., et al. (2015). International Society of Human and Animal Mycology (ISHAM)-ITS reference DNA barcoding database—the quality controlled standard tool for routine identification of human and animal pathogenic fungi. Med. Mycol. 53(4): 313–337.
Kambouris, M. E., Manoussopoulos, Y., Kritikou, S., Milioni, A., Mantzoukas, S. and Velegraki, A. (2018). Toward Decentralized Agrigenomic Surveillance? A Polymerase Chain Reaction–Restriction Fragment Length Polymorphism Approach for Adaptable and Rapid Detection of User-Defined Fungal Pathogens in Potato Crops. OMICS: J. Integr. Biol. 22(4): 264–273.
Kambouris, M. E., Siamoglou, S., Kordou, Z., Milioni, A., Vassilakis, S., Goudoudaki, S., Kritikou, S., Manoussopoulos, Y., Velegraki, A., Patrinos, G. P., et al. (2020). Point-of-need molecular processing of biosamples using portable instrumentation to reduce turnaround time. Biosaf. Health 2(3): 177–182.
Priye, A., Wong, S., Bi, Y., Carpio, M., Chang, J., Coen, M., Cope, D., Harris, J., Johnson, J., Keller, A., et al. (2016). Lab-on-a-Drone: Toward Pinpoint Deployment of Smartphone-Enabled Nucleic Acid-Based Diagnostics for Mobile Health Care. Anal. Chem. 88(9): 4651–4660.
Psarias, G., Iliopoulou, E., Liopetas, I., Tsironi, A., Spanos, D., Tsikrika, A., Kalafatis, K., Tarousi, D., Varitis, G., Koromina, M., et al. (2020). Development of Rapid Pharmacogenomic Testing Assay in a Mobile Molecular Biology Laboratory (2MoBiL). OMICS: J. Integr. Biol. 24(11): 660–666.
Velegraki, A., Kambouris, M., Kostourou, A., Chalevelakis, G. and Legakis, N. J. (1999a). Rapid extraction of fungal DNA from clinical samples for PCR amplification. Med. Mycol. 37(1): 69–73.
Velegraki, A., Kambouris, M., Skiniotis, G., Savala, M., Mitroussia-Ziouva, A. and Legakis, N. (1999b). Identification of medically significant fungal genera by polymerase chain reaction followed by restriction enzyme analysis. FEMS Immunol. Med. Microbiol. 23(4): 303–312
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4,797 | https://bio-protocol.org/en/bpdetail?id=4797&type=0 | # Bio-Protocol Content
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Functional Assay for Measuring Bacterial Degradation of Gemcitabine Chemotherapy
SS Serkan Sayin
AM Amir Mitchell
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4797 Views: 511
Reviewed by: Chiara AmbrogioIstvan Stadler Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Drug biotransformation by the host microbiome can impact the therapeutic success of treatment. In the context of cancer, drug degradation can take place within the microenvironment of the targeted tumor by intratumor bacteria. In pancreatic cancer, increased chemo-resistance against the frontline chemotherapy gemcitabine is thought to arise from drug degradation by the tumor microbiome. This bacterial–drug interaction highlights the need for developing rapid assays for monitoring bacterial gemcitabine breakdown. While chemical approaches such as high-performance liquid chromatography are suitable for this task, they require specialized equipment and expertise and are limited in throughput. Functional cell-based assays represent an alternate approach for performing this task. We developed a functional assay to monitor the rate of bacterial gemcitabine breakdown using a highly sensitive bacterial reporter strain. Our method relies on standard laboratory equipment and can be implemented at high throughput to monitor drug breakdown by hundreds of strains simultaneously. This functional assay can be readily adapted to monitor degradation of other drugs.
Key features
• Quantification of gemcitabine breakdown by incubating bacteria that degrades the drug and subsequently testing the growth of a reporter strain on filtered supernatant.
• Use of an optimized reporter strain that was genetically engineered to be a non-degrader strain and highly sensitive to gemcitabine.
• A high-throughput assay performed in microplates that can be adjusted for identifying bacteria with a fast or slow gemcitabine degradation rate.
• The assay results can be compared to results from a standard curve with known drug concentrations to quantify degradation rate.
Graphical overview
Protocol overview. (1) Bacteria are incubated with gemcitabine for a set period of time. (2) Samples are removed from co-incubated suspensions and filtered to remove bacteria to halt gemcitabine degradation. (3) A gemcitabine-sensitive reporter strain is then added to the conditioned supernatant and is supplemented with growth media. (4) Growth of the reporter strain is monitored over time. (5) Results from the growth experiments are used to infer the concentration of gemcitabine in the co-culture supernatant and the drug degradation rate.
Keywords: Drug degradation Drug breakdown Drug modification Gemcitabine Gemcitabine deamination Bacterial Biotransformation Functional assay Tumor-microbiome Cytidine deaminase
Background
The human microbiome can modulate the efficacy of different therapeutic interventions. In some cases, the effectiveness of host-targeting (non-antibiotic) drugs can be influenced by direct drug–microbiome interactions, such as bacterial drug biotransformation and bioaccumulation (Spanogiannopoulos et al., 2016; Zimmermann et al., 2019 and 2021; Klünemann et al., 2021). In the context of cancer, microbial drug biotransformation can take place in natural microbiome sites, such as the gut, or by tumor-colonizing bacteria. Recent work on pancreatic cancer revealed that gammaproteobacteria, frequently infecting pancreatic tumors in humans, can inactivate the frontline chemotherapy drug gemcitabine [2’,2’-difluoro 2’ deoxycytidine (dFdC)] and increase chemoresistance in murine tumor models (Geller and Straussman, 2017). Published clinical data further provided circumstantial evidence supporting the notion that bacterial drug inactivation may be impacting gemcitabine treatment efficacy in patients (Gao et al., 2020; Meriggi and Zaniboni, 2021; Mohindroo et al., 2021). However, an important and related observation that is often overlooked is that gemcitabine is a potent antimicrobial (Sayin et al., 2023), similar to many other antineoplastic drugs (Maier et al., 2018). We previously proposed and showed that bacteria can rapidly evolve resistance against anticancer chemotherapies that are also antimicrobial, and by doing so can also alter their drug metabolism and therefore inadvertently influence chemoresistance in the treated host (Rosener et al., 2020; Sayin et al., 2023).
Investigating bacterial biotransformation of host-targeted drugs is key for our understanding of cancer–microbiome interactions (Xavier et al., 2020). Chemical methods, such as high-performance liquid chromatography and mass spectrometry, are typically used to quantify drug biodegradation rate by monitoring extracellular drug concentration after bacterial incubation. However, despite their prevalence, these methods require specialized equipment and expertise, and are limited in experimental throughput. In contrast, functional cell-based assays represent an alternative category of approaches for gauging drug concentration by using toxicity as a proxy for its concentration. Specifically, the growth of drug-sensitive cells can be leveraged as a functional reporter for quantifying drug concentration. The use of commonplace microplate optical density (OD) readers for high-throughput monitoring of bacterial growth allows for multiplexing such assays and measuring drug breakdown across hundreds of bacterial cultures with common lab equipment.
We have developed a functional assay to monitor the rate of gemcitabine breakdown in bacteria (Sayin et al., 2023). We used the assay to uncover how loss-of-function mutations in dozens of genes that confer bacterial gemcitabine resistance influence the drug breakdown rate. For this approach, we first incubated the strain of interest in a saline buffer that contained a known drug concentration for set incubation periods. We then filtered the cell suspension and aliquoted the conditioned supernatant into fresh growth media. We inoculated into this media a genetically engineered reporter strain that was optimized for the functional assay—a highly sensitive strain that is incapable of drug degradation. We compared the growth of the reporter strain in multiple supernatants and identified supernatants with altered growth. Increased or decreased growth of the reporter strain was indicative of changes in drug concentration during the incubation period.
Materials and reagents
Biological materials
Reporter strain: Escherichia coli Δcdd mutant (KEIO knockout collection, Dharmacon, GE Life Sciences)
Test strain: any other mutant from KEIO knockout collection or any other bacterial species of interest
Reference strain: for KEIO mutants, the parent strain E. coli BW25113 (Dharmacon, GE Life Sciences)
Reagents
Kanamycin monosulfate (TCI, catalog number: K0047)
Gemcitabine hydrochloride salt > 99% (LC labs, catalog number: G-4177)
Phosphate-buffered saline (PBS) (Corning, catalog number: 21-040-CV)
M9 minimal salts (Difco, catalog number: 248510)
NaCl (Fisher Scientific, catalog number: S271-3)
Yeast extract (Gibco Bacto, catalog number: 212720)
Tryptone (Fisher Scientific, catalog number: BP1421-2)
MgSO4, anhydrous (Sigma-Aldrich, catalog number: M8266-100G)
CaCl2 (Fisher Scientific, catalog number: C79-500)
Protein hydrolysate amicase (Sigma-Aldrich, catalog number: 82514-1KG)
Glucose/Dextrose anhydrous (Fisher Scientific, catalog number: D14-500)
Solutions
Lysogeny broth (LB) (see Recipes)
M9 minimal medium (see Recipes)
Kanamycin stock solution (50 mg/mL in water) (see Recipes)
Gemcitabine stock solution (25 mg/mL in water corresponding to 83.4 mM) (see Recipes)
Recipes
Lysogeny broth
Reagent Final concentration Quantity
Tryptone 1% 10 g
Yeast extract 0.5% 5 g
NaCl 1% 10 g
Milli-Q H2O n/a 1 L
Autoclave the solution after dissolving all of the components.
M9 minimal medium
Reagent Final concentration Volume
M9 minimal salt solution (5×) 1× 200 mL
20% glucose solution 0.4% 20 mL
1 M MgSO4 2 mM 2 mL
1 M CaCl2 0.1 mM 0.1 mL
2% protein hydrolysate amicase 0.2% 100 mL
Autoclaved Milli-Q H2O n/a 678 mL
Filter sterilize the components of the M9 minimal medium except the M9 minimal salt solution, by passing through a 0.22 μm syringe filter or 0.22 μm vacuum-driven filter. Autoclave 5× M9 minimal salt solution to sterilize. Heat amicase solution at 42 °C water bath for dissolving. Combine all components in sterile conditions to prepare the final medium.
Kanamycin stock solution
Dissolve kanamycin in autoclaved Milli-Q water at 50 mg/mL and filter sterilize by passing through a 0.22 μm PES filter. Keep the stock solution at -20 °C.
Gemcitabine stock solution
Dissolve gemcitabine in autoclaved Milli-Q water at 25 mg/mL and filter sterilize by passing through a 0.22 μm PES filter. Keep the stock solution at -20 °C.
Laboratory supplies
96-well plate 0.22 μm filters (PALL Corporation, catalog number: 8119)
0.22 μm syringe filter, PES membrane, 30 mm diameter (Celltreat, catalog number: 229747)
10 mL syringes without needle (Air-tite, catalog number: ML10)
0.22 μm vacuum driven filter, PES membrane, 250 mL (GenClone/Genesee, catalog number: 25-225)
96-deep-well plates (Eppendorf, catalog number: 951033502)
96-well plates (Thermo Fisher Scientific, catalog number: 167008)
Cuvettes for spectrophotometer (Olympus Plastics, Genesee catalog number: 21-136)
Air permeable 96-well plate seals (Excel Scientific, catalog number: BS-25)
Adhesive 96-well plate seals (Thermo Scientific catalog number: AB-0558)
Equipment
Plate reader (BioTek Eon/TECAN Spark)
Spectrophotometer (Eppendorf, BioPhotometer plus)
Microcentrifuge (Eppendorf, Centrifuge 5424)
Centrifuge with multi-well plate carriers (Beckman Coulter Avanti centrifuge J-26 XPI with rotor JS-5.3)
Shaker incubator (New Brunswick Scientific, Excella E25)
Tabletop microplate shaker incubator (Heidolph Inkubator1000)
Optional: 96-channel handheld electric pipette (Integra, Viaflo 96)
Software and datasets
MATLAB or any other statistical software (e.g., R, Excel, SPSS)
Procedure
Preparing the conditioned supernatant
Figure 1A shows the overview of the procedure. For biological replicates, we recommend repeating the entire section A on different days.
Figure 1. Schematic representation of the protocol for determining gemcitabine degradation rate. A. Protocol for preparing the conditioned supernatants. Bacterial cultures, normalized to an identical optical density, are incubated with gemcitabine in PBS for defined periods of time. Samples are removed from the incubated cultures and filtered to yield conditioned supernatants. The assay can be paused after the filtering stage and supernatants can be frozen for later use. B. Protocol for the functional assay. The reporter strain (E. coli Δcdd) is grown overnight and aliquots of the conditioned supernatant are then added to the growth culture. Growth of the reporter strain is monitored in a microplate optical density (OD) reader. Changes in growth curves are used to infer the gemcitabine concentration in the conditioned supernatant of different strains (red: fast degrader; blue: slow degrader) compared to a reference strain (green). Parts of the figure were created with BioRender.com.
Inoculate your test bacterial strains into 1 mL of LB in a 96-deep-well plate and grow overnight at 37 °C, 200 rpm shaker. Also inoculate the reference strain for comparison. Critical: Inoculate multiple replicates of reference strain (at least six technical replicates)
On the next day, pellet the cells by centrifuging the plate at 5,000× g for 10 min using a centrifuge with multi-well plate holders.
Discard the supernatant without disrupting the pellet and resuspend the cells in 1 mL of PBS.
Pellet the cells by centrifugation at 5,000× g for 10 min using a centrifuge with multi-well plate holders.
Repeat steps A3–A4 two additional times.
Let the cells incubate in 1 mL of PBS for 1 h and then measure the OD (600 nm) after 1:4 dilution in PBS at a final volume of 200 μL (e.g., 50 μL of cell suspension, 150 μL of PBS).
Dilute the cultures to an OD of 0.5 in a 96-deep-well plate at a final volume of 1.1 mL of PBS.
Sample 200 μL from the diluted cultures and measure optical density to validate if the dilution was accurate. Critical: Accuracy of dilution is important since any deviation from the intended cell number will influence the drug breakdown speed and will be incorrectly interpreted as a change in drug breakdown rate.
Add gemcitabine at a final concentration of 200 μM: prepare 21× (4.2 mM) gemcitabine and pipette 45 μL into 900 μL of bacterial suspension. Critical: Make sure your gemcitabine stocks are kept sterile and avoid multiple freeze-thaw cycles.
Caution: Always wear appropriate personal protective equipment when working with gemcitabine and follow institutional health and safety regulations for managing gemcitabine waste. Gemcitabine is a cytotoxic chemotherapy agent.
Incubate the plate on a tabletop microplate shaker at 37 °C and 900 rpm.
At designated timepoints (15 and 45 min), transfer 400 μL of liquid culture into a 96-well plate filter placed onto a 96-deep-well plate and centrifuge immediately at 5,000× g for 5 min using a centrifuge with multi-well plate holders. Critical: A 15 min timepoint is suitable for identifying fast degraders, and a 45 min timepoint is suitable for identifying the slow degraders. These incubation periods can be adjusted for work with other bacterial species that degrade gemcitabine at considerably different rates than E. coli.
Safe stopping point: if you are not proceeding immediately to section B, cover the supernatant with sterile plastic foil and freeze at -20 °C.
Performing the functional assay: testing the conditioned supernatants
Figure 1B shows the overview of the procedure. We recommend performing section B in triplicates using three independent biological replicates that are collected by performing section A on three different days.
Inoculate a single colony of reporter mutant Δcdd into 3 mL of M9 media with 50 μg/mL kanamycin and grow overnight at 37 °C and 200 rpm in a shaker.
On the next day, measure the OD of the culture (after 1:20 dilution in M9) and dilute the culture to OD 1 in 1 mL of M9.
Further dilute the culture 1:500 in M9 (final volume of 25 mL) and pipette 190 μL into each well of a 96-well microplate.
Add 10 μL of conditioned buffer from section A into each well. This step should be performed quickly, so all individual cultures start the exposure at the same time point. Caution: If the conditioned supernatants to be tested are frozen, thaw them at room temperature in advance.
Optional: This step can be performed using a 96-channel handheld electric pipette to make sure all wells receive the conditioned supernatant simultaneously.
Monitor the growth of the culture in the microplate using a suitable microplate optical density reader. Define a repeating measurement cycle that includes incubation at 37 °C and double orbital shaking at 360 rpm with reading the OD every 10 min over 8 h (plate lid on). Make sure to measure M9 alone for a blank optical density value.
Data analysis
Identifying the fast and slow degraders
Subtract the blank optical density value from all data points. Alternatively, you can treat the average optical density at the first timepoint across all wells as the background optical density.
Plot the growth curves for each strain as time on x-axis and OD on y-axis for each replicate.
Calculate the area under the curve (AUC) up to 7 h of growth for each replicate.
Perform one-tailed paired Student’s t-test with test strain AUCs and reference strain AUCs.
Important: pick the correct tail of the distribution, depending on your interest in slow or fast degraders. For example, for determining fast degraders, using 15 min conditioned supernatant, use a right tail.
Adjust the p-values that were calculated with a false discovery rate (FDR) or Bonferroni correction. Determine the hits based on your p-adjusted cutoff (generally < 0.1 or < 0.05).
Validation of protocol
This protocol was used to produce data on panel B of Figure 2 in our previous publication (Sayin et al., 2023). Section A has been repeated as three biological replicates and section B has been performed three times with the supernatants collected from biological replicates performed on different days. For the reference strain, six technical replicates have been used with the wild-type E. coli strain. Other controls used for validation were a no-bacteria control and no-gemcitabine control. Since multiple hypothesis testing was performed, p-values calculated using one tailed Student’s t-test were adjusted using FDR and a cutoff of < 0.1 was used. Measurement of drug degradation from our functional assay for a fast degrader and a slow degrader were validated with a chemical method (gas chromatography–mass spectrometry) that measured drug concentration and its degradation product. Figure 2 shows examples of fast and slow gemcitabine degraders.
Figure 2. Representative results from the assay. Graphs show the growth curves of the reporter strain on conditioned supernatants after incubation with three E. coli strains: wild type, ΔcytR (fast degrader), and ΔnupC (slow degrader). The graph shows the mean of three replicates.
General notes and troubleshooting
General notes
We expect that the rate of gemcitabine degradation may differ considerably between different bacterial species. The functional assay can be adjusted to account for such differences. The following parameters may need to be optimized for work with other species:
Bacterial concentration: bacterial concentration in step 7 of section A should be lowered for work with fast degraders.
Incubation period: incubation period in step 11 of section A should be shortened for working with fast degraders.
Initial gemcitabine concentration: drug concentration in step 9 of section A can be decreased for slow degraders.
Multiple controls in section A must be included for sensible interpretation of the experiment results:
No-bacteria, no-gemcitabine control: for observing unhindered growth of the reporter strain.
No-bacteria, +gemcitabine: for validating that maximum gemcitabine concentration is indeed inhibitory for the reporter strain.
+bacteria, no-gemcitabine: for validating that bacteria are not secreting molecules into the supernatant that inhibit growth of the reporter strain.
+reference bacteria, +gemcitabine: for determining the gemcitabine breakdown in a reference strain (e.g., wild-type strain).
Limitations of the protocol
Technical artifacts in the functional assay can be incorrectly interpreted as strains with fast or slow degradation. Great attention should be therefore dedicated to validating that cell numbers and incubation periods are identical across all cultures. Moreover, microplate optical density readers can typically have slightly uneven growth conditions across the multi-well plates (e.g., non-uniform heating). To minimize these artifacts, we recommend shuffling the positions of biological replicates across the plate.
Applicability to other experimental systems/model organisms
This protocol can be used with any bacteria that can be cultured in the lab. This experimental approach can also be used to measure the degradation of other drugs, as long as a sensitive reporter strain can be identified.
Acknowledgments
Work in this article was supported by NIGMS and NIAID of the National Institutes of Health under award numbers R35GM133775 and R01AI170722. We would like to thank Dr Caryn Navarro for her comments on the manuscript. This protocol was previously used in our reported results (Sayin et al., 2023).
Competing interests
The authors declare no conflicts of interest or competing interests.
References
Gao, Y., Shang, Q., Li, W., Guo, W., Stojadinovic, A., Mannion, C., Man, Y. g. and Chen, T. (2020). Antibiotics for cancer treatment: A double-edged sword. J. Cancer 11(17): 5135–5149.
Geller, L. T. and Straussman, R. (2017). Intratumoral bacteria may elicit chemoresistance by metabolizing anticancer agents. Mol. Cell. Oncol. 5(1): e1405139.
Klünemann, M., Andrejev, S., Blasche, S., Mateus, A., Phapale, P., Devendran, S., Vappiani, J., Simon, B., Scott, T. A., Kafkia, E., et al. (2021). Bioaccumulation of therapeutic drugs by human gut bacteria. Nature 597(7877): 533–538.
Maier, L., Pruteanu, M., Kuhn, M., Zeller, G., Telzerow, A., Anderson, E. E., Brochado, A. R., Fernandez, K. C., Dose, H., Mori, H., et al. (2018). Extensive impact of non-antibiotic drugs on human gut bacteria. Nature 555(7698): 623–628.
Meriggi, F. and Zaniboni, A. (2021). Antibiotics and steroids, the double enemies of anticancer immunotherapy: a review of the literature. Cancer Immunol., Immunother. 70(6): 1511–1517.
Mohindroo, C., Hasanov, M., Rogers, J. E., Dong, W., Prakash, L. R., Baydogan, S., Mizrahi, J. D., Overman, M. J., Varadhachary, G. R., Wolff, R. A., et al. (2021). Antibiotic use influences outcomes in advanced pancreatic adenocarcinoma patients. Cancer Medicine 10(15): 5041–5050.
Rosener, B., Sayin, S., Oluoch, P. O., García González, A. P., Mori, H., Walhout, A. J. and Mitchell, A. (2020). Evolved bacterial resistance against fluoropyrimidines can lower chemotherapy impact in the Caenorhabditis elegans host. eLife 9: e59831.
Sayin, S., Rosener, B., Li, C. G., Ho, B., Ponomarova, O., Ward, D. V., Walhout, A. J. and Mitchell, A. (2023). Evolved bacterial resistance to the chemotherapy gemcitabine modulates its efficacy in co-cultured cancer cells. eLife 12: e83140.
Spanogiannopoulos, P., Bess, E. N., Carmody, R. N. and Turnbaugh, P. J. (2016). The microbial pharmacists within us: a metagenomic view of xenobiotic metabolism. Nat. Rev. Microbiol. 14(5): 273–287.
Xavier, J. B., Young, V. B., Skufca, J., Ginty, F., Testerman, T., Pearson, A. T., Macklin, P., Mitchell, A., Shmulevich, I., Xie, L., et al. (2020). The Cancer Microbiome: Distinguishing Direct and Indirect Effects Requires a Systemic View. Trends Cancer 6(3): 192–204.
Zimmermann, M., Patil, K. R., Typas, A. and Maier, L. (2021). Towards a mechanistic understanding of reciprocal drug–microbiome interactions. Mol. Syst. Biol. 17(3): e202010116.
Zimmermann, M., Zimmermann-Kogadeeva, M., Wegmann, R. and Goodman, A. L. (2019). Mapping human microbiome drug metabolism by gut bacteria and their genes. Nature 570(7762): 462–467.
Article Information
Copyright
© 2023 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 > Antibacterial assay
Biological Sciences > Biological techniques > Microbiology techniques
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4,798 | https://bio-protocol.org/en/bpdetail?id=4798&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Detection of Cytoplasmic and Nuclear Circular RNA via RT-qPCR
KT Ke-En Tan
WN Wei Lun Ng
CE Chee-Kwee Ea
YL Yat-Yuen Lim
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4798 Views: 1415
Reviewed by: Durai SellegounderSalim Gasmi Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in Scientific Reports Jul 2021
Abstract
Circular RNA (circRNA) is an intriguing class of non-coding RNA that exists as a continuous closed loop. With the improvements in high throughput sequencing, biochemical analysis, and bioinformatic algorithms, studies on circRNA expression became abundant in recent years. However, functional studies of circRNA are still limited. Subcellular localization of circRNA may provide some clues in elucidating its biological functions by performing subcellular fractionation assay. Notably, circRNAs that are predominantly found in the cytoplasm are more likely to be involved in post-transcriptional gene regulation, e.g., acting as micoRNA sponge, whereas nuclear-retained circRNAs are predicted to play a role in transcriptional regulation. Subcellular fractionation could help researchers to narrow down and prioritize downstream experiments. The majority of the currently available protocols describe the steps for subcellular fractionation followed by western blot analysis for protein molecules. Here, we present a protocol for the subcellular fractionation of cells to detect circRNA via RT-qPCR with divergent primers. Moreover, detailed steps for the generation of specific circRNAs-enriched cDNA included in this protocol will enhance the amplification and detection of low-abundance circRNAs. This will be useful for researchers studying low-abundance circRNAs.
Key features
• This protocol builds upon the method developed by Gagnon et al. (2014) and extends its application to circRNA study.
• Protocol for amplification of low levels of circRNA expression.
• Analysis takes into consideration the ratio of cytoplasmic RNA concentration to nuclear RNA concentration.
Graphical overview
Keywords: Subcellular fractionation RT-qPCR Circular RNA circRNA Cytoplasmic circRNA Nuclear circRNA Low-abundance circRNA
Background
Circular RNA (circRNA) is an intriguing class of non-coding RNA that is formed through a unique mechanism known as backsplicing, in which the 5′ and 3′ termini are covalently joined (Jeck and Sharpless, 2014). Due to the absence of free termini in the circular structure, circRNA can easily escape from hydrolysis by numerous cellular exonucleases. CircRNAs can be derived from exons, introns, or both, with a great diversity in length. Emerging evidence has shown that circRNAs are involved in many different biological processes and human diseases by functioning as miRNA sponges, RNA-binding protein sponges, transcriptional regulators, mRNA traps, regulators of assembly and transport of cellular proteins, as well as templates for translation.
Several computational methods, such as CIRI2, CIRCexplorer2, and find_circ, have been developed to identify circRNA by employing alignment-based strategies to recognize the back-spliced junction (BSJ), which is a unique feature of circRNAs (Szabo and Salzman, 2016; Cai et al., 2020). To confirm the presence of a circRNA identified in silico, an experimental approach is applied to validate the computationally predicted circRNAs (Zhang et al., 2016).
Reverse-transcription PCR (RT-PCR) is commonly used for the validation of circRNAs identified by RNA sequencing using divergent primers. In this approach, circRNA is converted to cDNA via reverse transcription (RT) with the presence of random hexamer or gene-specific reverse primers. Priming with random hexamers offers flexibility by ensuring RT of all RNA sequences, but gene-specific RT enhances the sensitivity of detecting the target of interest by converting the target of interest—instead of transcribing all RNA in the mix—into cDNA. Priming with gene-specific primers could enhance the detection of circRNA of interest, especially for those present in low abundance (Bustin and Nolan, 2004).
The conventional RT-PCR assay using convergent primers is unable to differentiate circRNAs from their linear transcript when the linear genome is used as template for primer design. Divergent primers are needed for the RT-PCR, as they allow direct detection of BSJ and quantification of circRNA. Divergent primers are a pair of outward-facing primers that allow amplification of circRNAs and not linear mRNA with the same sequence in cDNA and not gDNA (Figure 1). Sanger sequencing is usually performed on the PCR product amplified with the divergent primers to confirm the presence of a BSJ, as to rule out any incorrect amplification or artifact. Quantitative RT-PCR (RT-qPCR) can be used to relatively quantify the expression of circRNA across a panel of samples.
Figure 1. Primer design for circular RNA (circRNA) detection. Divergent primers are a pair of outward-facing primers that allow amplification of circRNA backsplice junction (BSJ), whereas convergent primers are a pair of inward-facing primers that allow amplification of linear mRNA from prepared cDNA.
Combining subcellular fractionation and RT-qPCR with divergent primers may provide some hints on circRNA’s biological functions. For example, circRNAs that are predominantly found in the cytoplasm are more likely to be involved in post-transcriptional gene regulation, whereas nuclear-retained circRNAs are predicted to play a role in transcription regulation. Currently available protocols are mainly for protein work, describing the steps for subcellular fractionation followed by western blot analysis, which is not entirely suitable for RNA work. Instead, this protocol allows the downstream detection of circRNA through RT-qPCR with divergent primers. A detailed protocol for the generation of specific circRNAs-enriched cDNA will also be included to better detect low-abundance circRNAs.
Materials and reagents
Biological material
GM12878 cell line
Reagents
Dulbecco’s phosphate-buffered saline (PBS) (Fisher Scientific, catalog number: BP399-1)
Trypsin-EDTA solution (0.5%) (Gibco, catalog number: 15400054)
Tissue culture media: Roswell Park Memorial Institute (RPMI) 1640 medium (Gibco, catalog number: 11875093)
Fetal bovine serum (FBS) (Gibco, catalog number: 10270106)
Penicillin-Streptomycin (Gibco, catalog number: 15140122)
Trypan blue (Alfa Aesar, catalog number: A1860)
Tris-Base (Fisher Scientific, catalog number: BP152-5)
Potassium chloride (KCl) (Fisher Scientific, catalog number: BP366-1)
Magnesium chloride (MgCl2) (Fisher Scientific, catalog number: BP214-500)
DL-Dithiothreitol (DTT) (Sigma-Aldrich, catalog number: D0632)
NP-40 (Sigma-Aldrich, catalog number: 9016-45-9)
Ribonucleoside vanadyl complexes (NEB, catalog number: S1402S)
Sodium acetate (Fisher Scientific, catalog number: BP333-500)
Molecular grade ethanol (Merck, catalog number: 64-17-5)
TRIzol (Life Technologies, catalog number: 15-596-018)
Chloroform:isoamyl alcohol (1:24) (Fisher Scientific, catalog number: BP1752I-400)
Molecular grade isopropanol (Fisher Scientific, catalog number: BP2618-1)
RNase-free water (Macherey Nigel, catalog number: 740378)
10× M-MuLV RT buffer (NEB, catalog number: #M0253)
DNase I (NEB, catalog number: #M0303)
M-MuLV reverse transcriptase (NEB, catalog number: #M0253)
RNase inhibitor (NEB, catalog number: #M0314)
dNTP mix (Thermo Scientific, catalog number: R0192)
Random hexamer (Invitrogen, catalog number: N8080127)
Primers (IDT, origin: USA)
SYBR Fast qPCR Master Mix (KAPA Biosystem, catalog number: KK4602)
Solutions
Complete media for GM12878 cell line (see Recipes)
Hypotonic buffer (see Recipes)
RNA precipitation solution (RPS) (see Recipes)
Recipes
Complete media for GM12878 cell line
Reagent Stock concentration Final concentration Volume (mL)
RPMI media 395
FBS 100% 20% 100
Penicillin-Streptomycin 10,000 U/mL 100 U/mL 5
Total 500
Hypotonic buffer
Reagent Stock concentration Final concentration Volume (μL)
Tris (pH 7.5) 100 mM 10 mM 500
KCl 100 mM 10 mM 500
MgCl2 150 mM 1.5 mM 50
DTT 100 mM 0.5 mM 25
NP-40 10% 0.075% 37.5
Ribonucleoside vanadyl complexes 200 mM 2 mM 50
H2O 3837.5
Total 5,000
RNA precipitation solution (RPS)
Reagent Stock concentration Final concentration Volume (mL)
Sodium acetate (pH 5.5) 3 M 150 mM 0.5
Ethanol 99.5% 9.5
Total 10
Final concentration of each component is 0.15 M sodium acetate (pH 5.5) in ethanol.
Keep at -20 °C for storage.
Equipment
Refrigerated tabletop centrifuge (15 or 50 mL conical tube adaptors) (Thermo Scientific, model: Multifuge X1R)
Refrigerated benchtop centrifuge (1.5 mL tube rotor) (Eppendorf, model: 5424R)
Light microscope (Olympus, origin: USA)
Vortexer (Biosan, model: V-1 plus)
Micropipettors (Gilson, origin: USA)
NanoDrop 2000c UV-Vis Spectrophotometer (Thermo Scientific, origin: USA)
Applied Biosystems Veriti Dx 96-Well Thermal Cycler PCR Thermocycler (Applied Biosystem, origin: USA)
Bio-Rad Connect Real-Time PCR System (Bio-Rad, origin: USA)
Procedure
Subcellular fractionation
Cell harvesting from culture plates
Grow the cells to a confluency of ~80%. Rinse the cells with 1× PBS.
For adherent cells, detach the cells with Trypsin-EDTA. For suspension cells, proceed to step A1d.
Add an equal volume of complete media to stop the trypsin proteolysis.
Collect the cell suspension in a conical tube and centrifuge at 200× g for 5 min.
Remove the supernatant and resuspend the cell pellet with 1× PBS.
Perform cell counting and aliquot 1 × 106 cells into a 1.5 mL microcentrifuge tube.
Centrifuge the tube at 200× g for 5 min and remove the supernatant.
Keep the cell pellet on ice prior to the next step.
RNA isolation from cytoplasmic and nuclear fractions of cells
Resuspend the cell pellet gently with 100 μL of ice-cold hypotonic buffer and incubate on ice for 5 min.
Note: Volume of the hypotonic buffer can be adjusted based on the size of cell pellet.
Check for membrane lysis with trypan blue. Proceed with centrifugation at 500× g for 10 min at 4 °C until > 90% of cells are lysed. Otherwise, increase the incubation time for step A2a until > 90% of cells are lysed.
Note: Do not overlyse the cells.
Collect the supernatant (cytoplasmic fraction) and wash the pellet (nuclear fraction) with 300 μL of ice-cold hypotonic buffer three times with centrifugation at 500× g for 5 min at 4 °C.
Add 1 mL of RPS to the cytoplasmic fraction and incubate at -20 °C for at least 1 h.
Vortex the cytoplasmic fraction in RPS and centrifuge at 18,000× g for 15 min at 4 °C.
Discard the supernatant and rinse the pellet with 70% (v/v) ice-cold ethanol.
Add 1 mL of TRIzol to the semi-dry nuclear and cytoplasmic pellets followed by the addition of 10 μL of 0.5 M EDTA.
Heat both fractions at 65 °C until the pellet dissolves with vortexing.
Cool the samples to room temperature and add 200 μL of chloroform:isoamyl alcohol (1:24).
Vortex the samples and centrifuge at 18,000× g for 10 min at room temperature.
Transfer the aqueous supernatant into a clean microcentrifuge tube.
Add an equal volume of isopropanol and incubate at -20 °C for at least 1 h.
Vortex the samples and centrifuge at 18,000× g for 15 min at room temperature.
Wash the pellet with 70% (v/v) ethanol and centrifuge at 18,000× g for 5 min.
Air-dry the pellet for 5–10 min.
Dissolve the air-dried RNA pellet in 30 μL of RNase-free water.
Quantify the RNA concentration and purity with NanoDrop 2000c UV-Vis spectrophotometer.
cDNA synthesis
DNase I treatment for isolated RNA
Prepare the RNA in PCR tube for DNase I treatment as shown in Table 1.
Table 1.Components used for DNase I treatment
Components Stock concentration Final concentration Volume (μL)
MuLV RT Buffer 10× 1× 1.4
DNase I 1 U 0.5
RNA 500–2,000 ng x
H2O Top up to 14
Incubate the tube in a thermal cycler at 37 °C for 30 min.
Reverse transcription (RT)
Prepare the RT reaction as shown in Table 2.
Table 2. Components used for reverse transcription
Components Stock concentration Final concentration Volume (μL)
Random hexamer Primer specific
MuLV RT Buffer 10× 1× 0.6 0.6
dNTP mix 10 mM 0.5 mM 1 1
Random hexamer 50 mM 2.5 mM 1 −
Reverse primers 10 μM 0.5 μM − 0.5 for each primer
RNase inhibitor 2 U 0.5 0.5
Reverse transcriptase 10 U 0.5 0.5
H2O 2.4 Top up to 6
Subtotal 6 6
DNAse I–treated RNA 14 14
Total 20 20
Perform the cDNA conversion on a thermal cycler with the setup of 42 °C for 60 min and follow by heat inactivation at 65 °C for 20 min.
Notes:
i. If more than four pairs of primer sets (targeting linear mRNAs and/or circRNAs) are needed for primer-specific cDNA conversion, prepare 5 μM primer mix and add 1 μL into the RT reaction.
ii. One may start with random hexamer for the cDNA conversion and switch to primer-specific cDNA conversion if the amplification of the targeted circRNA BSJs is weak, absent, or unspecific.
iii. Random hexamer-generated cDNA allows amplification of any circRNA target of interest at any one time, while primer-specific-generated cDNA only allows amplification of the planned circRNA targets and does not allow any amplification of any unplanned circRNA targets.
iv. Caution is needed when generating circRNA-specific cDNA, as the remaining unused reverse primer in the RT reaction could possibly interrupt or participate in qPCR process and lead to undesired amplification from linear mRNA.
Quantitative PCR (qPCR)
Prepare the PCR reaction as shown in Table 3.
Table 3.Components used for SYBR Green RT-qPCR
Components Stock concentration Final concentration Volume (μL)
SYBR Fast qPCR Master Mix 2× 1× 5
Forward primer 10 μM 0.1 μM 0.1
Reverse primer 10 μM 0.1 μM 0.1
6-fold diluted cDNA 3
RNase-free water 1.8
Total 10
Perform the qPCR on a real time thermal cycler with the cycling parameter of 95 °C for 3 min, followed by 40 cycles of 95 °C for 2 s and 60 °C for 20 s.
Calculate the relative expression of each gene based on the equation below:
RNA ratio is the ratio of cytoplasmic RNA concentration to nuclear RNA concentration eluted in a similar volume of water. Gene expression for a cytoplasmic marker is calculated using formula (a), whereas gene expression of a nuclear marker is calculated using formula (b).
Data analysis
Representative data
Subcellular fractionation was performed using GM12878 cells, which is a lymphoblastoid cell line that contains Epstein-Barr Virus (EBV). The concentration and purity of each fraction were measured using NanoDrop 2000c (Table 4). The cytoplasmic RPL30 and nuclear MALAT1 transcripts were used as positive controls to indicate the purity of cytoplasmic and nuclear fractions, respectively. As shown in Figure 2, RPL30 was enriched in the cytoplasm, whereas MALAT1 was enriched in the nuclear fraction, indicating that subcellular fractionation was successfully performed. In this representative data, the linear EBV latent membrane protein 2 (LMP-2) transcripts were found in both cytoplasmic and nuclear fractions. The subcellular localization pattern of circLMP-2_e5 is similar to that of linear LMP-2A and LMP-2B.
Note that the protocol described here requires a real-time PCR machine to generate the data, which may not be available in all laboratories. In such a case, conventional agarose gel–based RT-PCR can be conducted to obtain a semi-quantitative result.
Table 4. Concentration and purity of cytoplasmic and nuclear fraction
Concentration (ng/μL) 260/280 260/230
Cytoplasmic fraction 1 1,279 1.96 1.95
Cytoplasmic fraction 2 1,242 1.93 1.74
Nuclear fraction 1 566 1.76 0.71
Nuclear fraction 2 430.3 1.87 1.46
1 and 2 represent biological replicates.
Figure 2. Subcellular localization of circLMP-2_e5. RT-qPCR analysis showed LMP-2 and circLMP-2_e5 were localized in both nucleus and cytoplasm of GM12878 cells. Data was normalized to the RNA yield ratio and represents the mean ± SD of two independent experiments. This result was published in Figure 3C in Tan et al. (2021).
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Tan et al. (2021). Identification and characterization of a novel Epstein-Barr Virus-encoded circular RNA from LMP-2 Gene. Scientific Reports (Figure 3, panel C).
Acknowledgments
The work was supported financially by the Ministry of Higher Education Malaysia via Fundamental Research Grant Scheme (FRGS/1/2017/SKK08/UM/02/11) and University of Malaya High Impact Research Grant (UM.C/625/1/HIR/MOE/CHAN/02/07).
The original research paper in which this protocol was described and validated is Tan et al. (2021).
Competing interests
The authors declare no competing interests.
References
Bustin S. A. and Nolan, T. (2004). Pitfalls of quantitative real-time reverse-transcription polymerase chain reaction. J. Biomol. Tech. 15(3): 155–166.
Cai, Z., Fan, Y., Zhang, Z., Lu, C., Zhu, Z., Jiang, T., Shan, T. and Peng, Y. (2020). VirusCircBase: a database of virus circular RNAs. Briefings Bioinf. 22(2): 2182–2190.
Gagnon, K. T., Li, L., Janowski, B. A. and Corey, D. R. (2014). Analysis of nuclear RNA interference in human cells by subcellular fractionation and Argonaute loading. Nat. Protoc. 9(9): 2045–2060.
Jeck, W. R. and Sharpless, N. E. (2014). Detecting and characterizing circular RNAs. Nat. Biotechnol. 32(5): 453–461.
Szabo, L. and Salzman, J. (2016). Detecting circular RNAs: bioinformatic and experimental challenges. Nat. Rev. Genet. 17(11): 679–692.
Tan KE, Ng WL, Marinov GK, Yu KH, Tan LP, Liau ES, Goh SY, Yeo KS, Yip KY, Lo KW et al. (2021). Identification and characterization of a novel Epstein-Barr Virus-encoded circular RNA from LMP-2 Gene. Sci Rep. 2021 Jul 13;11(1):14392.
Zhang, Y., Yang, L. and Chen, L. L. (2016). Characterization of Circular RNAs. In: Feng, Y. and Zhang, L. (Eds.). Long Non-Coding RNAs (pp. 215–227). Methods in Molecular Biology. Humana Press, New York.
Article Information
Copyright
© 2023 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
Molecular Biology > RNA > qRT-PCR
Cell Biology > Organelle isolation > Fractionation
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4,799 | https://bio-protocol.org/en/bpdetail?id=4799&type=0 | # Bio-Protocol Content
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Peer-reviewed
Confocal Imaging and 3D Reconstruction to Determine How Genetic Perturbations Impact Presynaptic Morphology at the Mouse Calyx of Held
CK Christian Keine
TR Tamara Radulovic
MA Mohammed Al-Yaari
SJ Samuel M. Young Jr.
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4799 Views: 820
Reviewed by: Alessandro DidonnaCarolina Tecuatl Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in eLIFE Oct 2022
Abstract
Neurons communicate via synapses—specialized structures that consist of a presynaptic terminal of one neuron and a postsynaptic terminal of another. As knowledge is emerging that mutations in molecules that regulate synaptic function underpin many neurological disorders, it is crucial to elucidate the molecular mechanisms regulating synaptic function to understand synaptic strength, plasticity, modulation, and pathology, which ultimately impact neuronal circuit output and behavior. The presynaptic calyx of Held is a large glutamatergic presynaptic terminal in the auditory brainstem, which due to its accessibility and the possibility to selectively perform molecular perturbations on it, is an ideal model to study the role of presynaptic proteins in regulating synaptic function. In this protocol, we describe the use of confocal imaging and three-dimensional reconstruction of the calyx of Held to assess alterations in gross morphology following molecular perturbation. Using viral-vector delivery to perform molecular perturbations at distinct developmental time points, we provide a fast and cost-effective method to investigate how presynaptic proteins regulate gross morphology such as surface area and synapse volume throughout the lifetime of a neuronal circuit.
Key features
• Confocal imaging and 3D reconstruction of presynaptic terminals.
• Used with a virus-mediated expression of mEGFP to achieve efficient, cell-type specific labeling of the presynaptic compartment.
• Protocol was developed with the calyx of Held but is suitable for pre- and postsynaptic compartments of various neurons across multiple mammalian and invertebrate species.
Keywords: Calyx of Held Presynaptic terminal Confocal imaging Three-dimensional reconstruction Transcardial perfusion Synaptic morphology
Background
The brain consists of billions of synapses, and many synaptic molecules have multiple roles regulating synapse morphology and synaptic transmission to control brain function (Shen and Cowan, 2010; Rosenberg et al., 2014). Therefore, combining physiological and imaging experiments provides powerful insight into the complex relationship between cellular structure and function (Marrone and Petit, 2002; Rollenhagen and Lübke, 2006; Wichmann and Moser, 2015; Sierksma et al., 2020). Genetic perturbations with viral-vector approaches allow for the targeted expression of different synaptic proteins in the pre- or postsynaptic cell in combination with a fluorescent reporter. Since the fluorescent reporter can readily be identified using light microscopy, this enables the use of laser scanning confocal microscopy in conjunction with the quantification of fluorescent signals in three-dimensional image stacks to examine the gross morphology of specific pre- or postsynaptic compartments at a fine scale. Here, we describe the use of confocal imaging of the calyx of Held presynaptic terminal followed by three-dimensional reconstruction to assess alterations in gross morphology following molecular perturbations (Montesinos et al., 2015; Radulovic et al., 2020; Keine et al., 2022). In combination with viral-vector delivery to perform targeted molecular perturbation at distinct developmental time points, this method allows for fast and cost-effective analysis of synaptic gross morphology of presynaptic terminals at different developmental stages. This protocol was developed with the mouse calyx of Held but it is applicable to other preparations and a wide range of model organisms, mammalian and invertebrate, where fluorescent labeling of the target structure can be achieved.
Materials and reagents
Biological materials
Laboratory mice (Rac1tm1Djk/J, P28, either sex) (Jackson Laboratory, catalog number: 005550)
Reagents
Sodium phosphate monobasic (Sigma-Aldrich, catalog number: S0751)
Sodium phosphate dibasic (Millipore, catalog number: 567550)
2,2,2-Tribromoethanol (Sigma-Aldrich, catalog number: T48402)
2-Methyl-2-butanol (Sigma-Aldrich, catalog number: 152463)
Paraformaldehyde (Sigma-Aldrich, catalog number: 158127)
Sodium hydroxide solution 1 N (Sigma-Aldrich, catalog number: S2770)
Hydrochloric acid solution 1 N (Sigma-Aldrich, catalog number: H9892)
Solutions
Phosphate buffer (PB) 0.1 M (see Recipes)
Phosphate buffer (PB) 0.5 M (see Recipes)
Fixative solution (4% paraformaldehyde in 0.1 M PB, pH 7.4) (see Recipes)
Avertin (see Recipes)
Recipes
Phosphate buffer (PB) 0.1 M
Reagent Final concentration Quantity
Sodium phosphate monobasic (solution A) 0.1 M 0.6 g in 50 mL of ultrapure H2O
Sodium phosphate dibasic (solution B) 0.1 M 2.4 g in 170 mL of ultrapure H2O
Total 0.1 M 220 mL
Prepare 50 mL of 0.1 M sodium phosphate monobasic solution by dissolving 0.6 g of sodium phosphate monobasic (MW = 119.98 g/mol) in 50 mL of ultrapure water (solution A) under constant stirring.
Prepare 170 mL of 0.1 M sodium phosphate dibasic solution by dissolving 2.4 g of sodium phosphate dibasic (MW = 141.96 g/mol) in 170 mL of ultrapure water (solution B) under constant stirring.
Slowly pour solution A into solution B while stirring and monitoring pH.
Stop when pH settles at 7.3–7.4.
Filter with a sterile bottle-top filter and store at 4 °C until ready to use for up to one month.
Phosphate buffer (PB) 0.5 M
Reagent Final concentration Quantity
Sodium phosphate monobasic (solution A) 0.5 M 3 g in 50 mL of ultrapure H2O
Sodium phosphate dibasic (solution B) 0.5 M 14 g in 170 mL of ultrapure H2O
Total 0.5 M 220 mL
Prepare 50 mL of 0.5 M sodium phosphate monobasic solution by dissolving 3 g of sodium phosphate monobasic (MW = 119.98 g/mol) in 50 mL of ultrapure water (solution A) under constant stirring.
Prepare 170 mL of 0.5 M sodium phosphate dibasic solution by dissolving 12 g of sodium phosphate dibasic (MW = 141.96 g/mol) in 170 mL of ultrapure water (solution B) under constant stirring.
Slowly pour solution A into solution B while stirring and monitoring pH.
Stop when pH settles at 7.3–7.4.
Filter with a sterile bottle-top filter and store at 4 °C until ready to use for up to one month.
Fixative solution (4% paraformaldehyde in 0.1 M PB, pH 7.4)
Reagent Final concentration Quantity
Paraformaldehyde 4% 4 g
Ultrapure H2O n/a 100 mL
Total n/a 100 mL
The process of preparing PFA solution should take place in a chemical fume hood.
Measure 70 mL of ultrapure water and heat to 55–60 °C while stirring on a hotplate stirrer. Avoid heating the solution above 65 °C, as this will degrade PFA.
Slowly add 4 g of paraformaldehyde.
Let stir and keep adding 1 N NaOH drop by drop until the paraformaldehyde is dissolved and the solution is clear. Let solution cool down.
Add 20 mL of 0.5 M PB and bring the final volume to 100 mL.
Adjust pH to 7.4 with 1 N HCl and 1 N NaOH.
Filter with a sterile bottle-top filter and store at 4 °C for up to one week or in 50 mL aliquots at -20 °C for up to one month.
Avertin
Reagent Final concentration Quantity
2,2,2-Tribromoethanol 44 mM 250 mg
2-methyl-2-butanol n/a 0.5 mL
Ultrapure H2O n/a 19.5 mL
Total n/a 20 mL
Dissolve 250 mg of 2,2,2-Tribromoethanol into 0.5 mL of 2-methyl-2-butanol at 40 °C and while stirring. Make sure not to exceed 40 °C or the solution might degrade.
Add ultrapure water to a final volume of 20 mL, filter with a sterile syringe filter, and store in 0.5 mL aliquots at -20 °C for up to one year. Protect from light by using amber/brown tubes and store in a separate light-protected box. Do not refreeze unused Avertin as it might reduce the anesthesiologic effect. Discard the solution and prepare fresh if any of the following conditions occur:
i. Expiration date (one year) exceeded.
ii. Solution has turned yellow in color.
iii. Solution started to crystallize.
Laboratory supplies
Magnetic stirring bars (VWR, catalog number: 442-0368)
Bottle-top filter (Thermo Fisher, Nalgene RapidFlow, catalog number: 596-3320)
Syringe filter (Millipore Millex, 0.22 μm, catalog number: SLGPR33RS)
Single-use hypodermic needles (Braun Sterican, 25G, 25 mm)
Three-way stop cock (GPC Medical, catalog number: DIS122)
Forceps (Fine Science Tools, Dumont #3, catalog number: 11231-30)
Extra Fine Bonn scissors (Fine Science Tools, catalog number: 14084-08)
Bonn scissors (Fine Science Tools, catalog number: 14184-09)
Surgical scissors (Fine Science Tools, catalog number: 14000-13)
12-well plate (CytoOne, catalog number: CC7682-7512)
Aqua-Poly/Mount (Polysciences, catalog number: 18606-20)
Microscope slides (Fisher Scientific, Fisherbrand Superfrost Plus, catalog number: 22-037-246)
Cover glass (VWR, catalog number: 48404-453)
Razor blades (Procter & Gamble, Astra Superior Platinum Double Edge)
Superglue (3M, Scotch)
Modeling clay (VWR, catalog number: 470149-616)
Zirconia ceramic blades (Cadence Blades, catalog number: EF-INZ10)
Equipment
Laboratory hotplate stirrer (VWR, catalog number: 442-1271)
Dissection tray (Fisher Scientific, Epredia Shandon, catalog number: 73092)
Mechanical pipette 0.5–10 μL (Eppendorf, Research Plus, catalog number: 3123000071)
Mechanical pipette 10–100 μL (Eppendorf, Research Plus, catalog number: 3123000047)
Digital pump (Ismatec, MS-4/12, catalog number: ISM597D)
Vibrating blade microtome (Leica Biosystems, VT1200 S, catalog number: 14048142066)
Confocal microscope (LSM 700, AxioObserver, Carl Zeiss Microscopy)
Software
ZEN blue (Carl Zeiss Microscopy, v3, RRID:SCR_013672)
Commercial software
Minimum hardware requirements: 3 GHz Intel i5 quad-core CPU, 4 GB RAM, 32 bit graphics adapter with 4 GB RAM
Imaris Microscopy Image Analysis Software (Bitplane, Oxford Instruments, v10.0, RRID:SCR_007370)
Used with Imaris Measurement Pro feature
Commercial software
Minimum hardware requirements: 3 GHz dual core CPU (64-bit), 8 GB RAM, NVIDIA Quadro P400 Graphics Card with 2 GB RAM
Alternative software: Amira (RRID:SCR_007353)
Free alternative software: Fiji/ImageJ, RRID:SCR_002285)
MATLAB (MathWorks, v9.12, RRID:SCR_001622)
Commercial software
Minimum hardware requirements: Intel or AMD x86-64 processor, 4 GB RAM
Free alternative software: Python (RRID:SCR_008394), R (RRID:SCR_001905)
Procedure
Transcardial perfusion
Note: A detailed protocol on transcardial perfusion including images and further methodological considerations can be found in Wu et al. (2021).
The transcardial perfusion should take place in a chemical fume hood.
Prepare solutions (PB and 4% PFA), pour into glass beakers, and connect to pump. Prefill tubing with respective solutions and remove air bubbles. Switching between solutions (step A15) can be aided by using a three-way stop cock connected to PB, 4% PFA, and the injection needle.
Weigh the animal.
Deeply anesthetize the animal via intraperitoneal injection of Avertin (250 mg/kg body weight).
Position the mouse in a supine position on a dissection tray and secure the limbs with clamps or laboratory tape.
Locate the most caudal end of the sternum indicating the caudal end of the rib cage.
Make a midline incision into the abdominal cavity through the skin and muscle layers just caudal of the rib cage.
Open thorax cavity by cutting through the diaphragm, taking care not to damage the heart.
Cut along the lateral wall on either side of the rib cage and lift it to expose the heart.
Locate the left ventricle and right atrium. The right atrium has a characteristic dark red color. The left ventricle forms the tip of the heart.
Insert a 25-gauge needle into the caudal portion of the left ventricle at a flat angle, being careful not to puncture the septum.
Make a small incision in the right atrium with scissors so that rapid blood flow occurs.
Turn on the pump to flush ice-cold 0.1 PB into the left ventricle at a flow rate of 1–2 mL/min and keep the needle in place using modeling clay or laboratory tape. The blood should be washed out through the right atrium.
Continue to flush with PB until the mouse’s body weight has been flushed three times and the liquid leaving the right atrium is clear and free of blood (e.g., if the animal’s weight is 10 g, inject a total volume of 30 mL of PB). Examine the liver; it should change color from dark red to pale yellow or white.
Inject fixative (4% PFA) using a three-way stop cock, making sure that the needle stays in place. Double-check tubing to make sure there are no air bubbles present. Organs and muscles should start to turn stiff after a few minutes.
Allow the fixative to run until at least three times the mouse’s body weight has been injected (e.g., if the animal’s weight is 10 g, inject a total volume of 30 mL of fixative). Liver, limbs, and tail should then be stiff.
Stop the pump and remove the needle.
Decapitate the animal with surgical scissors.
Carefully remove the brain from the skull, taking care not to touch the area of interest (e.g., cochlear nucleus or MNTB). Cut larger nerves with fine scissors instead of tearing them to minimize damage to the brain.
Using forceps, carefully remove the meninges from the ventral surface of the brainstem.
Immerse brain in 4% PFA in a 12-well plate at 4 °C overnight. The volume of PFA solution used should be at least three times the volume of the brain (e.g., > 1.5 mL for an adult mouse brain). Make sure the whole brain is fully submerged in the solution.
After 12 h of post fixation, discard PFA and replace it with PB. The volume of PB solution used should be at least three times the volume of the brain (e.g., > 1.5 mL for an adult mouse brain). Make sure the whole brain is fully submerged in the solution. Keep the brain immersed in PB at 4 °C until further use. For best results, use the brain within a few hours after being placed in PB. Storing the brain in PB for several days might reduce fluorescence and impact imaging quality.
Slicing of fixed brain and mounting of brain slices
Remove the brain from PB and excise the rostral half of the cerebrum with razor blades.
Using super glue, adhere the brain on the cut surface to the specimen holder. Handle the brain with caution to avoid touching the brainstem.
Immerse the brain into PB and orient the specimen holder so that the ventral surface of the brainstem faces the vibrating microtome blade.
Using the microtome, slice the brain at a thickness of 40 μm, moving the blade at a rate of 20–50 μm/s.
Identify the region of interest and collect the slices containing this region in PB solution.
Mount the slices on microscope slides and remove any excess PB. Allow the slices to dry almost completely; then, add one drop of Aqua-Poly/Mount to each slice, taking care to avoid the formation of air bubbles. Cover the slice with a cover glass.
Allow Aqua-Poly/Mount to harden for 24 h at room temperature. Store completed samples at 4 °C in the dark when not in use. Properly prepared samples can be stored for several months and imaged multiple times.
Confocal imaging of presynaptic terminals
Locate the MNTB region containing EGFP-labeled calyx terminals using a low-magnification objective. Select calyx terminals that are well separated from neighboring terminals for subsequent reconstruction.
Switch to a 63× oil immersion objective lens (Plan-Apochromat 63×/1.4 Oil DIC M27) and select an imaging area that contains multiple well-separated calyx terminals.
Image the membrane-bound EGFP expressed in the cells using an excitation wavelength of 488 nm and an emission wavelength of 518 nm.
Select the Scan Mode Frame and set the image size to 1,024 × 1,024 pixels.
Set Speed to 8 and Averaging to 4 lines. Set bit depth to 16 bit and the scan mode to bidirectional.
Set the pixel size to 0.1 μm × 0.1 μm × 0.35 μm.
Optimize the microscopy image by adjusting the laser power, gain, and offset. Images suitable for 3D reconstruction will have low background noise, high contrast, and are not saturated (Figure 1). The fluorescence signal should be within the range of the detector. Use the range indicator to adjust laser power and gain. If saturated areas are detected, lower laser power and gain until there are no saturated pixels in the image.
Figure 1. Adjusting laser power and signal gain to improve confocal images. When acquiring confocal images, the fluorescence signal should be within the range of the detector. (A) Signal too weak, structural details are lost due to low laser power and signal gain. (B) When laser power and signal gain are adjusted, structural details become visible and calyceal terminals can be imaged. (C) Setting laser power and signal gain too high will result in saturation of image details and loss of structural details. Images were obtained using a 63× oil immersion objective.
Set the imaging depth in z-axis so that the whole presynaptic terminal and at least 5 μm on either end are within the imaging volume.
3D reconstruction of presynaptic terminals
Start Imaris, select Arena, and drag and drop the image file containing the acquired z-stack into Arena.
Double-click on the file to convert it to IMS format.
Double-click to open the IMS file.
Inspect the image volume in 3D View and locate a well-isolated calyx terminal that is fully captured within the image volume.
Select Edit > Crop 3D and crop image volume to contain the selected calyx terminal. Make sure to crop the calyx as close as possible including 1 μm margins around the calyx volume. Then, click OK. The 3D View should now contain the selected calyx terminal only.
In the Surpass tree tab on the left, select Add new Surfaces.
Select the newly created surface object. In the Create tab, select Skip automatic creation, edit manually.
Click Settings and select Render on Slicer.
Use the slider Slice Position in the Draw tab to select a slice in the middle of the image volume in which the calyx terminal is clearly visible.
In the Mode tab under Drawing Mode, select Magic wand, click Draw.
Using the mouse cursor, create line-abound surfaces with the same pixel intensity.
When not in Draw mode, vertices of the detected surface can be re-positioned with the mouse while holding down the T button and deleted using the D button.
Adjust the surface area carefully to properly represent the dimensions of the calyx terminal.
When satisfied with the selection, select the next slice from the Slice position slider and repeat steps D9–D13. Repeat this process for all slices until the whole calyx terminal is selected (Figure 2).
Figure 2. 3D reconstruction of the calyx of Held presynaptic terminal. (A) Volumetric image containing mEGFP-labeled calyx terminals of a P28 mouse. Rectangle indicates the calyx terminal in B–E. Scale bar: 20 μm. (B) Volumetric image of single calyx terminal after three-dimensional cropping. Scale bar: 5 μm. (C) Single section through the middle of the calyx terminal shown in B. Scale bar: 5 μm. (D) Detection of surface borders (cyan) in the single section. Scale bar: 5 μm. (E) Full 3D reconstruction of fluorescent labeled calyx. Scale bar: 5 μm.
Click Create Surface at the bottom of the Draw tab and turn off Render on Slicer.
Using the tools in the Edit tab, the generated surface can be visually inspected and verified.
If multiple surfaces have been created that should be merged into a single surface, select the surfaces to merge and click the Unify button located in the Edit tab.
In the Statistics tab, select Selection. In the drop-down menu, select Specific Values. Then, select Area and Volume from the drop-down menu below. Data can be exported to CSV or XLS format using the Export Statistics buttons.
Data analysis
Presynaptic surface area and volume were extracted from the 3D reconstructions and compared between treatment and control animals. To quantify the calyx’s overall shape, surface-area-to-volume ratios can be calculated by dividing the calyx’s surface area by its volume. Additional measures such as sphericity can be calculated and exported from Imaris but should be used with caution and depending on the expected shape of the structure of interest. Individual calyx terminals were considered independent samples, and calyx terminals that expressed only GFP served as control. Statistical analysis was conducted in MATLAB, but other statistical analysis software (e.g., R, GraphPad Prism, SPSS) can be used. Data distributions were tested for Gaussianity using the Shapiro-Wilk test (function swtest). Comparison of two groups was performed using a two-tailed unpaired Student’s t-test with Welch’s correction (normal distribution, function ttest2) or a two-tailed Mann-Whitney U test (non-normal distribution, function ranksum). Effect sizes were calculated using the MES toolbox in MATLAB (Hentschke and Stuttgen, 2011). Calyx terminals should be sampled from the same MNTB region in treatment and control mice to minimize bias introduced by location (Ford et al., 2009). The final dataset should contain a minimum of 15 calyces from at least three different animals per group, to minimize inter-individual differences. When using Cre-expressing viral vectors in combination with conditional knock-out mice to ablate synaptic proteins, animals were injected with viral vectors expressing either mEGFP (control) or both Cre recombinase and mEGFP (knock-out).
Validation of protocol
This protocol has been validated independently by multiple researchers and results have been published in peer-reviewed publications (Montesinos et al., 2015; Radulovic et al., 2020; Keine et al., 2022). The procedure can be applied to different developmental stages and has been successfully applied in young, juvenile, and young adult mice. When using viral vectors to express Cre recombinase in combination with mEGFP to ablate proteins in conditional knock-out animal models, sufficient expression of Cre-recombinase should be validated by injecting the virus into a suitable Cre-reporter mouse line (e.g., B6.Cg-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J, RRID:IMSR_JAX:007909) with Cre-dependent expression of a fluorescent protein. To assess the effectivity and time course of Cre-mediated protein ablation, mRNA or protein levels can be quantified with qRT-PCR, Western plot, or immunohistochemistry in either cell cultures or brain slices.
General notes and troubleshooting
General notes
All surgical procedures should be performed in accordance with a protocol approved by the respective Institutional Animal Care Use Committee. Slight variations in anesthesia, surgical procedures, pain management, and post-surgical care and follow-up may be required. Experiments have been performed in Rac1tm1Djk/J mice at age P28, but the protocol can be applied to other laboratory animals and developmental stages if mEGFP can be expressed in the target structure. However, large structures are more suited for this approach due to the resolution limit of confocal imaging. Subtle changes in cell morphology might go unnoticed using confocal imaging and require the use of super resolution or electron microscopy. All reagents are of molecular biology grade and should be stored according to the manufacturer’s recommendation. All solutions are prepared using ultrapure water (> 18 MΩ·cm at 25 °C) unless noted otherwise. For best results, solutions should be prepared daily. Solutions should be stored at 4 °C or -20 °C and can be used for one week or up to one month, respectively. Avoid multiple (> 3) freeze-thaw cycles.
Troubleshooting
Problem observed Possible reason Solution
During perfusion, lungs expand, fluid outflow from nose/mouth Heart septum pierced with needle Withdraw needle and insert at flat angle
After perfusion, brain contains blood, liver does not turn pale Ineffective washout of blood due to air bubbles in perfusion system Remove air bubbles in the perfusion system
Ineffective washout due to blood clotting Reduce anesthesia dose to avoid cardiac arrest and minimize time between anesthesia and perfusion
After perfusion, blood is washed out, but brain remains soft 4% PFA solution not working properly Make sure to fully dissolve PFA, avoid overheating (> 60 °C), and use fresh solution
Images oversaturated with no details visible Laser power or gain too high during imaging Lower laser power and gain, use range indicator
During reconstruction, the calyx borders cannot be determined properly Overlap of passing axons or neighboring terminals Use well-isolated calyx terminal for reconstruction
Acknowledgments
We thank the members of the Young lab for their comments on the manuscript. This protocol was derived from and validated in the original research papers (Montesinos et al., 2015; Radulovic et al., 2020; Keine et al., 2022). The work has been supported by grants from NIDCD (R01 DC014093), NINDS (R01 NS110742), and NCATS (R03TR004161-0) to S.M.Y. and a postdoctoral fellowship from the DFG (420075000) to C.K.
Competing interests
The authors declare no competing interest. The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.
Ethical considerations
All experiments were performed following animal welfare laws and approved by the Institutional Committee for Care and Use of Animals at the University of Iowa PHS Assurance No. D16- 00009 (A3021- 01) (Animal Protocol 0021952) and complied with accepted ethical best practices.
References
Ford, M. C., Grothe, B. and Klug, A. (2009). Fenestration of the calyx of held occurs sequentially along the tonotopic axis, is influenced by afferent activity, and facilitates glutamate clearance. J. Comp. Neurol. 514(1): 92–106.
Hentschke, H. and Stüttgen, M. C. (2011). Computation of measures of effect size for neuroscience data sets. Eur. J. Neurosci. 34(12): 1887–1894.
Keine, C., Al-Yaari, M., Radulovic, T., Thomas, C. I., Valino Ramos, P., Guerrero-Given, D., Ranjan, M., Taschenberger, H., Kamasawa, N., Young, S. M., et al. (2022). Presynaptic Rac1 controls synaptic strength through the regulation of synaptic vesicle priming. eLife 11: e81505.
Marrone, D. F. and Petit T. L. (2002). The role of synaptic morphology in neural plasticity: structural interactions underlying synaptic power. Brain Res. Rev. 38(3): 291–308.
Montesinos, M. S., Dong, W., Goff, K., Das, B., Guerrero-Given, D., Schmalzigaug, R., Premont, R. T., Satterfield, R., Kamasawa, N., Young, S. M., et al. (2015). Presynaptic Deletion of GIT Proteins Results in Increased Synaptic Strength at a Mammalian Central Synapse. Neuron 88(5): 918–925.
Radulovic, T., Dong, W., Goral, R. O., Thomas, C. I., Veeraraghavan, P., Montesinos, M. S., Guerrero‐Given, D., Goff, K., Lübbert, M., Kamasawa, N., et al. (2020). Presynaptic development is controlled by the core active zone proteins CAST/ELKS. J. Physiol. 598(12): 2431–2452.
Rollenhagen, A. and Lübke, J. H. R. (2006). The morphology of excitatory central synapses: from structure to function. Cell Tissue Res. 326(2): 221–237.
Rosenberg, T., Gal-Ben-Ari, S., Dieterich, D. C., Kreutz, M. R., Ziv, N. E., Gundelfinger, E. D. and Rosenblum, K. (2014). The roles of protein expression in synaptic plasticity and memory consolidation. Front. Mol. Neurosci. 7: e00086.
Shen, K. and Cowan, C. W. (2010). Guidance Molecules in Synapse Formation and Plasticity. Cold Spring Harbor Perspect. Biol. 2(4): a001842.
Sierksma, M. C., Slotman, J. A., Houtsmuller, A. B. and Borst, J. G. G. (2020). Structure–function relation of the developing calyx of Held synapse in vivo. J. Physiol. 598(20): 4603–4619.
Wichmann, C. and Moser, T. (2015). Relating structure and function of inner hair cell ribbon synapses. Cell Tissue Res. 361(1): 95–114.
Wu, J., cai, y., Wu, X., Ying, Y., Tai, Y. and He, M. (2021). Transcardiac Perfusion of the Mouse for Brain Tissue Dissection and Fixation. Bio Protoc 11(5): e3988.
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Cell Biology > Cell imaging > Confocal microscopy
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Peer-reviewed
Methods to Quantify the Dynamic Recycling of Plasma Membrane Channels
Rawad Hodeify
KM Khaled Machaca
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4800 Views: 680
Reviewed by: Emilia KrypotouXin Xu Anonymous reviewer(s)
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Bio-protocol journal peer-reviewed
Sep 05, 2023 | This version
Preprint
Oct 27, 2022
Original Research Article:
The authors used this protocol in Science Advances Sep 2018
Abstract
Store-operated Ca2+ entry (SOCE) is a ubiquitous Ca2+ signaling modality mediated by Orai Ca2+ channels at the plasma membrane (PM) and the endoplasmic reticulum (ER) Ca2+ sensors STIM1/2. At steady state, Orai1 constitutively cycles between an intracellular compartment and the PM. Orai1 PM residency is modulated by its endocytosis and exocytosis rates. Therefore, Orai1 trafficking represents an important regulatory mechanism to define the levels of Ca2+ influx. Here, we present a protocol using the dually tagged YFP-HA-Orai1 with a cytosolic YFP and extracellular hemagglutinin (HA) tag to quantify Orai1 cycling rates. For measuring Orai1 endocytosis, cells expressing YFP-HA-Orai1 are incubated with mouse anti-HA antibody for various periods of time before being fixed and stained for surface Orai1 with Cy5-labeled anti-mouse IgG. The cells are fixed again, permeabilized, and stained with Cy3-labeled anti-mouse IgG to reveal anti-HA that has been internalized. To quantify Orai1 exocytosis rate, cells are incubated with anti-HA antibody for various incubation periods before being fixed, permeabilized, and then stained with Cy5-labeled anti-mouse IgG. The Cy5/YFP ratio is plotted over time and fitted with a mono-exponential growth curve to determine exocytosis rate. Although the described assays were developed to measure Orai1 trafficking, they are readily adaptable to other PM channels.
Key features
• Detailed protocols to quantify endocytosis and exocytosis rates of Orai1 at the plasma membrane that can be used in various cell lines.
• The endocytosis and exocytosis assays are readily adaptable to study the trafficking of other plasma membrane channels.
Graphical overview
Keywords: Endocytosis Exocytosis Trafficking Orai1 Internalization Plasma membrane Immunofluorescence Recycling Store-operated Ca2+ entry
Background
Orai1 is the pore-forming subunit of the store-operated Ca2+ entry (SOCE) channel. It mediates Ca2+ influx into the cell following direct coupling with ER Ca2+ sensor STIM1, in response to store depletion following Ca2+ release from stores (Luik et al., 2006; Prakriya et al., 2006; Vig et al., 2006; Yeromin et al., 2006). SOCE is central to various physiological processes, and its dysfunction is associated with serious pathologies in humans (Feske, 2011, Feske et al., 2015; Gruszczynska-Biegala et al., 2021; Yu et al., 2021). A key feature in SOCE regulation is the residence of Orai1 at the plasma membrane (PM) and the amount of reserve Orai1 in intracellular compartments that can potentially contribute to the PM pool following SOCE activation.
The steady-state amount of Orai1 at the PM is determined by the rates of endocytosis and exocytosis. Alterations in these rates modulate Orai1 PM levels. Several protocols have been used to quantify the amount of surface and intracellular Orai1, including imaging of fluorescently tagged Orai1 and biotinylation (Yu et al., 2009; Cox et al., 2013; Yeh et al., 2020; Kim et al., 2021; Wrennall et al., 2022). These methodologies provide crude estimates of Orai1 distribution, but they lack the sensitivity and time resolution to determine Orai1 recycling kinetics at the PM. Herein, we describe protocols to quantify the kinetics of Orai1 trafficking using a dually tagged human Orai1 (YFP-HA-Orai1) with a cytosolic YFP at the N-terminus and a hemagglutinin (HA) tag inserted in the second extracellular loop (Park et al., 2009; Hodeify et al., 2015 and 2018). This dual reporter allows quantification of total Orai1 using YFP fluorescence and concomitantly follows its trafficking using the HA-antibody at the single-cell level. The PM and intracellular pools can be differentially labeled by staining unpermeabilized and permeabilized cells with different secondary antibodies (Cy5- or Cy3-labeled). Combining these approaches with time courses of anti-HA antibody feeding allows one to carefully quantify Orai1 endocytosis and exocytosis rates. The presented protocols can be easily adapted to study recycling dynamics of PM channels in a wide range of mammalian cells.
Materials and reagents
Biological materials
TRVb-1 cells, a CHO cell line lacking the endogenous transferrin receptor and stably expressing the human transferrin receptor (McGraw et al., 1987). Any readily transfectable cell line can be substituted for CHO cells
Materials
Plasmid pDS-YFP-HA-Orai1 encoding a dually tagged Orai1 with YFP at the N-terminus and an HA tag inserted in the second extracellular loop (see Hodeify et al., 2015)
Bicarbonate-buffered Ham’s F-12 medium (Invitrogen, catalog number: 11765054)
Cy3- and Cy5-conjugated anti-mouse secondary antibodies (Invitrogen, catalog numbers: A10521 and A10524)
Purified monoclonal anti-HA.11 Epitope Tag antibody (Covance, catalog number: MMS-101P)
Poly-D-lysine coated glass-bottomed plates (MatTek Corporation, catalog number: P35GC-1.5-14-C)
Lipofectamine 2000 (Invitrogen, catalog number: 11668019)
GibcoTM Opti-MEMTM I reduced serum medium, 500 mL (Thermo Fisher Scientific, catalog number: 31985047)
Dulbecco’s phosphate buffered saline (PBS) (Sigma, catalog number: D8537-500ML)
Penicillin-streptomycin (GIBCO REF, catalog number: 15140122)
Paraformaldehyde (PFA) (Sigma-Aldrich, catalog number: 16005-1KG-R)
Triton X-100 (Sigma, catalog number: T8787-50ML)
10% heat-inactivated FBS (Sigma, catalog number: F4135-500ML)
FalconTM 15 mL conical centrifuge tubes, polypropylene
FalconTM conical centrifuge tubes, 15 mL (Corning, catalog number: 352095)
Microcentrifuge tubes, 1.5 mL (Sigma-Aldrich, catalog number: HS4323)
250 mL glass Erlenmeyer flask (VWR, catalog number: 89000-362-250ML)
Solutions
Paraformaldehyde, 4% in PBS (see Recipes)
PBS containing 0.1% (w/v) Triton X-100 (see Recipes)
Recipes
Paraformaldehyde, 4% in PBS
Add 80 mL of 1× PBS into a 250 mL glass Erlenmeyer flask with a clean magnetic stir bar.
Heat the solution while stirring to approximately 60 and add 4 g of PFA powder (powder will not immediately dissolve).
Slowly raise the pH by adding 1 N NaOH dropwise until the solution clears.
Remove from heat and allow to cool to room temperature (RT).
Adjust the volume to 100 mL with 1× PBS.
Filter through aqueous filter paper to remove undissolved particles.
PBS containing 0.1% (w/v) Triton X-100
Add 5 μL of Triton X-100 to 50 mL of 1× PBS.
Equipment
Leica SP inverted confocal imaging microscope (Leica; Lasertechnik; S/N: 5100001464)
Magnetic stirrer with hotplate (IKA magnetic stirrer RCT basic, Ident. No.: 0025006052)
Cell culture incubator model: Sanyo MIR-262 Lab Incubator 153 L Capacity w/ Two Shelves S/N: 08020051 (Sanyo)
Software
ImageJ software (http://imagej.net/mbf/)
Adobe Photoshop CS3
Origin (v 2019b)
Procedure
Endocytosis assay protocol
To measure Orai1 endocytosis rate, cells are incubated with excess anti-HA antibody (Ab) to label the entire Orai1 pool at the PM (Figure 1). As Orai1 recycles at the plasma membrane, HA-associated Orai1 will be internalized by endocytosis, thus labeling the intracellular pool. Over time, one can then label the PM Orai1 pool by incubating with a Cy5 secondary Ab. This is followed by permeabilizing the cells and incubating with Cy3 secondary to label intracellular Orai1. The time course of the ratio of intracellular (Cy3) over surface (Cy5) Orai1 is fitted to quantify the endocytosis rate.
Figure 1. Outline of the endocytosis assay. CHO cells expressing YFP-HA-Orai1 are incubated with monoclonal anti-HA antibody for various periods of time. Cells are fixed and stained with Cy5-conjugated anti-mouse IgG, then washed and fixed again, permeabilized, and stained with Cy3-conjugated anti-mouse IgG. Cy5 and Cy3 intensities are then measured using quantitative fluorescence microscopy.
Cell preparation and transfection (Figure 2)
Figure 2. Flowchart depicting cell preparation and transfection with YFP-HA-Orai1 DNA
Cells are grown at 37 °C and 5% CO2 in bicarbonate-buffered Ham’s F-12 medium, supplemented with 10% heat-inactivated FBS and 5% (v/v) of a stock solution containing penicillin and streptomycin. For transient transfections, seed 300,000 cells on poly-D-lysine coated glass-bottomed plates for 18–20 h to 50%–60% confluency to initiate the experiment on the following day.
For transfection of each glass-bottom plate, prepare the transfection mixture as follows:
In two separate Eppendorf tubes, add 50 μL of Opti-MEMTM I reduced serum medium.
Add 1.5 μg of pDS-YFP-HA-Orai1 plasmid DNA to one tube and 2 μL of Lipofectamine 2000 to the other tube and vortex gently.
Incubate for 5 min at RT.
Combine the two mixtures in one tube, mix gently, and incubate for 20 min at RT.
While waiting, remove media from plates and wash with 500 μL of Opti-MEMTM.
After 20 min of incubation, add 300 μL of Opti-MEMTM to each mixture and add to the cells. Incubate for 18–20 h at 37 °C.
After overnight incubation, replace transfection mixture with normal medium and incubate cells for another 18–20 h.
Labeling and imaging (Figure 1)
Remove media from the transfected cells and incubate with 400 μL of monoclonal anti-HA.11 Epitope Tag antibody (1:100 dilution in 1× PBS) at 37 °C.
At various timepoints (2, 4, 6, 10, and 20 min), remove the PBS containing HA antibody and immediately fix cells for 10 min in 500 μL of 4% PFA at RT.
Then, remove PFA and wash cells twice with 1 mL of PBS.
Note: If a break point is needed, a few drops of PBS can be added to the cells, keeping those at RT for staining.
Stain cells with Cy5-conjugated anti-mouse secondary antibody (1:400 dilution in 1× PBS with 5% FBS) for 30 min at 37 °C.
Remove secondary antibody, wash cells with 1 mL of PBS, re-fix with 500 μL of 4% PFA for 10 min at RT, and then wash with 1 mL of PBS.
Permeabilize cells using 1 mL of 1× PBS with 0.1 % Triton for 10 min at RT.
Wash cells with 1 mL of PBS and then stain with Cy3-conjugated anti-mouse secondary antibody (1:400 dilution) for 30 min at 37 °C, before washing twice with 1 mL of PBS.
Notes:
During and after labeling with secondary antibodies, cover the plates in aluminum foil to minimize light exposure.
If a break point is needed, add a few drops of PBS to the cells and cover them with aluminum foil to reduce light exposure.
The fluorescence intensities of Cy3 and Cy5 are measured using quantitative fluorescence microscopy. The imaging parameters were as follows: for YFP, λex = 514, λem = 516–545; for Cy3, λex = 561, λem = 566–622; and for Cy5, λex = 633, λem = 648–703. The Cy5 signals show surface Orai1, whereas the Cy3 signal measures accumulation over time of intracellular Orai1 pool bound to the anti-HA antibody.
The (Cy3/YFP)Internal/(Cy5/YFP)Surface ratio is plotted over time and fitted with a linear regression function.
Imaging
The fluorescence intensities of the Cy3 and Cy5 were measured using quantitative fluorescence microscopy using Leica SP inverted confocal imaging microscope equipped with a 63× 1.4 NA oil lens. At early time points in the assay, Cy5 labeling of surface Orai1 was apparent, and intracellular Orai1 gradually became Cy3 positive (Figure 3).
Figure 3. Representative images of CHO cells expressing YFP-HA-Orai1 and fed with anti-HA antibody. Cells showing YFP distribution and stained for surface and internal Orai1 using an anti-HA antibody followed by a Cy5- and Cy3-conjugated secondary antibodies. Surface staining (Cy5 signal) is indicated by arrows and internal staining (Cy3) is represented by arrowheads. Scale bar, 5 μm.
Data analysis
Fluorescence of YFP, Cy5, and Cy3 was analyzed using ImageJ software at the single-cell level. Non-transfected cells (no YFP expression) were used for background correction. YFP, Cy3, and Cy5 signals at single-cell level were determined using ImageJ measure intensity tool after individually selecting cells (oval selection tool). Intensity was measured for both the Cy3 and YFP channels to allow for ratio calculations. Similarly, the ratio of Cy5 over YFP fluorescence at the single-cell level was determined. Unlabeled cells that did not express YFP-HA-Orai1 were used for background correction. The HA-bound to surface YFP-HA-Orai1 is labeled with Cy5-conjugated anti-mouse IgG. Cy5 labeling is followed by staining with an anti-mouse IgG conjugated to Cy3. The initial staining with Cy5-anti-mouse IgG blocks all PM HA epitopes, thereby limiting Cy3 labeling to internal HA epitope on YFP-HA-Orai1. The (Cy3/YFP)Internal/(Cy5/YFP)Surface ratio was plotted over time and fitted with a linear regression function to yield a straight line, whose slope is proportional, but not equal, to the Orai1 endocytosis rate, because two different fluorophores are used (Figure 4). The endocytosis rate constant can be calculated from levels of surface and internal Orai1 and the exocytosis rate constant using the equation:
Orai1Surface × Kendo = Orai1Internal × Kex
These methods have been validated for characterization of Orai1 recycling kinetics with CHO cells and HEK293 cells in Machaca Lab (Hodeify et al., 2015). A similar method was used by McGraw lab to successfully characterize GLUT4 transporter endocytosis (Karylowski et al., 2004; Blot and McGraw, 2008).
Figure 4. Intracellular accumulation of anti-HA-bound Orai1 measured as internal (Cy3/YFP) over surface (Cy5/YFP) Orai1 vs. time. Data were fitted by a linear regression. The slope of the straight line is proportional to endocytosis rate.
Exocytosis assay protocol
To quantify Orai1 exocytosis rates, cells are incubated with excess anti-HA Ab for various times to allow for HA-labeled Orai1 molecules to be internalized, and intracellular Orai1 to reach the PM. Initially, anti-HA antibody will bind to YFP-HA-Orai1 at the PM and binding will gradually increase overtime, as more intracellular Orai1 is translocated to the surface. As new Orai1 molecules reach the PM through recycling, they will be labeled by the HA Ab until the entire recycling Orai1 pool is labeled. Quantifying the amount of cell-associated anti-HA over time allows the quantification of Orai1 exocytosis rate.
Cell preparation and transfection
Prepare and transfect cells with YFP-HA-Orai1 as described above in the endocytosis assay.
Labeling and imaging (Figure 5)
Figure 5. Outline of the exocytosis assay. CHO cells expressing YFP-HA-Orai1 were incubated with monoclonal anti-HA antibody for various periods of time. Cells were fixed, permeabilized, and stained with Cy5-conjugated anti-mouse IgG. YFP and Cy5 intensities were measured using quantitative fluorescence microscopy.
Remove media from the transfected cells and incubate with monoclonal anti-HA.11 antibody (1:100 dilution in 1× PBS) at 37 °C.
At various timepoints (2, 5, 15, 30, and 45 min), remove the 1× PBS containing HA antibody and immediately fix cells for 10 min in 4% PFA at RT.
Remove PFA, wash cells with 1× PBS, and then permeabilize using 0.1% Triton for 10 min at RT.
Wash cells with 1× PBS and then stain with Cy5-conjugated anti-mouse secondary antibody (1:400 dilution) for 30 min at 37 °C.
The fluorescence intensities of YFP and Cy5 are measured using quantitative fluorescence microscopy.
Plotting the Cy5/YFP ratio over time and fitting with a mono-exponential growth curve
Data analysis
Fluorescence of YFP and Cy5 was analyzed using ImageJ software at the single-cell level. To correct for background, non-transfected cells (no YFP expression) were used for background correction. The ratio of Cy5/YFP fluorescence was calculated. The Cy5/YFP ratio was plotted over time and fitted with a mono-exponential growth curve (Figure 6). The ratio increases over time until it reaches a plateau, when the total pool of YFP-HA-Orai1 had sampled the PM and became attached to the anti-HA antibody. The exocytosis rate constant can be determined using the equation:
where A is the plateau of maximal uptake, B is the amplitude of the signal, t is the time, and Kex is the exocytosis rate constant. (Cy5/YFP)t is the fraction of YFP-HA-Orai1 bound by anti-HA monoclonal antibody.
Figure 6. Representative images from the exocytosis assay. A. Cells fed with anti-HA antibodies for various periods of time were fixed, permeabilized, and stained with Cy5 anti-mouse IgG. Scale bar, 5 μm. B. Cy5/YFP ratio was plotted and fitted with mono-exponential growth curve.
Surface/total ratio assay protocol
Cell preparation and transfection
Prepare and transfect cells with YFP-HA-Orai1 as described above in the endocytosis and exocytosis assay.
Labeling and imaging
Remove media from the transfected cells, wash once with 1× PBS, and fix immediately with 4% PFA for 10 min at RT.
In parallel, detect internal Orai1 in similar cells fixed with 4% PFA for 10 min at RT, washed with 1× PBS, and then permeabilized using 0.1 % Triton for 10 min at RT.
Wash non-permeabilized and permeabilized cells with 1× PBS and then incubate with saturating concentrations (1:300) of monoclonal anti-HA antibodies for 45 min at 37 °C. Wash cells with 1× PBS and then stain with Cy5-conjugated anti-mouse secondary antibody (1:400 dilution) for 30 min at 37 °C.
The fluorescence intensities of YFP and Cy5 are measured using quantitative fluorescence microscopy.
Surface Orai1 is determined by dividing the Cy5/YFP for non-permeabilized cells by the Cy5/YFP value for permeabilized cells.
Calculating Kexo and Kendo
As an example for calculating Kexo and Kendo, we used the data shown in Figures 4 and 6.
The exocytosis data in Figure 6 showing a time course of Cy5/YFP is fit to a mono-exponential curve using this formula:
The maximum uptake plateau A = 0.7969,
The initial value Y0 = 0.07393,
The amplitude of the signal, B = plateau – Y0 = 0.7895.
Solving the equation with these values for Kex gives 0.048 ± 0.006 min-1
Using the surface to total ratio protocol described above, we have also calculated that 45% of total Orai1 is at the surface at steady state. This provides a relative Orai1 surface of 0.45 and internal of 0.55. Using the calculated Kex from the exocytosis assay and this value, we can calculate Kendo using the formula:
Orai1Surface × Kendo = Orai1Internal × Kex
So, Kendo = 0.05866/min.
General notes and troubleshooting
This protocol can be adapted to be used in any transfectable cell line. One limitation is due to the non-homogenous expression of the protein of interest following transient transfection, which may lead to overexpression and saturation of the endogenous trafficking machinery. To assess this possibility, one can assess the surface/total ratio of the protein of interest as a measure of total expression using the fluorescent tag (YFP, in this case). If the trafficking machinery is not saturated, surface/total ratio should be stable over a wide range of expression. Alternatively, one can generate stable cell lines that express low levels of the tagged protein (see Hodeify et al., 2015).
Precautions should be taken when preparing PFA, as it is irritating to eyes, respiratory system, and skin. Wear appropriate personal protective equipment and prepare solution in a certified chemical fume hood.
Acknowledgments
This work was supported by NPRP 9-767-3-208 grant from the Qatar National Research Fund to K.M and by the Biomedical Research Program at Weill Cornell Medical College in Qatar, a program funded by Qatar Foundation.
This protocol was derived from the original work of Hodeify et al. (2018).
Competing interests
The authors declare no conflict of interests.
References
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4,801 | https://bio-protocol.org/en/bpdetail?id=4801&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Expression Stability Analysis of Candidate References for Normalization of RT-qPCR Data Using RefSeeker R package
PP Patrick H.D. Petersen *
JL Joanna Lopacinska-Jørgensen *
CH Claus K. Høgdall
EH Estrid Vilma Høgdall
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4801 Views: 488
Reviewed by: AYŞE NUR PEKTAŞUte Angelika HoffmannHélène Léger
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Original Research Article:
The authors used this protocol in PLOS ONE May 2023
Abstract
When performing expression analysis either for coding RNA (e.g., mRNA) or non-coding RNA (e.g., miRNA), reverse transcription quantitative real-time polymerase chain reaction (RT-qPCR) is a widely used method. To normalize these data, one or more stable endogenous references must be identified. RefFinder is an online web-based tool using four almost universally used algorithms for assessing candidate endogenous references—delta-Ct, BestKeeper, geNorm, and Normfinder. However, the online interface is presently cumbersome and time consuming. We developed an R package, RefSeeker, which performs easy and straightforward RefFinder analysis by enabling raw data import and calculation of stability from each of the algorithms and provides data output tools to create graphs and tables. This protocol uses RefSeeker R package for fast and simple RefFinder stability analysis.
Key features
• Perform stability analysis using five algorithms: Normfinder, geNorm, delta-Ct, BestKeeper, and RefFinder.
• Identification of endogenous references for normalization of RT-qPCR data.
• Create publication-ready graphs and tables output.
• Step-by-step guide dialog window for novice R users.
Graphical overview
Simple workflow diagram. Two main workflow paths are presented. A) Using the RefSeeker wizard allows non-R programmers to easily load data and choose between selected output formats. B) Command line interface provides more options to control input and output formats and to automate analysis.
Keywords: RT-qPCR Normalization Coding and non-coding RNA expression RefFinder Expression stability R package
Background
Whether coding or non-coding, gene expression research represents a large field of investigation, including molecular biomarker research, drug research, cancer diagnostics, pathway research, RNA interference studies, stem cell research, and much more. In many of these fields, reverse transcription quantitative real-time polymerase chain reaction (RT-qPCR) is used to validate results and investigate changes in expression of a variety of RNA types. The Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guideline requirements have been widely adopted by the scientific community (Bustin et al., 2009). These guidelines assert that the use of three or more stably expressed endogenous references should be used for normalization of target RNAs (e.g., genes/mRNA or miRNAs). Additionally, these references should preferably be of the same type of RNA as the targets.
When performing expression analysis, it is often required to analyze the stabilities of reference genes used to normalize the data from targets of interest (Bustin et al., 2009). The selection of a sufficient number of adequately stable references, typically three or four, is a crucial step since the choice may significantly influence the results and could lead to wrong conclusions (Faraldi et al., 2019). Expression data is often obtained as a set of quantification cycle (Cq), crossing point (Cp), cycle threshold (Ct), or take-off point (TOP) values (Bustin et al., 2009). These expression values are typically obtained by performing the RNA quantification in technical triplicates or quadruplicates, averaging these data results in one value per RNA target (e.g., gene or miRNA) for each sample.
Four different algorithms are commonly used for identifying stable RNAs: (1) Normfinder calculates intra- and inter-group variations (Andersen et al., 2004); (2) geNorm uses an average pairwise standard deviation with all other candidates as the stability measure M (Vandesompele et al., 2002); (3) BestKeeper calculates a range of statistics but bases the individual stability on the mean absolute deviation of the raw Cp values (Pfaffl et al., 2004); and (4) the delta-Ct (ΔCt) method compares each candidate average standard deviation of ΔCt values for each combination of candidates (Silver et al., 2006). Further details on the different algorithms are beyond the scope of this protocol, and further information on their strengths and weaknesses can be found elsewhere (De Spiegelaere et al., 2015; Sundaram et al., 2019).
To deal with differences in results from these algorithms, Xie et. al. created RefFinder, which combines the rankings of the four algorithms and gives a geometric mean of these ranks (Xie et al., 2012 and 2022). RefFinder is available as an online tool, allowing researchers and others to perform the analysis by copying their expression data into a textbox and pressing the analyze button; results are then presented on the webpage.
Given that RefFinder is an online tool, data needs to be copied and pasted into a web-based interface. After the analysis, the results need to be copied and pasted back into a statistical or spreadsheet software of choice for further processing, table generation, and/or graphical depiction. This involves extensive manual work, especially in cases when multiple datasets are used simultaneously or when analyses need to be redone. Moreover, the process can be error prone, considering that copying and pasting manually to and from many different sources and destinations can be disorienting. Therefore, we aimed to develop a straightforward method to perform RefFinder analysis on preprocessed RT-qPCR data, providing easy generation of tables, datasheets, and graphical output: the RefSeeker package.
Software description
RefSeeker is a package developed in R designed to be compatible across different operating systems. R is a widely used, free, and open-source statistical environment, thus providing a great basis for expression data analysis (R Core Team, 2022). R provides great tools for working with data in tabular format as well as for plotting and graphing. The RefSeeker package utilizes widely available tools either available through base R or through The Comprehensive R Archive Network (CRAN).
As for RefFinder, to use the RefSeeker package, expression data need to be prepared in a tabular format. However, data can be prepared either as a data object prepared in R or as one of the supported file types (.xls, .xlsx, .ods, .csv, .tsv, or .txt). Each column represents a named target (gene, miRNA, or other) and each row represents a sample. If the data file is created using R, an index column might be added to the .csv file by default. This should be avoided by setting row.names = FALSE. In case of spreadsheets (Excel or .ods) where more than one dataset is included, each sheet in the spreadsheet file can contain a dataset. Naming the sheets will make it easier to identify the data later, since the name is carried over. In case of txt-based files, e.g., .csv, .tsv, or plain .txt tables, each dataset must be in a separate file in the same folder.
The package functions can be divided into four categories:
1. Data import functions, which import data from different sources (.csv, .tsv, .txt, .xls, .xlsx, and .ods) and arrange it for further processing. A wrapper function, rs_loaddata(), identifies the file extensions and calls the proper import function.
2. Data processing functions. RefSeeker uses four main functions to perform the RefFinder analysis: rs_normfinder(), rs_genorm(), rs_bestkeeper(), and rs_deltact(). These functions are all called by the rs_reffinder() function to determine stability rankings. The comprehensive rank is then calculated as the geometric mean of these stability rankings.
3. Data export functions for further analysis, visualization, and publication of results. The function rs_graph() handles printing and optionally exporting of graphs as .png, .tiff, .jpeg, or .svg file formats. Likewise, the function rs_exporttable() handles export of data tables, either as spreadsheets (.ods or .xlsx), txt-based formats (.csv, .tsv, .txt), or formatted tables in docx format.
4. Interactive implementation of the above functionality through the rs_wizard(). This function provides a graphical user interface dialog window for selecting data and output table and graphical formats.
See Figure 1 for an overview of the main functions and their association to the workflow.
Figure 1. Simple data analysis workflow diagram. Data can be loaded from outside sources via the rs_load() or rs_wizard() functions. RefFinder analysis can be performed on the data using rs_reffinder(), and the analyzed data can be visualized and exported via the rs_exporttable() and rs_graph() functions. Examples of output can be seen in Figure 3, Figure 4, Figure 5, and Table 1.
Equipment
Computer with Windows, MacOS, or Linux-based operating system compatible with R (≥ 4.1.0)
Software and datasets
R software environment (≥ 4.1.0) (https://www.r-project.org/)
RStudio integrated development environment (≥ 1.4.0) (optional, https://rstudio.com/)
Datasets can be prepared in several ways, and processing of raw expression data is outside the scope of this protocol. However, in general, data should be cleaned, quality checked, and adjusted for interplate variability (Petersen et al., 2022).
Data can be prepared in either one of the supported file types (.xlsx, .ods, .csv, .tsv, or .txt) or as a data frame in R.
Sample data can be downloaded from: https://github.com/Hannibal83dk/RefSeeker/blob/main/SampleData/RefSeekerSampleData.xlsx (see Figure 2). These data have been previously described, and details about experiment design, data acquisition, and processing can be found in Petersen et al. (2022), from where the data have been obtained.
• No matter the input source, the following requirements are the same:
• Each column must be representing a gene/target and each row an individual sample*.
• Each column must be named.
• Row names must be excluded.
• No missing data is allowed**.
*Although the RefSeeker package can handle spaces and dashes in column names, some downstream R processes might not be able to. Best practices therefore recommend avoiding these characters in column names.
**Missing data can be handled in several ways. If samples need to be preserved, targets can be removed; if it is more desirable to keep targets, samples can be removed. If both are important, a percentage threshold for allowed missing data can be chosen. This threshold is individually selected; however, it should be as low as possible. A specific recommendation cannot be provided here; however, a threshold of 20% missing data has been used before and seems to be an approximate upper limit. Following target exclusion, remaining missing datapoints can be imputed using different tools [e.g., MissForest (missForrest package), k-Nearest Neighbor (VIM package), Multiple Imputation by Chained Equations (mice package), or max + 1 (manually implemented)].
Figure 2. Example of expression data in an Excel file used for RefSeeker analysis. These may be averages of triplicates or quadruplicates and should be adjusted for e.g., possible interplate variation. Targets are given in the first row. Each of the following rows represents raw Cp values (adjusted for interplate variance) obtained from each sample. The file contains three spreadsheets: fresh frozen, formalin fixed and paraffin embedded (FFPE), and Plasma, each containing different datasets.
Procedure
Installing R and RStudio
The installation of R and RStudio are out of the scope of this protocol; however, more information on how to install these can be found on their respective websites:
R base: https://cran.r-project.org/
RStudio: https://posit.co/download/rstudio-desktop/
Installing dependencies
The RefSeeker has a few dependencies that need to be installed first by typing:
install.packages(c('ctrlGene', 'ggplot2', 'reshape2', 'readxl', 'openxlsx', 'data.table', 'readODS', 'flextable', 'officer'))
Installation of RefSeeker package
After installation of dependencies, the package can be installed in two ways:
Download Package Archive from GitHub:
Download the latest version of RefSeeker_latest.tar.gz file to your computer:
The latest version can be found at:
https://github.com/Hannibal83dk/RefSeeker/releases/latest/download/RefSeeker_latest.tar.gz
Open R or RStudio.
In the R Console, type:
install.packages("<PATH/TO/RefSeeker_latest.tar.gz>", repos = NULL, type = "source")
Note: Please note that the entire part of <PATH/TO/RefSeeker_latest.tar.gz> needs to be changed to the specific location of the downloaded file on your computer.
Alternatively, if RStudio is being used: from the menu bar, open the dropdown menu Tools > Install Packages > Select Package from Archive File in the Install from drop-down menu. Browse for the downloaded Package archive > press Install.
Use devtools to download from GitHub:
Make sure the devtools package is installed in your R environment. In the R Console, devtools can be installed from CRAN using:
install.packages("devtools")
In the R console type:
devtools::install_github("Hannibal83dk/RefSeeker", build_vignettes = TRUE)
RefFinder analysis
To illustrate the usability and ease of use of the package, three expression datasets with Cp data of six miRNAs from 20 patients in three sample types [Fresh frozen, formalin fixed and paraffin embedded (FFPE), and blood plasma] will be used (Petersen et al., 2022). The data are stored as individual sheets in an .xlsx file (Figure 2).
The aim is to identify the most stable miRNAs for normalization for each dataset. Firstly, the package needs to be loaded into an R environment. Two options are available for analysis.
Quick all-in-one analysis wizard:
Load the RefSeeker package:
In the R console first type:
library(RefSeeker)
Running the RefSeeker wizard will open a graphical interface dialog window, providing quick selection of input data and output formats.
Now, run the following in the console:
rs_wizard()
This will result in a dialog window popping up (see Figure 3).
Figure 3. The rs_wizard() dialog allowing interactive quick analysis. The dialog consists of nine sections: i) selection of input file(s), ii) selection of folder for output files, iii) optional prefix for output file names, iv) selection of bar graph format, v) selection of order of the target axis, vi) selection of the orientation of the bar graph, vii) selection of the graph output file format, viii) selection of the table format, and ix) option for proceeding or quitting the analysis.
Pressing the Select input files button (Figure 3-i) will open a file manager. From here, navigate to the .xlsx file and select it.
Press the Change output folder button (Figure 3-ii) and select an output directory from the file manager.
A file name prefix can be selected to identify the output files (Figure 3-iii). From the radio buttons, the desired output can be modified.
First, select the desired type of graph (Figure 3-iv): Individual will make a graph for each dataset and Multi will create a faceted graph combining all datasets in one graph (see Figure 4).
The ordering of the x-axis can also be changed (Figure 3-v). This defaults to Comprehensive Rank, meaning that all the target arrangements on the x-axis will be ordered from most to least stable, based on the ranking provided by the comprehensive rank.
The desired direction of the bar plot can be selected (Figure 3-vi).
Select desired file format to output the graph (Figure 3-vii).
Lastly, select the desired table output format (Figure 3-viii).
Press Ok and collect your outputs in the selected output folder (Figure 3-ix).
From the default selections, a .png file and an .xlsx file will be created in the output folder selected (Figure 4 and Figure 5). In step D1i (Figure 3-viii), a docx type table can be selected instead of the .xlsx file. An example of a docx type table format can be seen in Table 1. This is a good choice for presenting stabilities in a publication.
From the generated output (Figure 4 and Figure 5), targets with lower stability values are considered more stable. It is seen that the most stable endogenous miRNAs are hsa-miR-191-5p and hsa-miR-23a-3p for plasma, hsa-miR-23a-3p and hsa-miR-30c-3p for fresh frozen tissue, and hsa-miR-30c-3p ahashsa-miR-191-5p for FFPE tissue. Note that, since the different algorithms evaluate stability differently, results may vary between these (De Spiegelaere et al., 2015). Since BestKeeper only evaluates standard deviations of each sample, it is common to observe high differences between BestKeeper and the other algorithms that are more interdependent, especially GeNorm and delta-Ct, which evaluate highly correlated targets as more stable. Sundaram et al. (2019) suggests an integrated approach to this problem by removing targets with high overall variance before performing the analysis again.
Figure 4. Example of graph output created using RefSeeker. A multi graph with data from three different datasets using horizontal layout and targets sorted by the Comprehensive Rank.
Figure 5. Example of Excel table output. Stability values from each algorithm are provided in the first sheet and targets are ordered by the Comprehensive Rank. The second sheet is similar to the first but contains rankings instead of stability values.
Table 1. Example of docx-Combi type table output format
Plasma
delta-Ct
BestKeeper
Normfinder
geNorm
Comprehensive Rank
Target
Avg. STDEV.
Rank
MAD
Rank
Stability
Rank
Avg.M
Rank
Geom. mean value
Rank
hsa-miR-191-5p
0.756
1
1.355
2
0.075
1
0.332
1
1.189
1
hsa-miR-23a-3p
0.765
2
1.526
3
0.274
3
0.332
1
2.060
2
hsa-miR-30c-5p
0.826
3
1.541
4
0.217
2
0.488
4
3.130
3
hsa-miR-451a
1.735
5
1.172
1
1.704
5
0.986
5
3.344
4
hsa-miR-103a-3p
0.850
4
1.639
5
0.603
4
0.408
3
3.936
5
Command line analysis:
To gain more control over the outputs and run additional analyses, the R command line can be utilized. It is recommended to create an R-script from where commands can be run. Additionally, an R-markdown document using the details and sessioninfo libraries is a good way to document session info and version of used packages and report on findings in a repeatable manner.
First, load the RefSeeker package:
library(RefSeeker)
Data import
Load in the data into an R variable:
inputData <- rs_loaddata()
From the file selection dialog, find and select the data file(s).
Alternatively, the file path(s) can be given as an argument to the function; this is recommended to increase reproducibility of code.
inputData <- rs_loaddata(c(‘path/to/file1’, ‘path/to/file2’))
RefFinder analysis
Perform the RefFinder analysis:
results <- rs_reffinder(inputData)
The results can be checked by typing:
results
See File S1 for an example of the output given. The output is returned as a list of lists containing the results of each dataset. Results from each dataset are given as a list of two tables: one for stability values and one for stability rankings of all targets.
Results for each individual dataset can be accessed by typing:
results$Fresh_Frozen
This will return the two tables for the Fresh Frozen dataset.
To access individual tables, type:
results$Fresh_Frozen$rankTable
This will return the rank table for the Fresh Frozen dataset.
From the results, a set of references can be selected. For the plasma set, hsa-miR-191-5p and hsa-miR-23a-3p seem to be most stable; hsa-miR-23a-3p and hsa-miR-30a-3p seem to be most stable in Fresh Frozen tissue; and hsa-miR-30a-3p and hsa-miR-191-5p are most stable in FFPE.
Graph export
To produce a preview of the bar graph illustrating the results, use:
rs_graph(results)
Add colors to the graph by creating a data frame matching targets and colors.
colors <- data.frame(targets = names(inputData[[1]]), color = c("#E69F00", "#0072B2", "#009E73", "#CC79A7", "#D55E00"))
Here, the target names for the first column are collected in the first dataset of the inputData dataset list. A custom color scheme is then created for the second column. It is recommended to use colors that account for different kinds of color visions. Colors provided here were selected for that purpose; however, a wider selection can be obtained through the package colorblindr.
Running the function rs_graph again, inputting the color data frame will create a preview of the colored graph in the plot pane (see Figure 6):
rs_graph(results, colors = colors)
The function will output a width and height. These will be used for the exported image if not changed later in step D2g.
> width set to 2400
> height set to 960
Other arguments can be used to adjust the graph. It is encouraged to type help(rs_graph) to get an overview. Here, the main arguments that can be adjusted before exporting to a file are shown:
rs_graph(results, forceSingle = FALSE, ordering = "Comprehensive Rank", orientation = "horizontal", colors = colors)
Figure 6. Example output of the colored .png file created with the rs_graph function
Once the graph has the desired appearance, it can be exported to a number of different image file types. Again, type help(rs_graph) to see a comprehensive list of argument options. Here, we will create the default graph with the colors that were selected previously. To create the image file, a filename should be passed to the function:
rs_graph(results, filename= "Ovarian Cancer", colors = colors)
Also here, the function will output width and height as well as a confirmation that a file was created and a file path:
> width set to 2400
> height set to 960
> A png file was created at: <PATH\TO\FILE\>
Inspect the image file that was created (Figure 6). If needed, adjust the size using the width and height given in the previous output as a reference. Resolution can be set using the res-parameter; smaller numbers will make lines and text smaller and finer:
rs_graph(results, filename = "Ovarian Cancer", colors = colors, width = 2000, res = 200)
Table export
Creating a table output can be useful in many ways:
• To store results in universal lightweight file formats like .csv or .tsv.
• To share via Excel or OpenDocument Spreadsheet.
• To present findings in a word file or in a publication. Here, we will create an Excel file (see Figure 5) as well as a docx-combi table (see Table 1) for each dataset.
Creating an Excel file: the default table type is .xlsx, so this does not need to be specified:
rs_exporttable(results, filename = "Ovarian Cancer")
In this case, three .xlsx-files are created, one for each dataset.
Creating a docx-combi type table can be done by setting the tabletype parameter:
rs_exporttable(results, filename = "Ovarian Cancer", tabletype = "docx-combi")
Running individual algorithms
Stability values from each algorithm can be obtained individually. In this case, datasets must be passed individually:
rs_genorm(inputData$Plasma)
rs_normfinder(inputData$Plasma)
rs_deltact(inputData$Plasma)
rs_bestkeeper(inputData$Plasma)
Normfinder ungrouped and grouped stability analysis and paired candidate stability
Since it is recommended to use more than one reference gene, it may be of interest to identify pairs of targets, which are most stable in combination. If a set of reference targets have been chosen, these can be crudely validated through the Normfinder algorithm, which is able to provide further stability statistics and assessment of stability between groups and of pairs of genes. This approach is beneficial for identifying stable pairs and to compare selected references.
In the following example, we will use the Normfinder algorithm as a quick check of the selected references selected previously for fresh frozen and FFPE tissues.
Perform individual Normfinder analysis on Fresh frozen and FFPE datasets.
rs_normfinderFull(inputData$Fresh_Frozen, Groups = FALSE)
Giving the output:
$Ordered
GroupSD
hsa-miR-23a-3p 0.512
hsa-miR-30c-5p 0.607
hsa-miR-191-5p 0.668
hsa-miR-103a-3p 0.830
hsa-miR-451a 1.075
$PairOfGenes
Gene1 Gene2 GroupSD
1 hsa-miR-30c-5p hsa-miR-23a-3p 0.424
This output indicates that, for the fresh frozen samples, a combination of hsa-miR-30c-5p and hsa-miR-23a-3p are the most stable pair. This is in agreement with the previous results that showed that hsa-miR-23a-3p and hsa-miR-30c-3p were the two most stable targets.
rs_normfinderFull(inputData$FFPE, Groups = FALSE)
Giving the output:
$Ordered
GroupSD
hsa-miR-30c-5p 0.344
hsa-miR-23a-3p 0.493
hsa-miR-191-5p 0.513
hsa-miR-103a-3p 1.179
hsa-miR-451a 1.857
This output indicates that, for the FFPE samples, hsa-miR-30c-5p and hsa-miR-23a-3p are the most stable. This is in agreement with the previous Normfinder results.
Note: Since no PairOfGenes are shown, no candidates were assessed as stable enough in a first round of calculations for paired analysis. This threshold is by default set to 0.25; however, it can be set via the pStabLim argument.
If differential expression is to be evaluated between two sample types, for example before or after treatment, it may be of interest to find target references that in combination show high stability between the two sample types. Here, we will use the Normfinder grouped analysis to identify stable pairs of references candidates for comparing expression across fresh frozen and FFPE tissue.
Perform a full Normfinder grouped analysis on the Fresh frozen and FFPE datasets.
First, we need to create a grouped dataset.
freshFrozen <- inputData$Fresh_Frozen
freshFrozen$group <- 1
FFPE <- inputData$FFPE
FFPE$group <- 2
Grouped <- rbind(freshFrozen, FFPE)
Now, perform the analysis:
rs_normfinderFull(Grouped, Groups = TRUE)$ PairOfGenes
Note: Here, we only select the PairOfGenes table to be printed.
Giving the output:
Gene1 Gene2 Stability
1 hsa-miR-30c-5p hsa-miR-103a-3p 0.128
2 hsa-miR-30c-5p hsa-miR-191-5p 0.086
3 hsa-miR-30c-5p hsa-miR-23a-3p 0.079
4 hsa-miR-103a-3p hsa-miR-191-5p 0.133
5 hsa-miR-103a-3p hsa-miR-23a-3p 0.127
6 hsa-miR-191-5p hsa-miR-23a-3p 0.088
According to Normfinder grouped analysis, hsa-miR-30c-5p and hsa-miR-191-5p are deemed the most stable pair across two groups.
Data analysis
Conclusions
In this protocol, we show how to perform stability analysis using widely used algorithms: geNorm, Normfinder, delta-Ct, BestKeeper, and RefFinder by the RefSeeker package for R. This protocol is easy to follow with its step-by-step guide and allows non-R programmers to perform stability analyses. It can be used to identify stable references in any kind of expression analysis and gives great options for data export of ready-to-publish graphs and tables.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article:
Lopacinska-Jørgensen et al. (2023). Strategies for data normalization and missing data imputation and consequences for potential diagnostic microRNA biomarkers in epithelial ovarian cancer cancer. PLoS One (Table 3, Table 5 and Table S3).
Acknowledgments
We thank Douglas Vinicius Oliveira for valuable discussions. Also, we thank Rasmus Adalbert Meldgaard for early testing of the package. We are grateful to the Danish CancerBiobank (Bio- and GenomeBank Denmark) and the Danish Gynecologic Cancer Database for making specimens and data available for use in the present study. This work was founded by: The Mermaid Foundation, URL: http://www.mermaidprojektet.dk/ (PHDP, JLJ, CKH, and EVH received the funding), Danish Cancer Research Foundation: URL:http://www.dansk-kraeftforsknings-fond.dk/ (EVH received the funding), and Herlev Hospital Research Council, URL: https://www.herlevhospital.dk/forskning/ (EVH received the funding).
Competing interests
The authors declare no competing interests.
References
Andersen, C. L., Jensen, J. L. and Ørntoft, T. F. (2004). Normalization of Real-Time Quantitative Reverse Transcription-PCR Data: A Model-Based Variance Estimation Approach to Identify Genes Suited for Normalization, Applied to Bladder and Colon Cancer Data Sets. Cancer Res. 64(15): 5245–5250.
Bustin, S. A., Benes, V., Garson, J. A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M. W., Shipley, G. L., et al. (2009). The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments. Clin. Chem. 55(4): 611–622.
De Spiegelaere, W., Dern-Wieloch, J., Weigel, R., Schumacher, V., Schorle, H., Nettersheim, D., Bergmann, M., Brehm, R., Kliesch, S., Vandekerckhove, L., et al. (2015). Reference Gene Validation for RT-qPCR, a Note on Different Available Software Packages. PLoS One 10(3): e0122515.
Faraldi, M., Gomarasca, M., Sansoni, V., Perego, S., Banfi, G. and Lombardi, G. (2019). Normalization strategies differently affect circulating miRNA profile associated with the training status. Sci. Rep. 9(1): e1038/s41598-019-38505-x.
Petersen, P. H. D., Lopacinska-Jørgensen, J., Oliveira, D. V. N. P., Høgdall, C. K. and Høgdall, E. V. (2022). miRNA Expression in Ovarian Cancer in Fresh Frozen, Formalin-fixed Paraffin-embedded and Plasma Samples. In Vivo 36(4): 1591–1602.
Pfaffl, M. W., Tichopad, A., Prgomet, C. and Neuvians, T. P. (2004). Determination of stable housekeeping genes, differentially regulated target genes and sample integrity: BestKeeper – Excel-based tool using pair-wise correlations. Biotechnol. Lett 26(6): 509–515.
R Core Team. (2022). R: A language and environment for statistical computing 2022.
Silver, N., Best, S., Jiang, J. and Thein, S. L. (2006). Selection of housekeeping genes for gene expression studies in human reticulocytes using real-time PCR. BMC Mol. Biol. 7(1): e1186/1471-2199-7-33.
Sundaram, V. K., Sampathkumar, N. K., Massaad, C. and Grenier, J. (2019). Optimal use of statistical methods to validate reference gene stability in longitudinal studies. PLoS One 14(7): e0219440.
Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A., and Speleman, F. (2002) Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol. 3: research0034.1.
Xie, F., Xiao, P., Chen, D., Xu, L. and Zhang, B. (2022). RefFinder. Accessed February 21, 2022.
Xie, F., Xiao, P., Chen, D., Xu, L. and Zhang, B. (2012). miRDeepFinder: a miRNA analysis tool for deep sequencing of plant small RNAs. Plant Mol. Biol. 80(1): 75–84.
Supplementary information
The following supporting information can be downloaded here:
File S1. Example of a rs_reffinder result output.
Article Information
Copyright
© 2023 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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4,802 | https://bio-protocol.org/en/bpdetail?id=4802&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
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Peer-reviewed
Isolation of Embryonic Cardiomyocytes and Cell Proliferation Assay Using Genetically Engineered Reporter Mouse Model
MB Maren Beall
DL Deqiang Li
JJ Jihyun Jang
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4802 Views: 768
Reviewed by: Pilar Villacampa AlcubierreKyoung-Han KimMarina Sánchez PetidierChao Wang
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Original Research Article:
The authors used this protocol in Circulation Research Jul 2022
Abstract
Congenital heart disease (CHD) is often associated with myogenic defects. During heart development, cardiomyocyte growth requires essential cues from extrinsic factors such as insulin-like growth factor 2 (IGF-2). To determine whether and how growth factors account for embryonic cardiomyocyte proliferation, isolation followed by culturing of embryonic cardiomyocytes can be utilized as a useful tool for heart developmental studies. Current protocols for isolating cardiomyocytes from the heart do not include a cardiomyocyte-specific reporter to distinguish cardiomyocytes from other cell types. To optimize visualization of cardiomyocyte proliferation, our protocol utilizes a Tnnt2-promoter-driven H2B-GFP knock-in mouse model (TNNT2H2B-GFP/+) for in vitro visualization of nuclear-tagged cardiomyocyte-specific fluorescence. A cardiomyocyte-specific genetic reporter paired with an effective proliferation assay improves the reproducibility of mechanistic studies by increasing the accuracy of cell identification, proliferated cell counting, and cardiomyocyte tracking.
Key features
• This protocol refines previous methods of cardiomyocyte isolation to specifically target embryonic cardiomyocytes.
• UsesH2B-GFP/+cardiomyocyte reporters as identified by Yan et al. (2016).
• Traces cell proliferation with Phospho-Histone 3 (p-H3) assay.
• Has applications in assessing the role of growth factors in cardiomyocyte proliferation.
Graphical overview
Keywords: Embryonic cardiomyocytes Cardiomyocyte isolation Heart development Cell proliferation Knock-in mice
Background
Myocardial expansion is one of the most critical cardiac developmental events. Failure of this process gives rise to many forms of congenital heart disease (CHD), which remains the most common cause of infant death (Zaidi and Brueckner, 2017). During heart development, cardiomyocytes intimately communicate with adjacent nonmyocytes, which promote cardiomyocyte proliferation through paracrine signals (Jang et al., 2022).
To study whether extrinsic cues contribute to embryonic cardiomyocyte proliferation, isolated cardiomyocytes from embryonic hearts can be used to explore the underlying mechanisms of myocardial growth. Here, a newly introduced genetically engineered reporter mouse model (Tnnt2H2B-GFP/+) (Yan et al., 2016) was paired with a proliferation assay. Since previous protocols have outlined the rudimentary trypsin digestion method for embryonic cardiomyocyte isolation (Rodgers et al., 2009), isolated cells were primarily cardiomyocytes but also included cardiac fibroblasts and cardiac endothelial cells. Thus, cardiomyocyte staining would be needed for further tracking of cardiomyocyte-specific proliferation within isolated cells. To monitor cardiomyocyte proliferation over time without the need of further staining, we used a cardiomyocyte specific reporter, cardiac Troponin T (Tnnt2), to definitively confirm cell type for accuracy in subsequent cell counting. With the inclusion of a proliferation assay, this protocol provides a reliable approach for in vitro assessments of growth factor effects on myocardial development.
This protocol consists of three key steps: embryonic cardiomyocyte isolation, cardiomyocyte culture, and immunostaining of cell proliferation markers. First, the hearts will be collected and digested with trypsin, and then further separated with dissociation buffer. For cardiomyocyte culture, cells will be pre-plated in culture media to remove other cell types, such as fibroblasts and endothelial cells, and will then be moved into a laminin-coated dish to support myocardial cells. To measure cell proliferation, serum-free media will be added to starve the cells and prevent residual growth factor influences; cells will then be treated with insulin-like growth factor 2 (IGF-2) to stimulate cardiomyocyte proliferation. Finally, immunocytochemistry staining with Phospho-Histone 3 (p-H3) will be applied for counting the proliferating cells as well as total GFP-positive cardiomyocytes.
Taken together, the proliferation assay with the genetic reporter mouse model will allow us to dissect the general mechanisms of the mitotic properties of cells, which can be applied to developmental research.
Materials and reagents
Biological materials
p-H3 (rabbit) (Ser10) antibody (Cell Signaling, catalog number: 9701S), store at -20 °C
Donkey anti-rabbit IgG (H+L), Alexa Fluor 555 (Life, catalog number: A31572)
100 ng/mL IGF-2 recombinant protein (R&D Systems, catalog number: 792-MG-050)
TNNT2H2B-GFP/+mice (from Dr. Chen-Leng Cai, Background Strain: C57BL/6J)
Reagents
1× PBS pH 7.4 (Gibco, catalog number: 10010-023)
1 M HEPES (Gibco, catalog number: 15630-060), store at 4 °C
0.5 M EDTA pH 8.0 (Gibco, UltraPure, catalog number: 15575), store at 4 °C
1:250 Trypsin (Gibco, catalog number: 27250-018)
HBSS solution (Gibco, catalog number: 14175-095)
Horse serum (Gibco, catalog number: 16050-114), store at -80 °C
FBS (Gibco, catalog number: 10437-028), store at -80 °C
100× antibiotic-antimycotic (Gibco, catalog number: 15240-062)
1× DMEM [-] Ca2+(Gibco, catalog number: 21068-028), store at 4 °C
Laminin (Corning, catalog number: 354232), store at -80 °C
1× DPBS (Ca2+/Mg2+) (Corning, catalog number: 21-030-CV)
1× Opti-MEM (Gibco, catalog number: 31985-070), store at 4 °C
IC fixation buffer (Invitrogen, catalog number: 00822249), store at 4 °C
Triton X-100 (Fisher Biotech, catalog number: BP151-500)
Donkey serum (Fisher Brand, catalog number: 03-395-464)
0.1 μg/mL DAPI (1:1,000 working concentration) (Sigma, catalog number: D9542)
1× DMEM [+] Ca2+(Gibco, catalog number: 11965-092), store at 4 °C
Solutions
Trypsin solution (see Recipes)
Dissociation buffer (see Recipes)
Rinse media (see Recipes)
Culture media (see Recipes)
Recipes
Trypsin solution (in 1× HBSS)
Reagent Final concentration Quantity (25 mL)
HEPES (1 M) 10 mM 250 μL
EDTA (0.5 M) 0.54 mM 27 μL
Trypsin 0.25% 0.0625 g
HBSS solution (1×) n/a 25 mL
Dissociation buffer (in 1× HBSS) at 4 °C
Reagent Final concentration Quantity (25 mL)
Horse serum 10% 2.5 mL
FBS 5% 1.25 mL
HEPES (1 M)
HBSS Solution (1×)
n/a
n/a
250 μL
21.2 mL
Rinse media (in Ca2+free DMEM)
Reagent Final concentration Quantity (50 mL)
Horse serum 10% 5 mL
FBS 5% 2.5 mL
Antibiotic-antimycotic
Ca2+-free DMEM
1%
n/a
0.5 mL
42 mL
Culture media (in Opti-MEM)
Reagent Final concentration Quantity (20 mL)
Horse serum 10% 2 mL
FBS 5% 1 mL
Antibiotic-antimycotic
Opti-MEM
0.1%
n/a
20 μL
17 mL
Note: In isolating and plating the cardiomyocyte step, high quality and optimized concentration of FBS and horse serum are critical for cell viability and cell adherence.
Laboratory supplies
Surgical scissors (Roboz, catalog number: RS-5882)
Curved forceps (Stoelting, catalog number: 52102-54P)
7 mL transfer pipette (Globe Scientific, catalog number: 135038)
24-well culture plate (Denville, catalog number: T1024)
1.5 mL micro-centrifuge tube (Stellar Scientific, T3 Tube, catalog number: T17-300)
60 mm cell culture dish (Eppendorf, catalog number: 0030701111)
70 μm mesh cell strainer (Swish, catalog number: TC70-MT-3)
15 mL centrifuge tube (Denville, Thomas Scientific, catalog number: 1158R11)
Equipment
M205 FCA microscope (Leica)
Hybridization oven/shaker (Amersham Biosciences)
Centrifuge 5425 R (Eppendorf)
TSX60086A -80 °C freezer (Thermo Scientific)
Isotemp CO2incubator (Fisher brand)
Centrifuge 5702 (Eppendorf)
Handheld automated cell counter (Millipore, Scepter 3.0, catalog number: PHCC30000)
60 μm sensors (Millipore, Scepter 3.0, PHCC360050)
-20 °C freezer (Fisher Scientific, Isotemp)
4 °C fridge (Insignia)
BZ-X800 microscope (Keyence)
Standard orbital shaker, Model 5000 (VWR, catalog number: 89032-100)
Software and datasets
ImageJ (https://imagej.nih.gov/ij/download.html)
Procedure
Cardiomyocyte isolation
Collect GFP-positive mutant hearts from E13.5–E14.5 embryos (see General note 1).
Make the solutions for the experiment and prepare a sterile workspace. Ensure that the PBS is refrigerated ahead of time and kept on ice throughout the duration of the procedure.
Anesthetize dam at E13.5–E14.5 in accordance with IACUC guidelines.
Use a scalpel to make an abdominal incision and extract the embryos with curved forceps. Separate the embryo from the placenta and yolk sac with surgical scissors. Gently remove each heart (see General note 2) using curved forceps and use a transfer pipette to move it into a 24-well plate containing cold PBS. Each heart should be placed in an individually labeled well.
Check the hearts for fluorescence under a fluorescence-emitting microscope (Figure 1) and label the wells accordingly.
Figure 1. Embryonic day (E) 13.5 hearts from the same litter (TNNT2H2B-GFP/+) imaged in brightfield and GFP fluorescence at 10× magnification. Compared to wild type (above), the TNNT2H2B-GFP/+ mouse (below) has a positive indicator for the GFP-tagged TNNT2 gene. Scale bar, 1 mm.
Isolate cells (see General note 3).
Using a transfer pipette, move each GFP-positive sample from the 24-well plate to a 1.5 mL microcentrifuge tube. All GFP-positive samples from the litter will be pooled into the same tube and digested together. Then, pipette out excess PBS with a micropipette. Add 500 μL of trypsin solution (see Recipes) to the tube and gently pipette the solution up and down with the micropipette to mix.
Note: Each heart can be dissociated individually per embryonic age and experimental needs (see General note 4).
Use tape to secure the microcentrifuge tube to the platform of a hybridization shaker. Ensure that it is positioned such that the solution can move throughout the tube and mix properly. Shake at 70 strokes/min at 37 °C for 20 min.
Remove the tube from the shaker and gently pipette the liquid up and down with a 1 mL pipette 10–15 times until there are no observable tissue pieces.
Transfer the liquid from the 1.5 mL microcentrifuge tube to a 15 mL centrifuge tube using the same micropipette. Add 5 mL of cold (4 °C) dissociation buffer (see Recipes) to the 15 mL centrifuge tube. Pipette up and down with a 7 mL transfer pipette until the cells are disaggregated in the solution.
Centrifuge at 400×gfor 5 min at 4 °C.
In a sterile vent-hood with sanitized surfaces to avoid contamination, vacuum-pipette and discard the resulting supernatant from the tube.
Add 1 mL of rinse media in Ca2+free DMEM (see Recipes) with a micropipette.
Centrifuge at 400×gfor 5 min at 4 °C.
Vacuum-pipette and discard the resulting supernatant from the tube. Add 1 mL of culture media in Ca2+free DMEM (see Recipes) with a micropipette.
Use a new micropipette to disaggregate the pellet and centrifuge again (400×gfor 5 min at 4 °C). The resulting supernatant will be used for plating and should not be discarded.
Protocol continues in step B2.
Plating cells in culture media (see General note 3)
Laminin coating
Note: This step should be completed the day before plating.
To avoid damaging the laminin via excessive freezing and thawing, pipette out laminin (final concentration: 10 μg/mL) with a micropipette and transfer to a 1.5 mL microcentrifuge tube. Return the remaining laminin to a -80 °C freezer.
Dilute the laminin with 1× DPBS (Ca2+/Mg2+) (see General note 5) and add the solution to a 24-well culture plate (see General note 6). Ensure the entire surface is covered by the laminin coating solution.
Seal the plate with parafilm to prevent evaporation and contamination.
Incubate at 2–8 °C overnight. If a more rapid coating is required, incubate at 37 °C for 2 h.
Plating the cardiomyocytes
Using a 5 mL serological pipette attachment for an assisted pipette controller, add 4 mL of culture media to an uncoated 60 mm cell culture dish.
Through a 70 μm mesh cell strainer, add 1 mL of the supernatant from the microcentrifuge tube to the cell culture dish.
Check that the cells are present in the dish under fluorescence microscope (Figure 2).
Figure 2. Visualization of cells collected from embryonic day (E) 13.5 hearts at 20× magnification plated in a 48-well cell culture plate with culture media. Brightfield imaging shows all the heart cells present in the dish, whereas GFP fluorescence identifies cardiomyocytes via the H2B-GFP tag. The combined brightfield/GFP image illustrates the ratio of non-cardiomyocyte to cardiomyocyte cells in the embryonic heart, necessitating the subsequent laminin-plating step for isolating cardiomyocytes. Scale bar, 100 μm.
Pre-plate the dish for 1 h in a cell culture incubator at 37 °C in 5% CO2to allow nonmyocytes (fibroblasts and endothelial cells) to adhere to the dish.
Centrifuge the whole dish in the Eppendorf 5702 centrifuge with bucket rotors at 400×g for 5 min.
Collect the suspended cells carefully to avoid adherent cells, as they are primarily non-cardiomyocytes (e.g., fibroblasts and endothelial cells).
Load cells into a 60 μm sensor and run through the Scepter 3.0 handheld automated cell counter. The output will determine the size of wells needed for the laminin plating.
Use a vacuum pipette to remove excess laminin solution from the premade 24-well plate.
Transfer the cells to the premade laminin-coated plate (10 μg/cm2).
Incubate at 37 °C in 5% CO2.
After 24 h, cardiomyocytes can be seen beating under a microscope (Video 1) (see General note 7). Remove the solution and replace it with serum-free media (DMEM) for one hour to starve the cells.
Video 1. Embryonic day (E) 13.5TNNT2H2B-GFP/+cardiomyocytes beating after 24 h in culture media
Wash the cells with PBS.
Treat the cells with 100 ng/mL recombinant IGF-2 protein growth factors.
Immunocytochemistry and cell counting (see General note 8)
Wash the cells with ice-cold PBS.
Fixate cells using 200 μL per well of IC fixation buffer for 10 min at room temperature.
Wash the cells three times with room temperature PBS for five minutes.
Permeabilize the sample by adding 50 µL (or enough to cover the cells) of PBS-T (PBS containing 0.1% Triton X-100) per well for 10 min at room temperature.
Wash the cells three times with room temperature PBS for five minutes.
Incubate cells with 50 µL (or enough to cover the cells) of 5% donkey serum in PBS per well for 1 h to block non-specific antibody binding.
Incubate cells with 50 µL (or enough to cover the cells) of 1:250 diluted primary antibody [p-H3 (rabbit) - donkey anti-rabbit] in 1% donkey serum in PBS per well overnight at 4 °C.
Decant the solution and wash the cells three times in PBS for five minutes.
Incubate cells with 50 µL (or enough to cover the cells) of 1:500 diluted secondary antibody [donkey anti-rabbit IgG (H+L), Alexa Fluor 555] in 1% donkey serum in PBS per well for 1 h at room temperature in the dark.
Decant the secondary antibody solution and wash the cells three times in PBS for five minutes in the dark.
Incubate cells in 50 µL (or enough to cover the cells) of 0.1 μg/mL 1:1,000 DAPI working concentration per well for 1 min.
Remove DAPI and add 200 µL of PBS to each well.
Image cell plate at 20× with Keyence microscope (Figure 3).
Figure 3. Fluorescence imaging at 20× magnification of p-H3 immunocytochemistry-stained embryonic day (E) 13.5 cardiomyocytes in serum-free media (above) and IGF-2 recombinant protein (below). When exposed to GFP fluorescence, H2B-GFP tagged cardiomyocytes are visible (column 1). Proliferating cells are identified with the p-H3 stain (column 2). The combined GFP/p-H3 image (columns 3 and 4) allows for quantification of GFP+p-H3+ vs. only GFP+ cardiomyocytes in serum free and IGF-2-treated E13.5 heart cells. Scale bar, 275 μm. Percentage of p-H3+ cardiomyocytes and total number of cardiomyocytes were quantitated. IGF2, n = 4 in each group. P-values were determined by the Mann-Whitney U test.
Both the total number of cells (Green) and pH3+ cells (Red) are counted by ImageJ image processing software. For total cardiomyocyte number, we counted GFP-positive cardiomyocyte cells in ImageJ. For percentage of p-H3 within total cardiomyocytes, we counted each of p-H3-positive cells and GFP-positive cells.
General notes and troubleshooting
General notes
E13.5 is the ideal collection day for cardiomyocytes. If the hearts are collected prior to E12.5, there may not be enough cells, whereas if the hearts are collected after E14.5 there will be too many fibroblasts present in the samples.
As fluorescence cannot be observed in the whole embryo, hearts must first be removed to check fluorescence. For the purposes of this study, the whole heart was collected, but it is possible at this stage to separate the atria or ventricles if necessary. Additionally, this protocol can be used in wildtype (WT) hearts by staining cardiomyocyte markers such as cardiac Troponin T. GFP+ cell sorting by FACS can be performed before plating the cells.
All cell culture steps require sanitized gloved hands and surfaces in a sterile fume hood to prevent contamination.
As E13.5 hearts are approximately 1–2 mm in size, 500 μL of trypsin solution is enough to digest as many hearts as are collected from one litter.
DPBS with Ca2+and Mg2+should be used to dilute the laminin, since divalent cations are important for protein structure and function.
Based on the number of cells present in the pre-plated cell culture dish, the 24-well plate can be replaced with a 48-well plate (for 1 × 104cells) or a 96-well plate (for 4 × 104cells).
Twenty-four hours after plating the isolated cells, cardiomyocytes visibly beat and will reach confluency 48 h later. In this protocol, we treated the recombinant protein under serum-free conditions and did not passage the cells over seven days. However, in normal serum conditions, we can observe the beating cardiomyocytes 14 days after isolating the cells from the heart.
For each wash step, add approximately 200 μL per well or enough to cover the cells and remove after 5 min with a vacuum pipette; then, proceed to the next step with haste to prevent the cells from drying out. Steps for the immunocytochemistry procedure are performed on a standard orbital shaker.
Acknowledgments
We gratefully acknowledge Dr. Chen-Leng Cai (Indiana University, Indianapolis, Indiana, USA) for providingTNNT2H2B-GFP/+mice. This work was partially supported by American Heart Association-Career Development Award 23CDA1046244 (J.J.), the National Heart, Lung, and Blood Institute R01 grant HL153406 (D.L.) and American Heart Association-Transformational Project Award 20TPA35490132 (D.L.). Graphical overview created with BioRender.com.
Competing interests
There are no conflicts of interest or competing interests.
Ethical considerations
Tnnt2H2B-GFP/+mice were previously described. Animal protocol was approved by the Nationwide Children’s Institutional Animal Care and Use Committee (IACUC).
References
Jang, J., Song, G., Pettit, S. M., Li, Q., Song, X., Cai, C. l., Kaushal, S. and Li, D. (2022). Epicardial HDAC3 Promotes Myocardial Growth Through a Novel MicroRNA Pathway. Circ. Res. 131(2): 151–164.
Rodgers, L. S., Schnurr, D. C., Broka, D. and Camenisch, T. D. (2009). An improved protocol for the isolation and cultivation of embryonic mouse myocytes. Cytotechnology 59(2): 93–102.
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4,803 | https://bio-protocol.org/en/bpdetail?id=4803&type=0 | # Bio-Protocol Content
Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
Endoplasmic Reticulum Isolation: An Optimized Approach into Cells and Mouse Liver Fractionation
ML Marc Leiro *
RV Raúl Ventura *
NR Nil Rojo-Querol
MH María Isabel Hernández-Alvarez
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4803 Views: 1261
Reviewed by: Davide BottaAmberley D. StephensMichael D Schultz
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Abstract
The subfractionation of the endoplasmic reticulum (ER) is a widely used technique in cell biology. However, current protocols present limitations such as low yield, the use of large number of dishes, and contamination with other organelles. Here, we describe an improved method for ER subfractionation that solves other reported methods' main limitations of being time consuming and requiring less starting material. Our protocol involves a combination of different centrifugations and special buffer incubations as well as a fine-tuned method for homogenization followed by western blotting to confirm the purity of the fractions. This protocol contains a method to extract clean ER samples from cells using only five (150 mm) dishes instead of over 50 plates needed in other protocols. In addition, in this article we not only propose a new cell fractionation approach but also an optimized method to isolate pure ER fractions from one mouse liver instead of three, which are commonly used in other protocols. The protocols described here are optimized for time efficiency and designed for seamless execution in any laboratory, eliminating the need for special/patented reagents.
Key features
• Subcellular fractionation from cells and mouse liver.
• Uses only five dishes (150 mm) or one mouse liver to extract highly enriched endoplasmic reticulum without mitochondrial-associated membrane contamination.
• These protocols require the use of ultracentrifuges, dounce homogenizers, and/or Teflon Potter Elvehjem.
• As a result, highly enriched/clean samples are obtained.
Graphical overview
Keywords: Endoplasmic reticulum (ER) Organelle isolation Cells Mouse liver Subcellular fractionation Mitochondrial-associated membranes (MAMs) Time efficient
Background
The endoplasmic reticulum (ER) is a continuous membrane system that forms a series of flattened sacs within the cytoplasm of eukaryotic cells and serves multiple functions. Some of its most relevant roles include the synthesis, folding, modification, and transport of proteins (Schwarz and Blower, 2016). The ER was first observed by electron microscopy in 1945 by Porter and Claude (Porter et al., 1945), but its isolation as a distinct organelle was not achieved until 1959 by Palade and Siekevitz (Siekevitz and Palade, 1959). In this study, authors used differential centrifugation to separate ER membranes from other cellular components. This method was later refined by adding density gradient centrifugations to obtain cleaner (non-other contaminating organelles marker) fractions (Lee et al., 2010).
The isolation of the ER has facilitated and enhanced the understanding of its structure, function, and interactions with other organelles. Some of these studies revealed that the ER forms contact sites with other membranes, such as plasma membrane–associated membranes (PAMs) (Suski et al., 2014) and mitochondria-associated membranes (MAMs) (Missiroli et al., 2018), which mediate transport of proteins, lipids, and metabolites (Ventura and Isabel Hernández-Alvarez, 2022).
However, despite being a highly common technique, the isolation of ER has some limitations. Firstly, it requires a large amount of starting material (usually several grams of tissue or cell number) and takes a long time to complete (6 h in cells because of the time needed to recover the starting material from the plates). Secondly, existing protocols may not be suitable for all tissues or cell types that have different membrane properties or distributions. Thirdly, it is quite difficult to obtain pure samples, and thus achieving highly enriched fractions can be challenging.
Most already existing methods are able to isolate ER indirectly, but only a very small number of protocols are dedicated to the extraction of this particular organelle (Croze and Morré, 1984). This scenario can be found in some of the published protocols that use mouse liver, such as the ones described by Wieckowski et al. (2009) or Suski et al. (2014), where they propose a method to isolate specific areas of the ER, such as MAMs and PAMs, respectively. This fact can also be a limiting factor.
In regard to HeLa cells, no protocols were found to specifically extract the ER as well. Moreover, some protocols that work with similar adherent cells require a large number of dishes (over 50 dishes) (Wieckowski et al., 2009; Williamson et al., 2015), which complicates the isolation and the previous cell culture work. Additionally, a similar situation happens with subcellular fractionations that use liver tissue. According to published protocols, fractionations of liver tissue are performed using rat liver rather than mouse liver, and they usually require from 8 to 10 g of tissue (Suski et al., 2014). When extrapolated to a mouse model, five or six animals for every subfractionation are needed.
Since our laboratory is mainly focused on the study of mitochondrial dynamics, ER–mitochondria contacts, and its relationship with metabolic diseases (Hernández-Alvarez et al., 2019), a protocol that overcame some of these limitations was necessary. Hence, in this article, we provide an optimized and upgraded version of already existing protocols for ER isolation achieving a high throughput using the minimum amount of starting sample. Furthermore, the protocols described below are optimized for time efficiency and designed for seamless execution in any laboratory, eliminating the need for special/patented reagents. Other protocols do not report the yield of ER.
Materials and reagents
Biological materials
Mouse liver or cell line of interest
Antibodies
Mouse monoclonal anti-PDI (C-2) (Santa Cruz Biotechnology, catalog number: sc-74551)
Rabbit monoclonal anti-Tom20 (D8T4N) (Cell Signalling Technology, catalog number: 42406)
Mouse monoclonal anti-VDAC1 (B-6) (Santa Cruz Biotechnology, catalog number: 390996)
Mouse monoclonal anti-Tim23 (H-8) (Santa Cruz Biotechnology, catalog number: sc-514463)
Rabbit monoclonal anti-FACL4 (EPR8640) (Abcam, catalog number: ab155282)
Mouse monoclonal anti-Na+/K+ ATPase alpha 3 subunit (Sigma-Aldrich, catalog number: 05-369-25UG)
Reagents
Isolation from cells
HEPES (Merck, Sigma-Aldrich, catalog number: 54461)
D-Sucrose (Thermo Fisher Scientific, Fisher bioreagents, catalog number: BP220-1)
EGTA (Merck, Calbiochem, catalog number: 324626)
KCl (Honeywell, Fluka analytical, catalog number: 31248)
Isolation from mouse liver
D-Mannitol (Merck, Calbiochem, catalog number: 443907)
D-Sucrose (Thermo Fisher Scientific, Fisher bioreagents, catalog number: BP220-1)
Albumin from bovine serum (BSA) (Merck, Sigma-Aldrich, catalog number: A7906)
EGTA (Merck, Calbiochem, catalog number: 324626)
Tris hydrochloride (Tris-HCl) for buffer solutions (PanReac AppliChem, catalog number: A1087)
10× PBS
NaCl (Thermo Fisher Scientific, catalog number: 11916388)
Na2HPO4·2H2O (Thermo Fisher Scientific, catalog number: 15613040)
KH2PO4 (Merck, catalog number: 104873)
KCl (Honeywell, Fluka analytical, catalog number: 31248)
Solutions
Microsome isolation buffer cells (MIB-C) (see Recipes)
Microsome isolation stability buffer (MIB-S) (see Recipes)
5× swelling buffer (SB) (see Recipes)
Microsome isolation buffer liver (MIB-L) (see Recipes)
10× PBS (see Recipes)
Recipes
Microsome isolation buffer liver (MIB-L)
Reagent Final concentration Quantity
D-Mannitol 225 mM 0.41 g
D-Sucrose 75 mM 0.26 g
EGTA (100 mM; pH 8) 0.5 mM 0.05 mL
Tris-HCl (1 M, pH 6.5) 30 mM 0.3 mL
BSA 0.5% 0.05 g
MQH2O n/a Up to 10 mL
Total n/a 10 mL
Microsome isolation buffer cells (MIB-C)
Reagent Final concentration Quantity
Tris-HCl (1 M, pH 6.5) 30 mM 450 μL
D-Mannitol 225 mM 0.61 g
D-Sucrose 75 mM 0.38 g
MQH2O n/a Up to 15 mL
Total n/a 15 mL
Microsome isolation stability buffer (MIB-S)
Reagent Final concentration Quantity
D-Sucrose 0.25 M 1.2 g
KCl 25 mM 0.028 g
HEPES (1 M) 10 mM 150 μL
EGTA (100 mM; pH 8) 1 mM 150 μL
MQH2O n/a Up to 15 mL
Total n/a 15 mL
5× swelling buffer (SB)
Reagent Final concentration Quantity
KCl 125 mM 0.14 g
HEPES (1 M) 50 mM 750 μL
EGTA (100 mM; pH 8) 5 mM 150 μL
MQH2O n/a Up to 15 mL
Total n/a 15 mL
10× PBS
Reagent Final concentration Quantity
NaCl 1.5 M 400 g
Na2HPO4·2H2O 0.1 M 80 g
KH2PO4 15 mM 10 g
KCl 25 mM 10 g
MQH2O n/a up to 5 L
Total n/a 5 L
Equipment
Corning® tissue culture–treated culture dishes (150 mm × 25 mm) (Merck, Corning®, catalog number: CLS430599)
Active Motif dounce homogenizer (Fisher Scientific, Active Motif 40401, catalog number: NC0569256)
NuncTM cell scrapers (Thermo ScientificTM, catalog number: 179707PK)
L-90K ultracentrifuge (Beckman coulter, catalog number: 8043-30-1191)
Type 90 Ti fixed-angle titanium rotor (Beckman coulter, catalog number: 355530)
Centrifuge 5810R (Eppendorf, catalog number: 5811000015)
15 mL polypropylene centrifugation tube (Thermo ScientificTM, catalog number: 17627105)
8.9 mL ultracentrifuge tube (Beckman coulter, catalog number: 361660)
Heidolph RZR 2051 control homogenizer (Fisher Scientific, catalog number: FIS13-880-115)
Tissue grinders, Potter Elvehjem type with PTFE pestle for soft tissue, 15 cm3 (Avantor, VWR®, catalog number 432-5041)
Western blotting equipment (Bio-Rad)
Procedure
ER isolation from HeLa cells (Figure 1)
Figure 1. Graphical summary of the endoplasmic reticulum (ER) isolation process using HeLa cells
Cell collection, rinsing, and scraping
For this protocol, five 150 mm dishes are scraped one at a time directly on the laboratory bench on ice or a cold surface (at 4 °C). In the meantime, while one dish is scrapped, the other dishes must remain in the incubator at 37 °C. Before scraping, aspirate the media and then add an approximate volume of 8 mL of PBS 1× to rinse the dish. After gently swirling to ensure the PBS covers the entire surface of the dish, aspirate the PBS and then add 4.5 mL of ice-cold MIB-S buffer. Once the MIB-S buffer is added, the dish is ready to get scrapped with a manual scraper, just in one direction.
⚠ Important step: The MIB-S buffer must be kept cold to ensure its proper functioning and facilitate the process.
Notes:
i. Ensure that all surfaces and equipment are disinfected with 70% ethanol before use.
ii. A standard cell culture incubator with controlled CO2 levels should be used. If not possible, use a 37 °C incubator (without CO2 control), ensuring that cells are kept only for a short period of time to prevent their unwanted detachment.
As a result of the first scraping, a dish containing a cell suspension in 4.5 mL of buffer will be obtained. At this point, the next dish will be rinsed as explained in previous steps and, instead of adding an additional 4.5 mL of buffer, the cell suspension of the first dish will be reused to scrape the following dish.
⚠ Important step: The 4.5 mL of MIB-S buffer is reused along all the five dishes. Hence, the same volume of buffer will be used to accumulate the cells from all the dishes.
Note: For this step, trypsin can also be used to detach and collect the cells. In this case, cells must be centrifuged, rinsed with PBS, and later resuspended with 4.5 mL of MIB-S buffer.
Finally, transfer the cell suspension to a 15 mL polypropylene centrifugation tube.
Note: The final volume obtained is expected to be approximately 6–8 mL due to the accumulation of the cells of each five plates.
Low speed centrifugations and incubations
Centrifuge the polypropylene tube containing the cell suspension at 600× g for 5 min at 4 °C. The pellet obtained will correspond to the cells collected in the last step.
Note: This centrifugation and all the following ones are performed at 4 °C as stated in section General notes and troubleshooting.
Measure the approximate volume (usually 0.5 mL) of the cell pellet obtained and prepare three times this volume of SB.
Aspirate the supernatant and resuspend the pellet in the volume calculated in the previous step of cold SB. In the case of having 0.5 mL, 1.5 mL of SB will be prepared.
Note: To resuspend, use a 1 mL Pasteur pipette to prevent breaking the cells.
Incubate the cell suspension at 4 °C for 35 min.
Once the incubation is complete, centrifuge again at 600× g for 5 min. This time, the resulting pellet will be comprised of swollen cells.
Carefully aspirate the supernatant and resuspend with 8 mL of MIB-S buffer to avoid osmotic disequilibrium.
Homogenization
To obtain the different fractions, cells must be first homogenized to preserve organelle structure. For this process, pour the 8 mL of cell suspension obtained in step A2f into a dounce homogenizer and homogenize using a tight pestle (using a 15 cm3 homogenizer) for 20 strokes.
⚠ CRITICAL: This is an important step, and the success of the protocol hinges on the precision with which it is carried out. For it to work correctly, the pestle must be pushed up and down while twisted (or rotated) simultaneously in the same motion. It is extremely important to perform this movement carefully in order to avoid the formation of bubbles.
Note: Before using the homogenizer and its pestle, rinse them with normal and ultra-pure water. Homogenization must be done on ice to minimize protease effects.
The result from the homogenization will be called Total Homogenate. Store a sample of approximately 150 μL at -20 °C to use as a control for the western blot analysis.
Low speed centrifugations: cell debris removal
Transfer the Total Homogenate to a 15 mL polypropylene centrifugation tube and centrifuge at 600× g for 5 min. The pellet obtained after centrifugation will correspond to cell debris and will be discarded.
Transfer the supernatant obtained in the last step to a new 15 mL polypropylene centrifugation tube and centrifuge again at 600× g for 5 min. This step is used to further clean the sample. As in the previous step, only the supernatant is kept.
Obtaining the main fractions
Transfer the supernatant obtained in step A4b to a new 15 mL polypropylene centrifugation tube and centrifuge at 1,200× g for 4 min. After this centrifugation, the resulting pellet will correspond to a sample highly enriched in the plasma membrane. In order to keep this fraction, transfer the supernatant to a new 8.9 mL ultracentrifuge tube and resuspend the pellet in 150 μL of MIB-C buffer. Store this sample at -20 °C for further western blot analysis and label it as Plasma Membrane sample.
Note: After this step, every centrifugation is performed in an ultracentrifuge using the 90Ti fixed-angle rotor from Beckmann.
Centrifuge the supernatant at 7,000× g for 15 min. The result from this centrifugation will be a pellet that corresponds to mitochondria and mitochondrial-associated membranes (MAMs) enriched fraction. As in the previous step, transfer the supernatant to a new 8.9 mL ultracentrifuge tube and resuspend the pellet in 150 μL of MIB-C buffer. Store the resuspended pellet at -20 °C for further western blot analysis and label it as Mitochondria and MAMs sample.
Note: It is possible to further isolate pure mitochondria and pure MAMs from this fraction by following the protocol described by Wieckowski et al. (2009).
Centrifuge the supernatant from the previous step at 20,000× g for 35 min. The result from this centrifugation will be a pellet that is mainly composed of plasma membrane contamination. As previously mentioned, just like in the previous step, transfer the supernatant to a new 8.9 mL ultracentrifuge tube and resuspend the pellet in 150 μL of MIB-C buffer. Store the resuspended pellet at -20 °C for further western blot analysis and label it as Plasma Membrane Contamination sample.
Centrifuge the supernatant from the previous step at 100,000× g for 65 min. Finally, the result from this centrifugation will be a highly enriched endoplasmic reticulum pellet. To store the sample, aspirate the supernatant and resuspend the endoplasmic reticulum pellet in 150 μL of MIB-C at -20 °C. Results of this fractionation as well as a graphical guide of this protocol can be observed in Figure 2 and Figure 3.
Figure 2. Graphical overview of key steps in the subcellular fractionation protocol using HeLa cells
Figure 3. Western blot results of subcellular fractionation protocol using HeLa cells. For this validation, 20 μg of protein were loaded in a 10% SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) gel. Each fraction was tested with antibodies against specific proteins using a 1:1,000 dilution. Secondary antibodies were also diluted following the same ratio in 5% milk diluted in PBS. In relation to the proteins tested, endoplasmic reticulum (PDI), mitochondria (outer membrane: TOM20, VDAC/inner membrane: Tim23), mitochondria-associated membranes (MAMs) (FACL4) and plasma membrane (Na+/K+ ATPase) were tested. These proteins were specifically chosen as they are markers of typically contaminating organelles in these protocols. TH: total homogenate; PM: plasma membrane; PMC: plasma membrane contamination; MM: mitochondria and MAMs; ER: endoplasmic reticulum.
ER isolation from HeLa cells: adding more dishes
When conducting experiments that require a larger pure ER sample size, it may be necessary to add more dishes. However, it is important to keep in mind that as more dishes were added, more contamination (meaning the presence of less pure fractions, not in terms of microorganisms) was observed along our experiments. According to our analyses, the cleanest sample is obtained using five dishes, with the possibility of increasing this to 10, with the following considerations:
4.5 mL of MIB-S Buffer 1× was used per each set of five dishes. This means that, in this case, a total of 9 mL of this buffer were used only in the scraping part.
Instead of using 8 mL of MIB-S buffer 1× for resuspension in step A3a, 16 mL of buffer was used instead.
Bigger polypropylene centrifugation tubes were used.
In section 5, the total volume was split into two ultracentrifuge tubes and centrifuged using the same rotor as described in the last section. Each pellet was resuspended using 100 μL of MIB-C and both resuspensions were added together in the end (we will call this making a pool).
Even though we observed that the protocol is still quite stable when using 10 dishes, we believe that this should be further improved. In these kinds of scenarios, we recommend performing several fractionations from five dishes simultaneously and making pools of reticulum. On average, we got to obtain approximately 250–300 μg of endoplasmic reticulum for each five dishes; so, if the quantity needed is 1 mg, a more effective and cleaner approach would be to have 20 dishes and perform this protocol four times simultaneously.
ER isolation using different cell lines
While the fractionation protocol was originally optimized for HeLa cells, we have observed that it can be adapted to other cell types. The main modification involves adjusting the homogenization parameters, including the type of homogenizer and the number of strokes, to account for differences in membrane hardness. As an example, we successfully isolated reticulum from mouse embryonic fibroblast (MEF) cells using an automatic homogenizer at 4,000 rpm for 20 strokes (Figure 4). These findings demonstrate the flexibility of the protocol and its potential to be tailored to other cell types.
Figure 4. Western blot results of subcellular fractionation protocol using mouse embryonic fibroblast (MEF) cells. For this validation, 20 μg of protein were loaded in a 10% SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) gel. Each fraction was tested with antibodies against specific proteins using a 1:1,000 dilution. Secondary antibodies were also diluted following the same ratio in 5% milk diluted in PBS. In relation to the proteins tested, endoplasmic reticulum (PDI), mitochondria (Tim23), mitochondria-associated membranes (MAMs) (FACL4), and plasma membrane (Na+/K+ ATPase) were tested. These proteins were chosen as they are markers of typically contaminating organelles in these protocols. TH: total homogenate; PM: plasma membrane; PMC: plasma membrane contamination; MM: mitochondria and MAMs; ER: endoplasmic reticulum.
ER isolation from mouse liver (Figure 5)
Figure 5. Graphical summary of the ER isolation process using mouse liver
Liver homogenization
Euthanize the animal by cervical dislocation, extract the liver in one piece, and remove the gallbladder. Immediately wash the liver two times in ice-cold PBS to remove the blood.
Transfer the liver and wash it again with ice-cold PBS. Then, cut the liver into small pieces (0.2 cm approximately) using scissors.
Transfer the liver pieces to the 15 mL glass/Teflon Potter Elvehjem homogenizer. Add 4 mL of MIB-L per gram of liver.
Note: Usually, a normal liver weighs 1.5 g and therefore 6 mL of MIB-L is typically needed. We used the protocol in fed and fast animals, and it worked in both conditions. This was performed in mice fed with a normal diet. The results of this protocol can vary when mice are treated with different diets, especially those that cause alterations in the lipidic composition of the liver. Usually, 500 μL at 15 μg/μL of ER is obtained.
Low-speed centrifugations
Transfer the homogenate to a 15 mL polypropylene centrifugation tube and centrifuge at 600× g for 5 min at 4 °C. In this step, nuclei, unbroken cells, and cell debris sediment at the bottom of the tube.
Notes:
i. This centrifugation and all the following ones are performed at 4 °C as stated in General Notes and Troubleshooting.
ii. In order to perform a correct ER isolation, it is important to aspirate the lipid before transferring it to a new centrifuge tube. This can be observed as a thin white layer floating on top of the supernatant.
Discard the generated pellet and transfer the supernatant to a new 15 mL polypropylene centrifugation tube. This tube is then centrifuged at 600× g for 5 min to remove remaining cell debris.
Note: As in the previous step, the lipid layer found floating at the top of the tube must be discarded.
Discard the pellet, transfer the supernatant to a new 8.9 mL polypropylene centrifugation tube, and centrifuge it at 9,000× g for 10 min using the Beckmann 90-Ti fixed angle rotor. This separates ER (supernatant) from crude mitochondria and MAMs, which in the end are going to be observed as a pellet. This fraction can be saved for analysis purposes as Mitochondria and MAMs.
Note: It is possible to further isolate pure mitochondria and pure MAMs from this fraction by using the protocol described by Wieckowski et al. (2009).
High-speed centrifugations
Centrifuge the supernatant at 20,000× g for 30 min using the same rotor as in the previous step. In this case, the pellet will contain mainly plasma membrane and other membranes with similar density characteristics.
Note: If a lipid layer is formed, discard it as in previous steps.
Transfer the supernatant to a new 8.9 mL polypropylene centrifugation tube and centrifuge at 100,000× g for 1 h with the same rotor.
Finally, aspirate the supernatant and resuspend the pellet with 500 μL of MIB-L. This pellet corresponds to a highly enriched ER fraction and will be stored at -20 °C for further analysis. Results of this fractionation as well as a graphical guide of this protocol can be observed in Figure 6 and Figure 7.
Figure 6. Graphical overview of key steps in the subcellular fractionation protocol using mouse liver
Figure 7. Western blot results of the subcellular fractionation protocol using mouse liver. 20 μg of protein was loaded in a 10% SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) gel. Each fraction was tested with antibodies against specific proteins using a 1:1,000 dilution. Secondary antibodies were also diluted following the same ratio in 5% milk diluted in PBS. In relation to the proteins tested, endoplasmic reticulum (PDI), mitochondria (outer membrane: TOM20, VDAC / inner membrane: Tim23), mitochondria-associated membranes (MAMs) (FACL4), and plasma membrane (Na+/K+ ATPase) were tested. These proteins were specifically chosen as they are markers of typically contaminating organelles in these protocols. TH: total homogenate; ER: endoplasmic reticulum.
Data analysis
In accordance with existing literature, the verification of this technique is qualitative rather than quantitative. Hence, the validity of this technique depends on observing markers specific to ER in the corresponding lanes and ensuring that no other organelles’ markers are present in this same sample. Therefore, western blots were used as the main method of monitoring enrichment since quantitative methods were not needed.
In our approach for subcellular fractionation, we used markers for endoplasmic reticulum (PDI), mitochondria (TOM20 for outer membrane and Tim23 for inner mitochondrial membrane), MAMs (FACL4), and plasma membrane (Na+/K+ ATPase). MAMs are comprised of both the mitochondria and endoplasmic reticulum. Therefore, the presence of a small quantity of MAMs in the ER fraction is to be expected. However, a large quantity of MAMs in the ER section would indicate a contaminated, non-pure reticulum fraction.
Validation of protocol
To achieve reproducibility, we fine-tuned the protocol by producing three replicates of our results. Additionally, three independent isolations on different days were performed to ensure the robustness of the protocol. The western blot analysis images presented previously in this protocol correspond to the optimal results achieved. These can be observed below for HeLa cells (Figure 8) and mouse liver (Figure 9) extracts:
Figure 8. Validation of the protocol using HeLa cells. 20 μg of protein was loaded in a 10% SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) gel. Each fraction was tested with antibodies against specific proteins using a 1:1,000 dilution. Secondary antibodies were also diluted following the same ratio in 5% milk diluted in PBS. In relation to the proteins tested, endoplasmic reticulum (PDI), mitochondria (outer membrane: TOM20), mitochondria-associated membranes (MAMs) (FACL4), and plasma membrane (Na+/K+ ATPase) were tested. Every western blot corresponds to an independent isolation. TH: total homogenate; ER: endoplasmic reticulum.
Figure 9. Validation of the protocol using Mouse Liver. 20 μg of protein was loaded in a 10% SDS-PAGE (sodium dodecyl sulphate polyacrylamide gel electrophoresis) gel. Each fraction was tested with antibodies against specific proteins using a 1:1,000 dilution. Secondary antibodies were also diluted following the same ratio in 5% milk diluted in PBS. In relation to the proteins tested, endoplasmic reticulum (PDI), mitochondria (outer membrane: TOM20), mitochondria-associated membranes (MAMs) (FACL4), and plasma membrane (Na+/K+ ATPase) were tested. Every western blot corresponds to an independent isolation. TH: total homogenate; ER: endoplasmic reticulum.
General notes and troubleshooting
General notes
In the case of the subcellular fractionation performed with cells, the quantity of reticulum obtained is approximately 300 μL for approximately 2.5 μg/μL. This amount can be sufficient for some analysis, such as checking the presence of some other proteins in that organelle, but might not be enough for more complex analysis.
All the buffers must contain a protease inhibitor cocktail and each sample must be kept at 4 °C (or on ice) to avoid degradation. For this reason, every centrifugation must be done at 4 °C as well.
The liver protocol was performed in mice fed with a normal diet. The results of this protocol can vary when mice are treated with different diets, especially those that cause alterations in the lipidic composition of the liver. Usually, 500 μL at 15 μg/μL of ER is obtained.
This protocol uses a fixed-angle rotor. We recommend the use of these types of rotors for these purposes, even though other models can also be used. Note that different rotors may alter the quality of the purification.
Troubleshooting
Cell confluence is very important and must be approximately 80%–90%; otherwise, the amount of sample obtained can be severely compromised. To solve this issue, one more dish can be added in case the others are not so confluent, in order to obtain an amount similar to what would be obtained with five dishes.
The results may change depending on the state of the cells. Cells with a high passage or that are undergoing senescence can cause different outcomes from the ones shown in this protocol. The recommended passage of HeLa cells should be approximately 1–80.
If an abnormal contamination in the fractions is observed, the problem may rely on the homogenization step. The speed, strength, temperature, and uniformity of homogenization and bubble formation during this process are common causes for this issue. Also, note that these characteristics as well as the number of strokes can be adapted for any specific laboratory conditions.
If, after the two low speed centrifugations (step 4 for the HeLa cells part, or 2a and 2b for the mouse liver part), a larger pellet is observed, the supernatant can be centrifuged again to clean the sample even more.
In the case of abnormal plasma membrane contamination in HeLa cell fractionation, an additional centrifugation can be performed just after step A5a following these same parameters.
Abnormal contamination can also occur if the supernatant is taken using a very thin tip. To avoid unwanted breakage, it is ideal to handle the supernatants with a tip that has a diameter of approximately 0.5 mm. Pellets that will be re-used for further purification steps should also be handled with these types of tips.
In most cases, the degree of contamination that the final result shows is mainly due to a bad homogenization. Ensure no bubbles are formed during this step and everything is performed on ice to avoid degradation. To improve this step, the number of strokes can be fine-tuned by homogenizing the cells and checking the degree of breakage using trypan blue stain and observing it under the microscope. A good degree of breakage can be considered when approximately 90% of the cells observed are broken.
After the homogenization, when the cells or the tissue homogenate are centrifuged, if a big pellet is observed, this can mean that there is a big volume of cells that has not been properly homogenized. In this case, the pellet can be resuspended using the same buffer as it was before the homogenization and a second homogenization can be done. The result of this second homogenization can be either added to the first one or you can perform the protocol in parallel with the first one and join the final pellets. This second way has been observed to be more successful, even though no exhaustive analysis has been done to prove its efficacy with certainty.
Some of the pellets can be a bit looser; for this reason, a lack of a band in the western blot analysis can be due to an accidental aspiration of some of the pellet.
If low yield is observed, this may be due to a low amount or bad quality of starting material (stressed cells, tissue undergoing degradation, apoptotic cells, …).
As a troubleshooting technique, the cell debris can be analyzed under the microscope or via western blot to determine if the homogenization was performed correctly.
Acknowledgments
We thank Maya Saidu Momot for comments and the critical reading of this manuscript.
Funding: This study was supported by research grants from MICINN (PID2019-105466RA-I00 AEI/ 10.13039/501100011033 and RYC2018-024345-I MCIN/AEI/ 10.13039/501100011033) and “la Caixa” Foundation, Health Research Grant 2021 (LCF/PR/HR21/52410007).
Competing interests
The authors are not affected by any conflict of interest.
Ethical considerations
All animal work was approved and conducted according to guidelines established. This project has been assessed favorably by the Institutional Animal Care and Use Committee from Parc Científic de Barcelona (IACUC-PCB) and the IACUC considers that the above-mentioned project complies with standard ethical regulations and meets the requirements of current applicable legislation (RD 53/2013 Council Directive; 2010/63/UE; Order 214/1997/GC).
References
Schwarz, D. S. and Blower, M. D. (2016). The endoplasmic reticulum: structure, function and response to cellular signaling. Cell. Mol. Life Sci. 73(1): 79–94.
Porter, K. R., Claude, A. and Fullam, E. F. (1945). A study of tissue culture cells by electron microscopy. J. Exp. Med. 81(3): 233–246.
Siekevitz, P. and Palade, G. E. (1959). A Cytochemical Study on the Pancreas of the Guinea Pig. J. Biophys. Biochem. Cytol. 5(1): 1–10.
Lee, Y. H., Tan, H. T. and Chung, M. C. M. (2010). Subcellular fractionation methods and strategies for proteomics. Proteomics 10(22): 3935–3956.
Suski, J. M., Lebiedzinska, M., Wojtala, A., Duszynski, J., Giorgi, C., Pinton, P. and Wieckowski, M. R. (2014). Isolation of plasma membrane–associated membranes from rat liver. Nat. Protoc. 9(2): 312–322.
Missiroli, S., Patergnani, S., Caroccia, N., Pedriali, G., Perrone, M., Previati, M., Wieckowski, M. R. and Giorgi, C. (2018). Mitochondria-associated membranes (MAMs) and inflammation. Cell Death Dis. 9(3): e1038/s41419-017-0027-2.
Ventura, R. and Isabel Hernández-Alvarez, M. (2022). Endoplasmic Reticulum: A Hub in Lipid Homeostasis. In: Updates on Endoplasmic Reticulum. IntechOpen.
Croze, E. M. and Morré, D. J. (1984). Isolation of plasma membrane, Golgi apparatus, and endoplasmic reticulum fractions from single homogenates of mouse liver. J. Cell. Physiol. 119(1): 46–57.
Wieckowski, M. R., Giorgi, C., Lebiedzinska, M., Duszynski, J. and Pinton, P. (2009). Isolation of mitochondria-associated membranes and mitochondria from animal tissues and cells. Nat. Protoc. 4(11): 1582–1590.
Williamson, C. D., Wong, D. S., Bozidis, P., Zhang, A. and Colberg‐Poley, A. M. (2015). Isolation of Endoplasmic Reticulum, Mitochondria, and Mitochondria‐Associated Membrane and Detergent Resistant Membrane Fractions from Transfected Cells and from Human Cytomegalovirus‐Infected Primary Fibroblasts. Curr. Protoc. Cell Biol. 68(1): ecb0327s68.
Hernández-Alvarez, M. I., Sebastián, D., Vives, S., Ivanova, S., Bartoccioni, P., Kakimoto, P., Plana, N., Veiga, S. R., Hernández, V., Vasconcelos, N., et al. (2019). Deficient Endoplasmic Reticulum-Mitochondrial Phosphatidylserine Transfer Causes Liver Disease. Cell 177(4): 881–895.e17.
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Determination of Antibody Activity by Platelet Aggregation
HL Halina H.L. Leung
JP Jose Perdomo
ZA Zohra Ahmadi
BC Beng H. Chong
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4804 Views: 576
Reviewed by: Meenal SinhaLuis Alberto Sánchez Vargas Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Communications Sep 2022
Abstract
Platelets play an important role in hemostasis by forming clots and stopping bleeding. In immune thrombotic conditions, platelets and leukocytes are aberrantly activated by pathogenic antibodies resulting in platelet aggregates and NETosis, leading to thrombosis and thrombocytopenia. A simple assay that assesses platelet function and antibody activity is light transmission aggregometry. This assay can be used to determine antibody activity in patients with disorders such as heparin-induced thrombocytopenia (HIT) and vaccine-induced thrombotic thrombocytopenia (VITT). Briefly, for detection of pathogenic antibody, platelet-rich plasma (PRP) is treated with a specific agent (e.g., patient sera or purified patient antibodies) with constant stirring. Upon activation, platelets undergo a shape change and adhere to each other forming aggregates. This causes a reduction in opacity allowing more light to pass through PRP. Light transmission through the cuvette is proportional to the degree of platelet aggregation and is measured by the photocell over time. The advantage of this protocol is that it is a simple, reliable assay that can be applied to assess antibody activity in thrombotic conditions. Light transmission aggregometry does not require the use of radioactive reagents and is technically less demanding compared with 14C-serotonin release assay, another common assay for detecting antibody activity.
Key features
• This protocol can be used to assess platelet function and to detect platelet activating antibodies in diseases such as HIT and VITT.
• Does not require radioactive reagents, requires an aggregometer; based on the light transmission aggregometry protocol, adapted for detection of VITT and other platelet-activating antibodies.
• Two positive controls are required for reliable detection of antibodies in diseases such as HIT/VITT, namely a weak HIT/VITT antibody and a physiological agonist.
• For detection of HIT/VITT antibodies, it is essential to use donors known to have platelets reactive to these antibodies to avoid false negative results.
Keywords: Platelet activation Antibody Aggregation Immune thrombosis Heparin-induced thrombocytopenia Vaccine-induced thrombotic thrombocytopenia
Background
Platelets play an essential role in hemostasis, bleeding, and thrombosis. Upon activation, platelets undergo shape changes and aggregate, leading to platelet clumping. The rate and extent of aggregation can be measured by light transmission via a platelet aggregometer. Platelet-rich plasma (PRP), which is turbid, is stirred in a test cuvette. When the agonist is added, the platelets will form increasingly larger aggregates and the PRP will begin to clear, allowing more light to pass through. This increase in light transmittance is directly proportional to the amount of aggregation.
First described by Born in the 1960s (Born, 1962), light transmission aggregometry is considered a gold standard assay for platelet function, often assigned as the first step for analyzing platelet dysfunction in hemorrhagic patients (Kottke-Marchant and Corcoran, 2002; Hayward, 2008; Gadisseur et al., 2009; Hayward et al., 2009; Podda et al., 2012). To detect the presence and activity of pathogenic antibodies, patient sera or purified patient IgG can be used. Purified IgG confirms that the IgG is the component that induces platelet activation, thus excluding other potential platelet activating agents in patient sera. Standardization of the use of light transmission aggregometry to assess platelet function has been published by the International Society of Thrombosis and Haemostasis (Cattaneo et al., 2013).
Another method of analyzing platelet activation is whole-blood aggregometry (Park et al., 2012; Mencarini et al., 2021). This method measures the change in electrical impedance between two electrodes to indicate platelet aggregation. Although this process accounts for the effect of all blood cells on platelet function, it cannot determine with accuracy the direct or indirect contributions of other blood cells to the platelet activation observed. To avoid confounding factors in whole blood, light transmission aggregometry using PRP is an ideal, relatively simple, and reliable method to specifically determine platelet activity.
The 14C-serotonin release assay (SRA) and the heparin-induced platelet activation (HIPA) test are also common assays used to test antibody activity (Arepally et al., 1995; Greinacher et al., 2012 and 2022). They are based on the principle that incubation of patient serum (containing the pathogenic antibody) will activate donor platelets resulting in release of intraplatelet serotonin (SRA) and platelet aggregation (HIPA). SRA is a sensitive method used to detect pathogenic antibody in heparin-induced thrombocytopenia (HIT) (Warkentin et al., 2015) and more recently, vaccine-induced thrombotic thrombocytopenia (VITT) (Leung et al., 2022). For SRA, washed platelets are incubated with radiolabeled serotonin. Upon activation, platelets release radiolabeled serotonin, and this is measured by a beta counter. Despite the high sensitivity of SRA, this method is labor intensive and technically demanding, requiring the user to have radiation training and certification, access to radiation laboratory, and waste disposal streams for the radioactive waste. Due to this, platelet aggregometry is a simpler, more convenient method of assessing platelet activation.
The HIPA test, developed in 1991, is based on the visual assessment of platelet aggregates (Greinacher et al., 1991). It is performed in microtiter plates, allowing multiple samples and platelet donors to be tested simultaneously, uses washed platelets, and is generally considered more sensitive than platelet aggregation tests (Greinacher et al., 1994; Tardy et al., 2020). Experienced personnel are required to conduct this assay, as platelet washing can over-activate platelets, giving false positive results, and estimation of platelet aggregation by eye can be imprecise. In comparison, platelet aggregometry is technically less demanding, does not rely on personnel to be experienced in identifying visual changes in platelet morphology, and the real-time trace clearly shows when platelet activation has taken place. By using platelets from donors pre-determined to be reactive to HIT antibodies, the sensitivity of the assay should approach that of HIPA (Chong et al., 1993). Platelet aggregometry has certain limitations as it cannot be used if donors have low platelet counts and, unlike whole-blood aggregometry, some sample preparation is required.
Here, we provide details on an updated version of Born’s method for determining antibody activity by platelet aggregation (Figure 1). This protocol is improved by standardization, with added appropriate controls and use of reactive donor platelets to increase accuracy, and adapted to test IgG, along with serum and plasma. This protocol can be used for research purposes, as well as in clinical laboratories for testing of HIT/VITT antibodies.
Figure 1. Schematic diagram of the platelet aggregometry assay. Centrifuged tubes show the different layers of blood cells and plasma. PPP: platelet-poor plasma; PRP: platelet-rich plasma.
Materials and reagents
15 mL centrifuge tubes (Greiner, catalog number: 188271)
1.5 mL microcentrifuge tube (Greiner Bio-One, catalog number: 616201)
Pasteur pipette (Transfer) (Labtek, catalog number: 650.050.523)
Direct detect assay-free cards (Merck, catalog number: DDAC00010-GR)
Stir bar, disposable siliconized for P/N 312 cuvettes (DKSH, Part No 311)
Cuvettes, 450 μL (DKSH, Part No 312)
100 kDa Amicon Ultra-15 centrifugal filter unit (Merck, catalog number: UFC910024)
Econo-Column® chromatography column, 1.5 cm × 10 cm (Bio-Rad, catalog number: 7374151)
Fresh or frozen plasma (stored at -80 °C) collected in ACD (BD, catalog number: 366645), trisodium citrate (BD, catalog number: 363095), or EDTA (BD, catalog number: 367839), or sera collected in clot activator/gel vacutainer (BD, catalog number: 367954) from patient for IgG purification
Fresh or frozen plasma (stored at -80 °C) collected in trisodium citrate or sera collected in clot activator/gel vacutainer from patient for direct testing in platelet aggregation
Whole blood from healthy donor, collected in trisodium citrate vacutainer (BD, catalog number: 363095)
Protein G agarose beads (SeraCare, catalog number: 5720-0002)
Dulbecco’s phosphate buffered saline (Sigma-Aldrich, catalog number: D8537)
Control agonists such as collagen (1 μg/mL), thrombin (1 U/mL), plasma sample from positive HIT patients
Sodium phosphate (Merck, catalog number: S8282)
Glycine (Astral Scientific, catalog number: BIOGB0235)
Tris (Amresco, catalog number: 0497)
HCl (Merck, catalog number: 320331)
NaOH (Sigma, catalog number: S-0899)
NaCl (Merck, catalog number: S9625)
EDTA (Invitrogen, catalog number: 15576028)
Starting buffer, pH 7.0 (see Recipes)
Elution buffer, pH 2.7 (see Recipes)
Neutralization buffer, pH 9.0 (see Recipes)
Washing buffer, pH 7.0 (see Recipes)
Equipment
Benchtop centrifuge (Spintron, model: GT-10 SV3)
Tabletop microfuge (Beckman Coulter, Microfuge 20 Centrifuge, model: B31603)
Centrifuge (Eppendorf, model: 5810 R)
Aggregometer (Chrono-Log, model: 700)
Chromatography system (GE Healthcare, model: AKTA Pure. catalog number: 29-0182-24)
Direct detect infrared spectrometer (Millipore, model: DDHW00010)
Rotatory tube mixer (Ratek, model: RSM7DC)
Magnetic stirrer (Bacto Laboratories, model: MS7-H550-Pro)
Calibrated pipettes
-80 °C freezer
4 °C fridge
Software
Aggro/Link8 software (Chronolog, USA)
GraphPad Prism 9 (GraphPad Software, USA)
Microsoft Excel (Microsoft, USA)
Procedure
Preparation of platelet-rich plasma (PRP)
Draw human whole blood into a vacuum tube with trisodium citrate.
Note: PRP is approximately 30%–40% of whole blood volume. Minimum volume of whole blood needed depends on number of assays to be performed [300 μL of PRP per assay is required, plus 500 μL to prepare platelet poor plasma (PPP) for background].
Centrifuge tube at 200× g with no brake for 10 min at room temperature.
Transfer PRP layer using a 3 mL Pasteur pipette to a 15 mL centrifuge tube.
Store the capped PRP sample at room temperature until ready for use, for no longer than 3 h.
Note: Platelets can be activated if stored at 4 °C, affecting subsequent platelet function assays.
Preparation of platelet-poor plasma
Transfer 0.5 mL of PRP to a 1.5 mL microcentrifuge tube.
Centrifuge tube at 1,000× g for 10 min at room temperature.
Transfer the supernatant to another 1.5 mL microcentrifuge tube and label the tube PPP.
Preparation of patient antibody from serum or plasma
Centrifuge patient serum or plasma at 10,000× g for 10 min at 4 °C to remove any cells or debris. Fresh or frozen patient serum or plasma collected in ACD, EDTA, or sodium citrate can be used.
Transfer a minimum of 0.5 mL of supernatant to a clean microcentrifuge tube.
Add Protein G agarose beads to patient serum or plasma at a 1:1 ratio; then, transfer the mixture to a clean 1.5 cm × 10 cm Econo-Column® chromatography column.
Incubate the column on a rotatory tube mixer at 4 °C for 2 h.
Place the column into the AKTA purifier chromatography system.
Wash the column with at least 10× the sample volume of washing buffer. Check the chromatography trace remains at baseline to confirm that the column has been washed sufficiently.
Add 100 μL of neutralization buffer to 1.5 mL microcentrifuge collection tubes and place them on ice.
Add 1× the sample volume of elution buffer to the column to elute bound immunoglobulins. Collect the fraction containing the protein peak in aliquots of 900 μL into the 1.5 mL microcentrifuge tubes containing 100 μL of neutralization buffer. Invert the tubes to mix.
Pool eluted antibody and transfer to a 100 kDa Amicon Ultra-15 centrifugal filter unit.
Centrifuge the Amicon Ultra-15 unit at 4,000× g for 15–20 min. Add 14 mL of PBS and repeat the centrifugation step twice to significantly reduce the concentration of the original buffer. Once buffer exchange to PBS is completed, transfer mixture and aliquot into 2 mL Eppendorf tubes.
Pipette 2 μL of sample into a sample membrane and 2 μL of buffer onto another membrane on a direct detect assay-free card.
Place card into the direct detect infrared spectrometer to dry and measure the sample concentration, setting the buffer as the blank. Expected sample concentration is approximately 7–10 mg/mL.
Note: Other methods such as Bradford assay can be used to determine antibody concentration. The quality of the purified IgG can be tested using assays such as enzyme-linked immunosorbent assay (ELISA) to confirm binding of purified IgG to antigen. Purity can be confirmed by SDS PAGE gel. The presence of multimeric IgG can be analyzed using size exclusion chromatography.
Keep antibody in -80 °C freezer for long-term storage until required or at 4 °C for use the following day.
Measuring antibody activity by platelet aggregation
Transfer 300 μL of PPP into a cuvette. Place cuvette into the reference slot of the aggregometer (100% light transmission, used as a blank) (Figure 2).
Figure 2. Aggregometer setup. A. Image of cuvette with stir bar. B. Location of reference slot for platelet-poor plasma (PPP), heating slots, and measuring slot for platelet-rich plasma (PRP). C. Aggregometer setup for running four samples concurrently.
Set temperature to 37 °C and stirring speed to 1,200 revolutions per minute (rpm) and select optical mode on the aggregometer.
For each agonist to be tested, transfer 300 μL of PRP into each cuvette containing a stir bar.
Place PRP cuvettes into heating slots for 2 min to warm up to 37 °C; then, place cuvettes into the measuring slot and ensure stir bars are stirring evenly.
Set baseline on the aggregometer.
Pipette reagent to be assessed into a cuvette containing 300 μL of PRP. Reagents include patient sera (5–50 μL), purified patient IgG (0.1–2 mg/mL), negative controls (equivalent amounts of sera or purified IgG from a healthy donor or buffer), or positive controls (0.5–1 U/mL thrombin, 1 μg/mL collagen, or 50 μL of weak positive HIT patient serum with 0.1 U/mL heparin to allow for detection of weak pathogenic antibodies).
Note: Agonist or control volumes should not exceed 150 μL.
Select Aggregometer, Run a new patient, or click Run new icon on the Aggro/Link software.
Acquire data for 15–30 min (or until platelet aggregation has completed).
Exporting and analyzing data
Export image and raw data in del file format from the Aggro/Link software.
Open raw data file in Excel.
Replot datapoints into GraphPad Prism.
Obtain AUC, maximum aggregation %, velocity, and latency time from Aggro/Link software.
Data analysis
Various parameters can be extrapolated from the platelet aggregometry data. The most essential parameters are amplitude of the platelet aggregation curve, indicating percentage of maximum aggregation, and latency time, indicating time of aggregation onset. Velocity of aggregation can also be analyzed by calculating percentage aggregation per unit of time (Figure 3). Area under the curve (AUC) and maximum aggregation (%) are calculated directly by the Aggro-Link software. These can also be recalculated following export of raw data and replotting using other programs such as Excel or Prism. Velocity and latency time can be extrapolated by selecting a particular aggregation % to compare between samples. It is not recommended to compare data from different days and donors. This assay is semi-quantitative.
Figure 3. Analysis of platelet aggregation data. Aggregometer traces indicating velocity of aggregation and maximum aggregation percentage following treatment with purified heparin-induced thrombocytopenia (HIT) patient antibody (shown in green). No aggregation is observed following treatment with negative controls (buffer shown in blue; normal IgGs shown in black and red).
To ensure the healthy donor’s platelets are functioning normally, at least one positive control is required. For the detection of platelet activating antibodies in diseases such as HIT and VITT, a weak positive patient serum or IgG is needed as the positive control. A positive result should be attained when using a weak control HIT/VITT antibody to ensure that the donor platelets are reactive enough to detect weak antibodies. If the donor platelets show a negative result for a known weak control HIT/VITT antibody, the donor platelets used are unlikely to detect a weak antibody, leading to false negative results. On the other hand, if the weak positive control gives a positive result, the donor platelets are suitable to detect even weak pathogenic antibodies.
Since only a proportion of healthy individuals are responsive to HIT antibodies, laboratories that regularly conduct HIT antibody testing should have several donors with reactive platelets to HIT antibodies to call upon to donate. These reactive donors are selected by prior testing, identified from a larger group of donors. Alternatively, two to four random donors of unknown platelet reactivity can be used together in the assay as at least one of the donors will have reactive platelets.
As with other platelet function assays, the platelet donor must not have taken drugs (e.g., aspirin) or foods (e.g., garlic), as these drugs/foods could suppress platelet reactivity. To detect this, a physiological agonist, often collagen 1 μg/mL, is used as an additional positive control (see Notes below for more detail). Abnormally shaped curves should be interpreted with caution and assay repeated to ensure samples (suitable donor, assay conducted within 3 h of blood collection) and/or reagents (correct pH and concentration) were prepared and functioning correctly.
Notes
Healthy platelet donors should not be taking any medications known to affect platelet function during the 10 days prior to the experiment. These may include cyclooxygenase (COX)-1 and COX-2 inhibitors, non-steroidal anti-inflammatory drugs (NSAID), ibuprofen, platelet receptor inhibitors [Abciximab (αIIbβ3), Clopidogrel (P2Y12), Prasugrel (P2Y12)], and anticoagulants. Aggregation times and extent of aggregation may vary depending on donor blood platelet concentration; therefore, positive controls are essential. Platelet aggregation assay should be conducted within 3 h of blood collection. The amount of patient sera or IgG needed varies from patient to patient depending on antibody titer, avidity, time of collection, and so on. To study the contribution of various platelet membrane receptors and/or signaling pathways to platelet aggregation, a range of physiological agonists such as arachidonic acid (thromboxane A2), adenosine diphosphate (ADP) (P2Y1 and P2Y12), collagen (GPVI and α2β1), epinephrine (α2A), or thrombin receptor activator peptide (TRAP)-6 [protease-activated receptor (PAR)-1] can be used.
The platelet aggregation assay described in this protocol is used for the detection of HIT and VITT antibodies and for assessment of platelet function in general, e.g., patients suspected of having a platelet function disorder. When this assay is used for the former purpose, specific controls and platelet donor requirements are needed, as described in the protocol. Platelet count of 250,000/μL is recommended. Platelet counts lower than 50,000/μL may result in inconsistent results and cause problems with setting the baseline.
The factors that make certain donor platelets more reactive to HIT/VITT antibodies are unknown. There have been conflicting data on whether reactivity is related to the FcγIIa R131 isoform compared to those with the H131 polymorphism. No significant correlation has been found between Fc receptor number and platelet response. Platelet aggregation induced by other agonists (ADP, collagen) is affected by factors such as age, body mass index, triglyceride levels, and certain foods and drugs. These, together with unknown factors, are also likely to influence reactivity to HIT/VITT antibodies.
Recipes
Starting buffer, pH 7.0
20 mM sodium phosphate
Prepare 800 mL of distilled water in a suitable container.
Add 3.28 g of sodium phosphate and stir.
Adjust solution to final desired pH using HCl or NaOH.
Add distilled water until the volume is 1 L.
Store at room temperature. Stable for more than one year.
Elution buffer, pH 2.7
100 mM glycine
Prepare 800 mL of distilled water in a suitable container.
Add 7.5 g of glycine to the solution and stir.
Adjust the pH to 2.7 with HCl.
Add distilled water until the volume is 1 L.
Store at 4 °C. Stable for one year.
Neutralization buffer, pH 9.0
1 M Tris·HCl
Prepare 800 mL of distilled water in a suitable container.
Add 121.14 g of Tris base to the solution.
Adjust solution to desired pH using HCl.
Add distilled water until the volume is 1 L.
Store at room temperature or at 4 °C. Stable for six months.
Washing buffer, pH 7.0
20 mM sodium phosphate
150 mM NaCl
2 mM EDTA
Prepare 800 mL of distilled water in a suitable container.
Add 3.28 g of sodium phosphate and 8.76 g of NaCl and stir.
Add 4 μL of 0.5M EDTA.
Adjust solution to final desired pH using HCl or NaOH.
Add distilled water until the volume is 1 L.
Store at room temperature. Stable for more than one year.
Dissolve salts in ddH2O and stir with magnetic stirrer. Adjust solution to desired pH by adding HCl or NaOH. All recipes given are for 1 L of solution. Different volumes may be prepared as required.
Acknowledgments
This work was supported by grants from National Health and Medical Research Council, Australia, Program Grant APP1052616 and New South Wales Capacity Program Senior Researcher Grant RG201677 to B.H.C. This protocol was adapted from a previous publication from our group (Leung et al., 2022).
Competing interests
The authors of this manuscript declare that there are no conflicts of interest or competing interests.
Ethics
This protocol was approved by the South Eastern Sydney Human Research Ethics Committee (17/211 LNR/17/POWH/501). Informed consent was obtained from all study participants.
References
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4,805 | https://bio-protocol.org/en/bpdetail?id=4805&type=0 | # Bio-Protocol Content
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Peer-reviewed
Isolation, Purification, and Culture of Embryonic Melanoblasts from Green Fluorescent Protein–expressing Reporter Mice
MC Melissa Crawford
KB Kevin Barr
LD Lina Dagnino
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4805 Views: 548
Reviewed by: Olga KopachPhilipp WörsdörferKrishna NakuluriSreejith Perinthottathil
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Original Research Article:
The authors used this protocol in Journal of Investigative Dermatology Jul 2019
Abstract
In this article, we provide a method to isolate embryonic melanoblasts from reporter mouse strains. The mice from which these cells are isolated are bred into the ROSA26mT/mG reporter background, which results in green fluorescent protein (GFP) expression in the targeted melanoblast population. These cells are isolated and purified by fluorescence-activated cell sorting using GFP fluorescence. We also provide a method to culture the purified melanoblasts for further analysis. This method yields > 99% purity melanoblasts specifically targeted, and can be used for a variety of studies, including gene expression, clonogenic experiments, and biological assays, such as viability, capacity for directional migration, or differentiation into melanin-producing melanocytic cells.
Graphical overview
Keywords: Melanoblast isolation and culture Green fluorescent protein Cre recombinase Cell type–specific gene targeting Melanoblast markers
Background
Melanocytes are melanin pigment–producing cells that are found in the skin, inner ear, nervous system, and in the heart (Li, 2014). Melanocytes and their precursor cells, melanoblasts, originate from neural crest cells. In mice, a subset of neural crest cells becomes specified as melanoblasts and begin their migration around embryonic day (E) 10.5 from the neural crest, to ultimately colonize the entire body (Mort et al., 2015). A second wave of melanoblast formation occurs around E14.5, when neural crest cells that migrated along a ventral path detach from nerve endings and differentiate into melanoblasts (Delmas et al., 2007; Adameyko et al., 2009). Melanoblasts proliferate, migrate, and interact with their surrounding environment to localize to specific sites in the body. By E16.5, melanoblasts from both waves will have homed to the basal layer of the epidermis, as well as the developing hair follicles (Mort et al., 2015). Understanding the ontogeny of melanocytic cells is a key area in developmental biology with significant implications for human health, as evidenced by the plethora of human diseases linked to defects in melanocytic lineage cells. In this protocol, we describe the isolation from reporter mice of embryonic melanoblasts which express green fluorescent protein (GFP). These mice express in a Cre recombinase enzyme fused to a modified estrogen receptor (CreET2). In these animals, CreET2 expression is restricted to melanocytic lineage cells, as its transcription is under the control of the Tyrosinase promoter. Tamoxifen administration to these mice induces CreET2 nuclear translocation in melanoblasts, allowing activation of GFP expression. As a result, targeted melanoblasts can be identified through GFP fluorescence. This protocol can also be used to purify and characterize embryonic melanoblasts in which a gene of interest flanked by loxP sites can be inactivated by Cre recombinase (Crawford et al., 2017). In this protocol, we also describe methods to culture the embryonic melanoblasts using ST2 feeder layers. Specifically, seeding the purified melanoblasts onto a confluent monolayer of ST2 cells maintains their viability and supports their maturation into melanin-producing cells (Crawford et al., 2020). This system allows studies including single-cell analyses and whole-genome screens of melanoblasts. Under the conditions used in our cultures, melanoblasts remain viable for at least seven days and transition into melanin-producing cells.
Materials and reagents
Biological materials
Mice. The experiments we describe are conducted with mice generated by breeding the reporter strain Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J (hereafter termed ROSAmT/mG, The Jackson Laboratory, strain number 007676) (Muzumdar et al., 2007) with B6.Cg-Tg (Tyr-cre/ERT2)13Bos/J (hereafter termed Tyr::CreERT2, The Jackson Laboratory, strain number 012328) (Bosenberg et al., 2006). Our protocol uses melanoblasts isolated from timed-pregnant dams that are homozygous for the ROSAmT/mG allele and homozygous for Tyr::CreERT2. These reporter mice can be further bred into other backgrounds in which a gene of interest is flanked by loxP sites and can therefore be used to generate GFP-tagged melanocytic cells in which a gene of interest has also been inactivated upon administration of tamoxifen (Crawford et al., 2020). Non-targeted cells can be identified by their expression of mTomato fluorescent protein. These experiments require time-pregnant mice, which are first treated with tamoxifen at 11.5 days post-coitus (dpc; midday of the vaginal plug is considered 0.5 dpc), as described below in the procedure.
ST2 cells. Bone marrow stroma–derived cell line (Riken BioResource Research Center, catalog number: RCB0224). ST2 cells express collagens and fibroblast growth factors and support melanoblasts in culture (Abdallah et al., 2019).
Reagents
Tamoxifen (Millipore Sigma, catalog number: T-5648)
Absolute ethanol (Millipore Sigma, catalog number: 493511)
70% ethanol (Millipore Sigma, catalog number: 63350-M)
Bovine serum albumin fraction V (Wisent, catalog number: 800-095-EG)
Cremophore/Kolliphor EL (Millipore Sigma, catalog number: C5135)
MGM-4 basal medium (Lonza, catalog number: CC-3250)
MGM-4 SingleQuot kit with supplements & growth factors (Lonza, catalog number: CC-4435)
Endothelin 3, lyophilized (ET3) (Lonza, catalog number: CC-4510)
2.5% trypsin (Invitrogen, catalog number: 15090046)
Trypsin neutralizing solution (Life Technologies, Thermo Fisher Scientific, catalog number: R-002-100)
Fetal bovine serum (FBS) (Thermo Fisher Scientific, Gibco, catalog number: 26140079)
0.25% trypsin/EDTA (Invitrogen, Thermo Fisher Scientific, catalog number: 25200072)
7-Amino-actinomycin D (7-AAD) (Millipore Sigma, catalog number: A9400)
Paraformaldehyde (PFA) (Millipore Sigma, catalog number: 158127)
Sterile RPMI medium (Millipore Sigma, catalog number: R8758)
NaCl (Millipore Sigma, catalog number: S5150)
KCl (Millipore Sigma, catalog number: P5405)
Na2HPO4 (Millipore Sigma, catalog number: S7907)
KH2PO4 (Millipore Sigma, catalog number: P5655)
NaOH (Millipore Sigma, catalog number: S8045)
Triton X-100 (Millipore Sigma, catalog number: X100-100ML)
Mouse Anti-TRP1 antibody (Abcam, catalog number: 3312)
Goat anti-mouse IgG Alexa Fluor® 594 Conjugate (Thermo Fisher Scientific, catalog number: A-11032)
Mounting medium with DAPI (ibidi, catalog number: 50011)
Solutions
Tamoxifen stock solution (50 mg/mL) (see Recipes)
Melanoblast culture medium (see Recipes)
ST2 culture medium (see Recipes)
Sterile Ca2+- and Mg2+-free phosphate-buffered saline (PBS), pH 7.4 (see Recipes)
FACS buffer: sterile Ca2+- and Mg2+-free PBS containing 2% FBS (see Recipes)
7-Amino-actinomycin D (7-AAD) stock solution (see Recipes)
7-Amino-actinomycin D (7-AAD) working staining solution (see Recipes)
1 N NaOH solution (see Recipes)
4% paraformaldehyde (PFA) (see Recipes)
Recipes
Tamoxifen stock solution (store at -20 °C)
Reagent Final concentration Quantity
Tamoxifen 50 mg/mL 25 mg
Absolute ethanol 50% 250 μL
Cremophore/Kolliphor 50% 250 μL
Total n/a 500 μL
Melanoblast culture medium (store at 4 °C)
Reagent Final concentration Quantity
MGM-4 basal medium n/a 500 mL
MGM-4 SingleQuot Kit n/a 1 kit (9 mL)
ET3 255 ng/mL 130 μg
Total n/a 509 mL
ST2 cell culture medium (store at 4 °C)
Reagent Final concentration Quantity
RPMI medium 90% 450 mL
FBS 10% 50 mL
Total n/a 500 mL
Ca2+- and Mg2+-free phosphate-buffered saline (PBS, store at 4 °C)
Reagent Final concentration Quantity
NaCl 137 mM 8 g
KCl 2.7 mM 0.2 g
Na2HPO4 10 mM 1.44 g
KH2PO4 1.8 mM 0.24 g
Total n/a to 1,000 mL
Autoclave to sterilize.
FACS buffer (store at 4 °C)
Reagent Final concentration Quantity
Sterile Ca2+- and Mg2+-free PBS 98% 98 mL
FBS 2% 2 mL
Total n/a 100 mL
Keep sterile.
7-Amino-actinomycin D (7-AAD) stock solution (store at -20 °C, protected from light)
Reagent Final concentration Quantity
7-AAD 1 mg/mL 1 mg
Methanol 5% 50 μL
Sterile Ca2+- and Mg2+-free PBS 950 μL
Total n/a 1 mL
7-Amino-actinomycin D (7-AAD) working staining solution (freshly prepared)
Reagent Final concentration Quantity
7-AAD stock solution 1% 100 μL
FACS buffer 99% to 10 mL
Total n/a 10 mL
1 N NaOH solution
Reagent Final concentration Quantity
NaOH 1 N 40 g
H2O n/a to 1,000 mL
Total n/a 1,000 mL
4% Paraformaldehyde (PFA) solution
Reagent Final concentration Quantity
PFA 4% 0.4 g
Ca2+- and Mg2+-free PBS n/a to 10 mL
Total n/a 10 mL
Prepare just prior to use.
Laboratory supplies
Bacterial grade 100 mm Petri dishes (Fisher Scientific, catalog number: FB0875713A)
Surgical scissors (Fine Science Tools, catalog number: 14002-12)
Two sets of forceps (Fine Science Tools, catalog number: 11000-12)
Scalpel (Fine Science Tools, catalog number: 10003-12)
Scalpel blades (Fine Science Tools, catalog number: 10011-00)
Sterile 15 mL conical tubes (Sarstedt, catalog number: 62.554.502)
Sterile 50 mL conical tubes (Sarstedt, catalog number: 62.547.205)
2 mL microcentrifuge tubes (VWR, catalog number: 20170-098)
Sterile 10 mL disposable plastic syringes (BD Biosciences, catalog number: 302995)
Sterile 21 G needles for syringe (BD Biosciences, catalog number: 305165)
Sterile 5 mL round-bottom snap cap tubes (BD Falcon, catalog number: 52054)
Sterile 40 μm nylon cell strainers (Fisher Scientific, catalog number: 08-771-1)
Neubauer hemocytometer (Hausser Scientific, catalog number: 15170-173)
BioLite T25 flasks, cell culture treated (ThermoFisher Scientific, catalog number: 130189)
35 mm μ-dishes with a 4-well insert (ibidi, catalog number: 80466)
Pipet-aid (Thermo Fisher Scientific, S1 Pipet Filler, catalog number: 9521)
Micropipettes (Eppendorf Research plus, P20, P200, P2000)
12 mm glass coverslips (Mandel Scientific, catalog number: NEU-GG-12)
Equipment
Stereomicroscope (Leica Stereo Zoom GZ6E)
Phase contrast and fluorescence inverted microscope with camera and humidified CO2 37 °C chamber for time-lapse videomicroscopy (Leica DMIRBE or equivalent)
Becton Dickinson FACSAria III cell sorter (BD Biosciences)
Becton Dickinson FACSCanto flow cytometer (BD Biosciences)
Centrifuge (Eppendorf 5810R)
Biosafety cabinet (Microzone BK-2-4)
CO2 cell culture incubator (VWR, catalog number: 10710-944)
Water bath (PolyScience, catalog number: WBE05A11B)
Rocking platform (Bio-Rad, catalog number: 166709 or equivalent)
Software
FACSDiva Software (v. 8.0.1) or equivalent
Volocity software for cellular imaging and analysis (PerkinElmer), or equivalent
Procedure
Preparation of tamoxifen stock solution
Weigh 25 mg of tamoxifen placing it in a microfuge tube. Caution: Handle the tamoxifen with protective gloves and in a fume hood to avoid inhalation.
Add 250 μL of absolute ethanol.
Vortex and incubate this suspension at 55 °C with vigorous shaking for 5–10 min, ensuring complete dissolution.
Caution: Using protective gloves, add 250 μL of Cremophor using a 1 mL sterile serological pipette, as Cremophor is highly viscous.
Mix by vigorous vortexing.
Store at -20 °C in single-use aliquots of 100 μL.
In vivo activation of CreERT2 recombinase
Immediately prior to use, thaw rapidly a 100 μL aliquot of tamoxifen stock solution (50 mg/mL) by incubating at 37 °C for 1–2 min. Caution: Use protective gloves when handling the tamoxifen solution.
Following institutional biosafety regulations, add 400 μL of sterile Ca2+- and Mg2+-free PBS to 100 μL of tamoxifen stock solution (50 mg/mL), to yield a final concentration of 10 mg/mL of tamoxifen (i.e., 1 mg/100 μL). Vortex to mix well, protect from the light, and use promptly to avoid tamoxifen precipitation from the solution.
Note: The final volume of diluted tamoxifen administered to each mouse will depend on its body weight and will be approximately 100 μL (steps B3 and B4).
Weigh the time-pregnant 11.5 dpc mice to be treated with tamoxifen.
Briefly restraining the conscious mice with one hand (Machholz et al., 2012), administer intraperitoneally (IP) the PBS-diluted tamoxifen solution at a dose of 1 mg/25 g body weight.
Note: Administration of tamoxifen in Cremophor EL-containing solutions reduces variability in pharmacokinetic parameters, including plasma concentrations and half-life (Chevalier et al., 2013). A single IP injection is required.
Return the mice to their housing cage, allowing normal housing conditions for four days.
Culture of ST2 stromal cells
ST2 cells are maintained in ST2 culture medium. Procedures to process cell cultures from T25 tissue culture flasks are described below. All steps are conducted aseptically in a laminar flow biosafety cabinet.
Once ST2 cultures reach 80% confluence, remove the growth medium by aspiration.
Gently add 5 mL of warm (37 °C) sterile Ca2+- and Mg2+-free PBS to rinse the cell monolayer and remove by aspiration.
Trypsinize the cells, adding 1 mL of 0.25% trypsin/EDTA solution, and place the cells in the cell culture incubator.
Examine the extent of cell detachment by phase contrast microscopy every 2–4 min.
Once ~ 90% of the cells have detached, add 7 mL of warm (37 °C) growth medium to inactivate the trypsin and gently mix the cell suspension to generate a single-cell suspension.
Transfer 1 mL of the cell suspension into a new T25 cell culture flask containing 5 mL of warm (37 °C) growth medium. Mix gently and place the cells in the incubator.
To maintain exponential cell proliferation, aspirate the growth medium every third day and replace with 5 mL of fresh growth medium. Do not allow the cultures to reach over 80% confluence.
Preparation of ST2 feeder cell monolayers for melanoblast culture
Trypsinize ST2 cells cultured in a T25 flask, as described in steps C1–C5.
Transfer the cell suspension to a 15 mL conical tube.
Centrifuge the cells at 200× g for 5 min at 22 °C. Aspirate the supernatant and gently resuspend the cell pellet in 1 mL of warm (37 °C) RPMI growth medium.
Determine the cell density using a hemocytometer and adjust to 70,000 cells/mL by adding the appropriate volume of warm RPMI growth medium.
Add 100 μL of the cell suspension to each well of a 4-well insert placed in a 35 mm μ-dish (7,000 ST2 cells/well) and culture overnight.
Note: The use of μ-dishes with optical grade plastic allows high quality imaging and optimal cell attachment and spreading.
The day after plating, and just prior to isolation and purification of targeted melanoblasts (see below), remove ST2 growth medium by aspiration.
Add 100 μL of warm (37 °C) fresh culture medium consisting of a 1:1 mixture of melanocyte growth medium and ST2 RPMI growth medium to each well.
Place the ST2 feeder cells back in the culture incubator.
Isolation of targeted embryonic melanoblasts
Four days after single-dose IP administration of tamoxifen (step B4), euthanize time-pregnant dams at 15.5 dpc by CO2 inhalation or isoflurane overdose, following appropriate institutionally approved animal care ethics procedures.
Lay the animal in a supine position on an absorbent pad and soak the thorax and abdomen with 70% ethanol.
Using a pair of forceps, gently pinch outwards the abdominal skin below the center of the belly and, using surgical scissors, make a small incision across the midline, ensuring not to cut through the abdominal cavity.
Cut the skin along the ventral midline and gently pull the skin to each side to separate it from the underlying abdominal cavity, thus exposing the uterus (Figure 1).
Using forceps, gently remove the uterus containing the string of embryos out of the body cavity. Transfer the uterus to a Petri dish containing sterile PBS kept at room temperature (22 °C). Separate the embryos by carefully cutting between implantation sites along the uterine horn (Figure 1).
Embryo isolation is conducted under a stereomicroscope. To this end, transfer a single embryo and associated uterine tissues to another Petri dish containing PBS. Maintaining the embryo in PBS, use a pair of forceps to peel away the muscular uterine layers from the embryo, followed by removal of the decidua and the ectoplacental cone.
With a pair of forceps, gently tear the yolk sac and fully separate it from the embryo by gently tearing the umbilical cord.
Transfer embryos to a sterile tissue culture dish containing PBS and place on ice. At this stage, a small fragment of the embryo tail can be removed and processed for genomic DNA extraction and genotyping if necessary (Figure 1).
Figure 1. Schematic representation of procedure to isolate individual E15.5 mouse embryos
Transfer an embryo to a Petri dish containing sterile PBS placed on the stage of the stereomicroscope.
Holding the embryo in a prone position with forceps, gently make a superficial longitudinal dorsal incision along the midline with a scalpel below the head and through the entire carcass, avoiding deep incisions that will result in exposure of the organs.
Using a pair of forceps, gently and gradually separate the skin from the carcass, starting at the incision point closest to the neck.
Note: Embryonic skin is very fragile. Use one set of forceps to hold the embryo through its head and the other set to gently separate the skin from the carcass.
Float each harvested skin in a sterile Petri dish containing ice-cold sterile PBS. Continue harvesting the skin of all remaining embryos.
Carefully transfer with forceps the embryonic skins into a sterile conical tube containing 500 μL of sterile 2.5% trypsin per skin.
Note: If analysis of individual embryos is necessary, each skin can be digested in individual 2 mL microfuge tubes containing 500 μL of 2.5% trypsin and then minced.
Mince skin using small scissors into fragments of approximately 2 mm × 2 mm and incubate at 37 °C for 10 min with gentle rocking.
Add 1 mL of trypsin neutralizing solution for every skin digested and mix gently.
Filter the cell suspension through a sterile 40 μm pore size cell strainer into a 15 mL sterile conical tube to remove tissue debris.
To obtain a single cell suspension, gently pass the filtered cell mixture 10 times through a 5 or 10 mL syringe fitted with a 21 G needle.
Centrifuge at 140× g for 5 min at 22 °C.
Carefully remove the supernatant by aspiration, avoiding disturbing the cell pellet.
Resuspend the embryonic cells in 300–500 μL of ice-cold FACS sorting buffer.
Note: Resuspending the cell suspension in larger volumes of FACS sorting buffer will result in decreased flow rates and increased sorting times, leading to decreased cell viability.
Transfer the cell suspension into 5 mL BD Falcon glass tubes and place on ice.
Immediately proceed to cell staining and FACS purification of melanoblasts.
FACS purification and culture of targeted, GFP-positive melanoblasts
Approximately 15 min prior to fluorescence-activated sorting of melanoblasts, add 7-AAD stock solution to the melanoblast suspension (5 μL of 7-AAD stock solution for every 450 μL of cell suspension), to obtain a final concentration of 10 μg/mL, and mix gently.
Note: 7-AAD will stain apoptotic and necrotic cells, allowing their separation from the viable melanoblasts to be cultured.
For FACS purification of melanoblasts, set up sorting at 4 °C, using a 100 μm nozzle at low pressure (20 psi), and at a maximum event rate of 2,000 events/s.
Note: Increasing the event rate will decrease the recovery yield of GFP-positive cells.
Configure the blue laser trigon to detect GFP from detector B (530/30 bandpass and 502 long pass filter). The yellow-green laser octagon is configured to detect mTomato from detector C (610/20 band pass and 600 long pass filter). 7-AAD is detected from detector B (670/14 bandpass and 630 long pass filter).
Exclude cell doublets using consecutive gates on side scatter (SSC)-height vs. SSC-width plots and forward scatter (FSC)-height vs. FSC-width (Figure 2A and 2B).
The FSC and SSC detectors are set in a linear scale, whereas the fluorescence channels are set to logarithmic scale. Use the appropriate voltage on each detector to generate a cell population with a wide distribution but ensure that all events remain visible in the FSC vs. SSC plots for the GFP fluorescence channel, as exemplified in Figure 2.
Figure 2. FACS gating strategy to purify targeted melanoblasts from single cell suspensions of embryonic skin cells. (A, B) Cell doublets are excluded using consecutive gates on side scatter (SSC)-height vs. SSC-width plots and forward scatter (FSC)-height vs. FSC-width. (C) Cells with low 7-AAD staining are selected to exclude non-viable cells. (D) Cells gated for GFP expression are selected as targeted melanoblasts for subsequent culture.
Select viable cells based on the exclusion of 7-AAD (Figure 2C).
Run the GFP-negative control samples and set gates for GFP-positive and negative regions.
Create a region of GFP-positive, 7-AAD-negative cells and display on a plot of mTomato vs. GFP as shown in Figure 2D. These are the viable GFP-positive melanoblasts that have been targeted.
Sort cells into sterile tubes containing ice-cold melanoblast culture medium.
Run a small aliquot of the sorted cell suspension in the sorter to verify that > 95% of purified cells are GFP-positive.
Seal the tubes containing the remainder of the sorted melanoblasts using their caps and transport into the laminar flow hood for subsequent seeding.
Seed the sorted GFP-positive melanoblasts into the insert wells of the 35 mm μ-dishes containing the previously prepared confluent monolayers of ST2 feeder cells (section D) and culture at 37 °C.
Note: The average number of GFP-positive melanoblasts obtained per embryo is approximately 500 (ranging from 50 to 1,400). Melanoblasts obtained from 4–8 embryos can be pooled into one well.
Sixteen hours after sorting, gently remove the growth medium aspirating with a pipette to avoid disturbing the cell monolayer and replace with fresh growth medium.
Note: Sixteen hours after seeding, approximately 70% of melanoblasts will be fully spread and begin to exhibit dendricity (Figure 3).
Figure 3. GFP-expressing melanoblasts. Targeted, GFP-expressing melanoblasts isolated from tamoxifen-treated reporter mice (15.5 dpc) were isolated, and FACS-sorted cells were seeded on ST2 cell feeder layers and cultured for 16 h. GFP fluorescence and GFP/phase contrast overlay micrographs show the GFP-positive melanoblasts. Note confluent monolayer of ST2 cells around and underneath the GFP-expressing melanocyte, visible in the phase contrast micrograph. Scale bar, 25 μm.
Replace the growth medium every 72 h.
Note: Remove growth medium gently using a pipette to avoid disturbing the cell monolayer. Melanoblasts will mature into melanin-producing cells and become highly dendritic approximately within one week after isolation.
Basic characterization of melanocytic lineage identity
The melanocytic lineage cell-restricted expression of Cre-ERT2 recombinase under the regulation of the Tyr promoter and its tamoxifen-dependent activation to induce melanocytic cell-specific expression of fluorescent reporter proteins (e.g., GFP, tdTomato) has been documented in several independent reports (Bosenberg et al., 2006; Crawford et al., 2017 and 2020; Sun et al., 2023). In addition, cultured melanoblasts can be morphologically identified by their characteristic ability to develop highly dynamic dendritic extensions (Figure 3). Although E15.5 melanoblasts are amelanotic, they express the specific transcription factor MITF-M (Thomas and Erickson, 2008), which can be detected by immunofluorescence microscopy. Further, 4–5 days after isolation, cultured melanoblasts will mature into melanocytic cells with dendritic projections that contain melanin granules distributed throughout the cell body (Figure 4). At this stage, the cells also express enzymes involved in melanin biosynthesis, such as tyrosinase and tyrosinase-related protein 1 (TRP1; Figure 4). The protocol to process these cultures for TRP1 detection by indirect immunofluorescence microscopy is described below.
Figure 4. Expression of GFP and tyrosinase-related protein 1 (TRP1) in purified melanoblasts. Targeted, GFP-expressing melanoblasts isolated from tamoxifen-treated reporter mice (15.5 dpc) were isolated, FACS-sorted, and seeded on ST2 cell feeder layers. The cells were cultured for seven days and processed for immunofluorescence microscopy. Two different melanoblasts are shown. Brightfield micrograph panels (labeled “Melanin”) illustrate dendritic cell morphology and presence of dark melanin granules distributed throughout the cell. Scale bar, 25 μm.
Place the culture dish with melanoblasts cultured for 3–7 days on ST2 feeder cells on ice and remove the insert with forceps, taking care to avoid disturbing the cell monolayer.
Remove the culture medium by aspiration.
If used, remove the insert from the culture dish.
Gently add 1 mL of ice-cold PBS to rinse the cells and remove by aspiration.
Fix the cells by adding 1 mL of freshly prepared 4% PFA in PBS and incubate with gentle rocking for 20 min at room temperature (22 °C).
Remove the PFA solution by aspiration.
Rinse once by adding 1 mL of PBS, as in step G4.
Note: Fixed cells can be processed immediately or stored in PBS at 4 °C for up to a week.
Permeabilize the cells by adding 1 mL of 0.1% Triton X-100 in PBS and incubate for 10 min at room temperature with gentle rocking.
Rinse thrice with PBS, as in step G4.
Mix 40 μL of goat serum with 756 μL of PBS and add 4 μL of the primary anti-TRP1 antibody. Mix well.
Note: The primary antibody can also be diluted in PBS containing 3% bovine serum albumin in place of goat serum.
Add the anti-TRP1 antibody solution from step G10 to the cells and incubate at 4 °C overnight.
Remove the anti-TRP1 antibody solution by aspiration.
Wash the cells by incubating with 2 mL of PBS for 10 min at room temperature (22 °C), with gentle rocking. Remove the PBS by aspiration.
Repeat step G13 two additional times.
Mix 40 μL of goat serum with 756 μL of PBS and add 1–2 μL of the secondary Anti-mouse IgG Alexa Fluor® 594 Conjugate. Protect sample from light from this step onwards.
Add the secondary antibody solution to the cells and incubate for 1 h at room temperature (22 °C) with gentle rocking and protected from the light.
Note: The secondary antibody can also be diluted in PBS containing 3% bovine serum albumin in place of goat serum.
Wash the cells by incubating with 2 mL of PBS for 10 min at room temperature, with gentle rocking. Remove the PBS by aspiration.
Repeat step G17 two additional times.
Immediately after aspirating the PBS from the last wash, gently add 50 μL of mounting medium to the center of the culture dish.
Gently place a 12 mm glass coverslip on the mounting medium, ensuring there are no bubbles.
Place samples flat and store at room temperature (22 °C) overnight, protected from light.
Samples are ready for fluorescence imaging.
Processed samples should be subsequently stored at 4 °C, protected from light.
Validation of protocol
This protocol was validated in: J. Invest. Dermatol. (2020), DOI: 10.1016/j.jid.2019.07.681
Acknowledgments
We thank K. Chadwick for expert help with FACS experiments. The authors acknowledge financial support for this work to LD from the Natural Sciences and Engineering Research Council and the Canadian Institutes of Health Research. Studies on melanoblasts isolated and cultured following this protocol have been published in Crawford et al. (2020).
Competing interests
The authors declare no competing interests or conflicts of interest.
Ethical considerations
All animal experiments were approved by the University of Western Ontario Animal Care Committee (Protocol No. 2018-169), according to regulations and guidelines of the Canadian Council on Animal Care.
References
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Recoil Measurements in Drosophila Embryos: from Mounting to Image Analysis
LS Luis Eduardo Sánchez-Cisneros
SB Sourabh Bhide
LR Luis Daniel Ríos-Barrera
Published: Vol 13, Iss 14, Jul 20, 2023
DOI: 10.21769/BioProtoc.4806 Views: 528
Reviewed by: Alberto RissoneKai Yuan Anonymous reviewer(s)
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Original Research Article:
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Abstract
Tension and force propagation play a central role in tissue morphogenesis, as they enable sub- and supra-cellular shape changes required for the generation of new structures. Force is often generated by the cytoskeleton, which forms complex meshworks that reach cell–cell or cell–extracellular matrix junctions to induce cellular rearrangements. These mechanical properties can be measured through laser microdissection, which concentrates energy in the tissue of interest, disrupting its cytoskeleton. If the tissue is undergoing tension, this cut will induce a recoil in the surrounding regions of the cut. This protocol describes how one can perform laser microdissection experiments and subsequently measure the recoil speed of the sample of interest. While we explain how to carry out these experiments in Drosophila embryos, the recoil calibration and downstream analyses can be applied to other types of preparations.
Key features
• Allows measuring tension in live Drosophila embryos with a relatively simple approach.
• Describes a quick way to mount a high number of embryos.
• Includes a segmentation-free recoil quantification that reduces bias and speeds up analysis.
Graphical overview
Keywords: Laser cuts Recoil measurements Tension measurements Laser microdissection Biomechanics Particle image velocimetry
Background
Tissue morphogenesis requires that cells change their individual shapes and rearrange with one another to form novel structures. Most often, these processes depend on tension and force propagation to induce cell shape changes (Barrera-Velázquez and Ríos-Barrera, 2021; Inman and Smutny, 2021). Forces can act within the same cell or specific subcellular compartments, but can also act in neighboring cells with different biochemical and mechanical properties (Bhide et al., 2021; Ríos-Barrera and Leptin, 2022).
Cytoskeletal elements like actin are often the molecular basis of force generation. Actin fibers interact with motors and crosslinkers to generate force that induces cell autonomous and non-autonomous shape changes (Belmonte et al., 2017; Miao and Blankenship, 2020). Nevertheless, the sole presence of actin meshworks within a cell or tissue may not be indicative of tissue tension. To show that a given structure is under tension, one must manipulate or measure the mechanical properties of the tissue of interest. Different tools have been developed to do this, with particular advantages and caveats. Optogenetics (Izquierdo et al., 2018; Krueger et al., 2019) has a high temporal and spatial specificity but requires the incorporation of genetic tools that are sensitive to light, requiring special handling conditions and restricting imaging possibilities. Atomic force microscopy (Haase and Pelling, 2015) provides a direct readout of tissue properties but uses dedicated equipment and only works in direct contact with the specimen of interest. Fluorescent tension sensors (Lemke et al., 2019) do not require external manipulations but depend on complex image analyses to obtain quantitative information. Laser microdissection was among the first tools that enabled probing mechanical properties of tissues. It consists in focusing high-frequency photons within a region of interest (ROI), which will then disrupt and destabilize actin fibers (Nishimura et al., 2006). Severing the actin cytoskeleton using laser microdissection alters the force distribution in the cell or tissue of interest, resulting in a recoil in the direction in which the tissue is being stretched. Initial recoil speed of the tissue is directly proportional to the tension of the sample immediately before the laser procedure (Rauzi and Lenne, 2014; Shivakumar and Lenne, 2016). Therefore, by measuring the recoil speed, it is possible to infer the tension exerted in the sample of interest. This provides a quantitative approximation of the forces participating in the process of interest and can be used to compare tension generation in different directions (anisotropy), in different regions and stages of embryo development, or in varying genetic conditions (genetic gain- and loss-of-function studies).
There are several ways in which one can measure recoil speed [for an alternative approach, see Bio-protocol by Liang et al. (2016)]. In this protocol, we use particle image velocimetry (PIV) analyses, a straightforward approach that does not rely on image segmentation or manual annotation of the signal of interest (Tseng et al., 2012). Instead, PIV compares the spatial distribution of the fluorescent signal within two consecutive time points by comparing the signal within small interrogation windows within the ROI. A shift in the signal in an interrogation window is converted into a vector, which depicts the strength and direction of recoil, if there is any.
As an example of this procedure, we use tracheal terminal cells of the Drosophila respiratory system, which uses actin to coordinate their direction of migration with the formation of a subcellular tube (Ríos-Barrera and Leptin, 2022). Nevertheless, this approach can be used to study a wide range of developmental processes in the fruit fly and in other model systems.
Materials and reagents
Strains of Drosophila melanogaster expressing the actin-binding domain of Utrophin fused to GFP under the control of a UAS promoter (Flybase ID FBtp0073094) or the ubiquitously-expressed sqh promoter [Flybase ID FBtp0073095 (Rauzi et al., 2010; Bhide et al., 2021; Ríos-Barrera and Leptin, 2022)].
Paintbrush #3, round (commercially available)
Scalpel blade, #23 (e.g., EMS, catalog number: 72049-23)
Wash bottle containing distilled water (e.g., VWR, catalog number: 215-4306)
Forceps Dumont no. 5 (Merck, catalog number: F6521-1EA)
Commercial bleach 50% in water (commercially available)
35 mm glass-bottom dish plates (MatTek, catalog number: P35G-1.5-7-C) or plastic dish plates compatible with confocal microscopy (SPL Life sciences, catalog number: 210350)
Halocarbon oil 27 (Sigma-Aldrich, catalog number: H8773-100ML)
Heptane glue (see Recipes)
200 mL heptane (Sigma-Aldrich, catalog number: H2198-1L)
3M cellophane, double-sided tape (commercially available)
Embryo collection plates (see Recipes, Figure 1A)
6 cm Petri dishes (Sigma-Aldrich, catalog number: P5481-500EA)
250 mL distilled water
10 g agar (Millipore, catalog number: 9002-18-0)
4.2 g sugar molasses or brown sugar (commercially available)
250 mL apple juice (commercially available)
5 mL 10% Nipagin (Merck, catalog number: H5501-100G) in EtOH (Sigma-Aldrich, catalog number: 64-17-5)
Dried yeast (commercially available)
100 mL plastic beakers (e.g., Sigma-Aldrich, catalog number: Z245380)
Embryo basket (see Recipes, Figure 1B and 1C)
50 mL conical tube (e.g., Corning, catalog number: CLS430828)
Stainless steel mesh, 100 wires per inch (e.g., Amazon, Brand LTKJ)
Figure 1. Materials required for embryo collection. (A) 6 cm embryo collection plates and 100 mL plastic beakers are combined to make embryo collection cages. (B) Materials required to make an embryo basket. (C) Assembled embryo basket.
Recipes
Heptane glue
Mix the tape and heptane for 3 h in a rocker.
Discard the tape and use the liquid for downstream applications.
Store the heptane glue in a dropper bottle.
Embryo collection plates
Dissolve the agar and brown sugar in 250 mL of water and autoclave.
Add 250 mL of apple juice, mix, and add the Nipagin solution.
Pour into 6 cm Petri dishes and store at 4 °C.
Before using, add yeast paste (dried yeast dissolved in water).
Punch small holes on the base of the plastic beakers. These will contain the flies, and the Petri dish with yeast will serve as a lid of the embryo collection cage.
Embryo basket
Make an open-ended 5 cm cylinder by cutting the conical tube using a cutter knife.
Cut a round window on the lid of the conical tube.
Cut the metallic mesh into a circle that fits inside the lid.
Screw in the lid containing the metallic mesh back into the conical tube.
Equipment
Dissecting microscope (e.g., Velab, VE-S7)
Laser scanning confocal inverted microscope (e.g., Zeiss, LSM780) with a 63× oil immersion objective, 1.4 NA, and a femtosecond-pulsed infrared laser (Chameleon Compact OPO Fammily, Coherent).
Software
Fiji version 1.53t, https://fiji.sc (Schindelin et al., 2012)
PIV analysis plugin version #2020/04/22, https://sites.google.com/site/qingzongtseng/piv (Tseng et al., 2012)
Procedure
Embryo collection
Set up embryo collection cages by transferring flies of the desired genotype to a plastic beaker. Cover the beaker with a collection plate (see Recipes). This should be done at least three days before the day of the experiment and the plate should be replaced every day. This allows flies to adapt to the new environment and ensures that enough eggs are deposited on the day of the experiment (for additional comments on embryo collection, see General notes and troubleshooting).
One day before the experiment, collect embryos in the embryo cages for 16–24 h (i.e., overnight, Figure 2A–2C).
Replace the embryo plate with a new one.
Take the plate that contains the embryos and add enough bleach to cover the entire surface of the agar. Incubate for 1–2 min or until the chorionic membrane is completely dissolved. You can verify when this is done by looking through the dissection scope. Waggling the embryo plate can also speed up the dissolution of the chorionic membrane.
Caution: Bleach is corrosive; wear gloves and a lab coat when using it.
Transfer the dechorionated embryos to the embryo basket. To ensure that most embryos are recovered from the collection plate, wash this several times with distilled water using a wash bottle. In addition, wash the embryos with abundant water using tap water (Figure 2D).
Critical: Embryo exposition to bleach must be as brief as possible. Therefore, it is important to wash the basket with abundant water as soon as the chorionic membrane is dissolved and to do extensive washes to ensure that bleach is completely washed off the embryos. This can be confirmed by smelling the embryo basket.
Using a paintbrush, transfer the dechorionated, washed embryos to a new embryo collection plate (prewarmed to room temperature).
Under the dissecting microscope, collect embryos of the desired stage of development. In this case, take embryos that are at approximately stage 15 of development. In this stage, when looking at the embryos from the dorsal side up, the yolk is no longer heart shaped, but instead has more irregular anterior/posterior edges (Figure 2E–2E’).
Critical: When selecting the embryos, the light source might increase the temperature of the embryo plate, especially when screening using the bottom light of the microscope. To avoid this, this step should be done as quickly as possible.
Figure 2. Staging and mounting embryos. (A) Assembled embryo collection cage. (B–B’) Collection plates before (B) and after (B’) embryo collection. (C) Zoom in to an embryo collection plate; embryos are mixed with yeast paste and still have chorionic membrane. (D) Zoom in to an embryo basket, after collecting dechorionated embryos. (E–E’) Embryos aligned in an embryo collection plate, with anterior up and dorsal facing the camera. The shape of the yolk allows staging of the embryos. From left to right: first embryo is in stage 13; the yolk looks like an inverted heart. Second embryo is a stage 16 embryo; the yolk has a two lobed appearance due to folding of the midgut. Third position is an unfertilized egg. The yolk is irregular and off centered, and the rest of the egg is more transparent relative to the other embryos. The fourth embryo is stage 15; the midgut has begun to fold. The last embryo is stage 14; the yolk seems more condensed compared to stage 13 but is more distended than stage 15. (F) All-at-once mounting method. The imaging plate is carefully brought towards the aligned embryos so that they stick to the glass (1). In the same movement, the plate is then moved upwards (2) with the embryos glued to the plate. (G) Embryos attached to the glass after the imaging plate was approached to the agar block.
Mounting, one-by-one method
Pour two drops of heptane glue (See Recipes) into an imaging plate and let the heptane evaporate.
Using the paintbrush, transfer ~5 embryos of the desired stage to the imaging plate, making sure that the region of interest is facing downwards.
Critical: Embryos should be mounted as quickly as possible to prevent the embryos from dehydrating.
Add enough halocarbon oil to completely cover the embryos. Do not add the oil directly on top of the embryos, as this might detach them from the heptane glue.
Mounting, all-at-once method
Pour two drops of heptane glue into an imaging plate and let the heptane evaporate.
Using forceps, select 5–10 embryos of the desired stage and align them, making sure that the region of interest is facing upwards. Using a blade, cut a cube of agar containing the embryos. Realign them if necessary (Figure 2E).
Invert the imaging plate with the dried heptane glue and put it on top of the embryos so that they attach to the glass (Figure 2F–2G). Because of the side walls of the imaging plate, it is necessary to place the cube of agar containing the embryos on a high surface as exemplified on Figure 2F. In this example, the agar cube with the embryos was placed on top of a nail polish bottle.
Critical: Avoid putting pressure on the embryos, as this might damage them or make them burst.
Add enough halocarbon oil to completely cover the embryos. Do not add the oil directly on top of the embryos as this might detach them from the heptane glue.
Imaging and laser ablation
In the confocal microscope, focus on one of the embryos of interest. For a detailed protocol on how to perform live imaging of Drosophila embryos, refer to Araújo and Llimargas (2023).
Using the bleaching module of the microscope’s software, define a ROI that will be the area intended to cut (Figure 3A–3A’). Set the excitation wavelength at 950 nm using the two-photon laser. Shorter wavelengths (i.e., 850 and 900 nm) give similar results, but 950 nm from a two-photon laser allows to also visualize the GFP actin reporter during the course of the experiment.
Figure 3. Laser microdissection and recoil measurements. (A–B’) Tracheal terminal cell expressing the actin-binding domain of Utrophin fused to GFP (UtrABD-GFP) under btl-gal4. The black box is the ROI where the laser microdissection was or will be done. (A, A’, B) before laser cut; (A’’, B’) after laser cut. (A’) The blue box shows the region where a control particle image velocimetry (PIV) analysis was done, before the laser procedure (time points A vs. A’). (A’’) The red box shows the region where the PIV analysis was done comparing the images before and after laser ablation (time points A’ vs. A’’). Inset shows the graphical representation of the measurements exactly as generated by the plugin. (B’) After a second laser exposition in the same ROI, the sample generates an autofluorescence signal (arrowhead), evidence of phototoxicity. (C–C’’) Higher-resolution reconstructions of the insets shown in (A’–A’’). (C) Control PIV values (from images in A vs. A’). (C’) Experimental PIV values, before and after laser microdissection. (C’’) Color scale for vector values shown in (C–C’).
To calibrate the laser settings, start by using 75% laser power and 8 iterations. If this results in tissue damage, you can lower the number of iterations by half until photo-damage disappears. Tissue damage can be seen as emergence of autofluorescence or permanent tissue deformation (Figure 3B–3B’, Video 1). In contrast, laser power that is not strong enough to induce ablation will only cause bleaching of the fluorescent signal but will not rupture the cytoskeleton.
Video 1. Laser micro-dissection in the tip of a tracheal terminal cell.The black box in (A) and (B) indicates the region where laser micro-dissection was performed. (A) Adequate laser settings. (B) Excessive laser power.
Once the optimal laser settings have been set, move to a different embryo to run the experiment.
Acquire a Z-stack still image of the cell or tissue that you intend to acquire. This will aid as a higher-resolution image of the initial condition of the sample.
Define a new ROI where you intend to induce the laser cut (Figure 3A–3A’). Adjust the bleaching module of the microscope’s software so that the laser cut is done after five frames of imaging and post-acquisition of ~50 frames (total duration of approximately 90 s to acquire the complete recoil, including its plateau phase). Within this time window, acquisition time should be set as fast as possible, with a maximum interval of one image every 10 s.
Immediately after acquisition, you can acquire a new Z-stack still image to record the final morphology of the treated tissue.
Save the image in a lossless file format like .tiff or any other recommended by the microscope software (.lsm, .lif, .nd2).
Data analysis
Image analysis
Image processing is done in Fiji. First, make sure that the plugin “iterativePIV” is installed. If it is not, install it by clicking on Command Menu > Help > Update… Then go to Manage update sites and then add “https://sites.imagej.net/iterativePIV/” as a new update site. Close Manage update sites and click on Apply changes. You will have to restart Fiji afterwards. If this does not work, you can directly download the plugin from https://sites.google.com/site/qingzongtseng/ and paste it in the plugins folder within Fiji.
Define a ROI that covers the area where you expect to see an effect. In the example used here, we used 107 × 50 pixel (5 μm × 2.3 μm) boxes on one side of the laser cut. Duplicate the ROI for further analyses (Figure 3).
The PIV plugin uses two consecutive time points. To measure the initial recoil speed, duplicate into a new image the time points right before and after the laser cut. As negative control, you can use two consecutive time points before the laser cut (Figure 3A and 3C).
To run the PIV plugin, you will need to define an interrogation window and a search window size. As suggested in the plugin documentation, you can start by using ¼ of the image dimension as an initial window size and the doubled value as search window size. In this example, we used a 20 pixel window size and 30 pixel search window size.
The PIV plugin will generate a graphical representation of the displacements identified (Figure 3A’–3A’’, insets; and 3C–3C’) and a .csv file with multiple values. Average the absolute magnitude values (fifth column) of the different interrogation windows (rows in the file). This is the value we will use as initial recoil speed.
As a validation of these results, you can use complimentary image analysis tools, for instance manual tracking of the edges of the ablated area as described elsewhere (Liang et al., 2016).
General notes and troubleshooting
To increase the number of embryos retrieved from embryo collection plates, ensure that the flies are young (1–2 weeks) and that they come from healthy vials. Once in the embryo cages, ensure that the plates always have yeast paste so that the adults are well fed. This can be achieved by replacing the plate with fresh yeast paste three times per day.
This protocol explains how to identify embryos at stage 15 of development, when tracheal terminal cells begin to elongate. However, the protocol can be used for other tissues and other developmental stages. To identify the stages of interest the reader might refer to https://www.sdbonline.org/sites/fly/atlas/00atlas.htm (Hartenstein, 1993).
A way to recover more embryos of the desired stage is to synchronize embryo collections. To do this, embryos are collected for 30 min. This first collection is not used for experiments in case females had retained eggs, in which case the staging would not be precise. A second 30-min egg collection is done, and the embryos are aged until the time when they are needed. In the example presented here, embryos would be harvested after 10–12 h of incubation. This approach can help beginners to identify the developmental stages of interest for a given experiment.
Acknowledgments
LESC is a master student from Programa de Maestría en Ciencias Bioquímicas, Universidad Nacional Autónoma de México, and received a fellowship from Consejo Nacional de Ciencia y Tecnología (CVU: 1099237). Work in the lab of LDRB is supported by Universidad Nacional Autónoma de México, Programa de Apoyo a Proyectos de Investigación e Innovación Tecnológica, México (UNAM-PAPIIT) grants #IA201921 and #IA202923 and an Early Career Return grant from the International Centre for Genetic Engineering and Biotechnology, Italy, #CRP/MEX21-04_EC.
This protocol was derived from the original work of Ríos-Barrera and Leptin (2022).
Competing interests
The authors declare no competing interests.
References
Araújo, S. J. and Llimargas, M. (2023). Time-Lapse Imaging and Morphometric Analysis of Tracheal Development in Drosophila. In: Margadant, C. (Ed.). Cell Migration in Three Dimensions (pp. 163–182). Humana, New York.
Barrera-Velázquez, M. and Ríos-Barrera, L. D. (2021). Crosstalk between basal extracellular matrix adhesion and building of apical architecture during morphogenesis. Biol. Open 10(11): e058760.
Belmonte, J., Leptin, M. and François, N. (2017). A theory that predicts behaviors of disordered cytoskeletal networks. Mol. Syst. Biol. 13: 941.
Bhide, S., Gombalova, D., Mönke, G., Stegmaier, J., Zinchenko, V., Kreshuk, A., Belmonte, J. M. and Leptin, M. (2021). Mechanical competition alters the cellular interpretation of an endogenous genetic program. J. Cell Biol. 220(11): e202104107.
Haase, K. and Pelling, A. E. (2015). Investigating cell mechanics with atomic force microscopy. J. R. Soc. Interface 12(104): 20140970.
Hartenstein, V. (1993). The development of Drosophila melanogaster. Cold Spring Harbor Laboratory Press.
Inman, A. and Smutny, M. (2021). Feeling the force: Multiscale force sensing and transduction at the cell-cell interface. Semin. Cell Dev. Biol. 120: 53–65.
Izquierdo, E., Quinkler, T. and De Renzis, S. (2018). Guided morphogenesis through optogenetic activation of Rho signalling during early Drosophila embryogenesis. Nat. Commun. 9(1): e1038/s41467-018-04754-z.
Krueger, D., Izquierdo, E., Viswanathan, R., Hartmann, J., Pallares Cartes, C., and De Renzis, S. (2019). Principles and applications of optogenetics in developmental biology. Development 146(20): dev175067.
Lemke, S. B., Weidemann, T., Cost, A. L., Grashoff, C. and Schnorrer, F. (2019). A small proportion of Talin molecules transmit forces at developing muscle attachments in vivo. PLoS Biol. 17(3): e3000057.
Liang, X., Michael, M. and Gomez, G. (2016). Measurement of Mechanical Tension at cell-cell junctions using two-photon laser ablation. Bio Protoc. 6(24): e2068.
Miao, H. and Blankenship, J. T. (2020). The pulse of morphogenesis: actomyosin dynamics and regulation in epithelia. Development 147(17): e186502.
Nishimura, N., Schaffer, C. B. and Kleinfeld, D. (2006). In vivo manipulation of biological systems with femtosecond laser pulses. SPIE Proceedings: e668886.
Matteo, R., Lenne, P.F., and Lecuit, T. (2010). Planar polarized actomyosin contractile flows control epithelial junction femodelling. Nature 468(7327): 1110–14.
Rauzi, M., and Lenne, P. F. (2014). Probing Cell Mechanics with Subcellular Laser Dissection of Actomyosin Networks in the Early Developing Drosophila Embryo. Tissue Morphogenesis: Methods and Protocols 1189: 209–218.
Ríos-Barrera, L. D. and Leptin, M. (2022). An endosome-associated actin network involved in directed apical plasma membrane growth. J. Cell Biol. 221(3): e202106124.
Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., et al. (2012). Fiji: An Open-Source Platform for Biological-Image Analysis. Nature Methods 9(7): 676–82.
Shivakumar, P. C., and Lenne, P. F. (2016). Laser Ablation to Probe the Epithelial Mechanics in Drosophila. In Drosophila, edited by Christian Dahmann, 1478:241–51. Methods in Molecular Biology. New York, NY: Springer New York.
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Article Information
Copyright
© 2023 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
Biophysics > Microscopy > Two-photon laser scanning microscopy
Cell Biology > Tissue analysis > Tissue imaging
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Improve Research Reproducibility
A Bio-protocol resource
Peer-reviewed
A Large-format Polyacrylamide Gel with Controllable Matrix Mechanics for Mammalian Cell Culture and Conditioned Media Production
CM Catherine S. Millar-Haskell
JG Jason P. Gleghorn
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4807 Views: 852
Reviewed by: Masashi AsaiXin Xu Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in bioRxiv Oct 2022
Abstract
Tissue culture plastic has been used for routine cell culture and in vitro experiments for over 50 years. However, cells are mechanically responsive and behave differently on hard surfaces than they do on softer substrates. Polyacrylamide gels have become a popular hydrogel of choice for controlling surface stiffness and ligand density for cell adhesion. Many synthesis methods use coverslips and small gel surface areas for cell culture, which are amenable to microscopy-based experiments. However, none of the currently published methods can be scaled up to increase the surface area to accommodate conditioned media production, high volume analyte collection, or cell line expansion. To overcome this size limitation, we developed a protocol for synthesizing polyacrylamide in glass dishes using commercially available materials. This enables routine cell culture on soft surfaces and facilitates experiments that require large amounts of analyte, especially studies involving extracellular vesicles and secreted factors.
Graphical overview
Keywords: Polyacrylamide Stiffness Matrix mechanics Cell culture Hydrogel Extracellular matrix Extracellular vesicles Secreted factors
Background
Culturing mammalian cells on plastic is regarded as the gold standard—it is quick, easy, and inexpensive. However, the mechanical properties of plastic surfaces are not physiological, and many studies have demonstrated phenotypic, secretory, and transcriptional differences in cells cultured on compliant surfaces. Most tissues in the body are soft and have an elastic modulus (E) that falls between 1 and 100 kPa (Engler et al., 2006;Chandler et al., 2011;Gleghorn et al., 2013). Plastic is rigid and its elastic modulus measures on the order of Gigapascals (Chen and Simmons, 2011). Some cell lines are more sensitive to mechanical stimuli than others. Well established cell lines that have been cultured on plastic for many generations (e.g., NIH 3T3 fibroblasts) are less likely to be affected by routine culture as they have already been conditioned to grow on hard surfaces. Primary cells, which are isolated directly from fresh tissue, are susceptible to undesirable phenotypic and genotypic changes when they are placed in these altered mechanical environments. When cultured on plastic, cells may transition to an activated or inflammatory state, de-differentiate, or lose their primary function. Primary cells are thus passage-limited and have a relatively short expansion window before new cells must be used.
Matrix mechanics directly modulate cell signaling and function (Nelson and Gleghorn, 2012; Millar-Haskell et al., 2019; Morgan et al., 2019; Trompeter et al., 2021). This has been observed from in vitro experiments using a variety of hydrogels of different compositions and structure. Polyacrylamide gels are particularly useful because they are elastic, the stiffness can be tightly controlled by varying the concentrations of prepolymer, and a variety of extracellular matrix proteins can be conjugated to the gel for cell attachment. Polyacrylamide gel systems have yielded important observations regarding durotaxis (Lachowski et al., 2017), mechanotransduction (Stanton et al., 2019), and differentiation (Engler et al., 2006). Over the course of a decade, protocols have been published detailing the creation of polyacrylamide coverslips or platforms for cell-based experiments (Aratyn-Schaus et al., 2010;Tse and Engler, 2010; Mih et al., 2011; Syed et al., 2015; Kumai et al., 2021). These protocols present a standardized methodology, allowing experiments across research groups to be compared more easily. Generally, the process involves functionalizing a glass surface (e.g., coverslips or multi wells) to facilitate polyacrylamide attachment during polymerization. Sulfo-SANPAH is used to conjugate an extracellular matrix protein of choice to the polyacrylamide gel for cell attachment. Cells are then seeded on top of these gel surfaces for 24–72 h to study morphological and phenotypic changes. Polyacrylamide gel coverslips are useful for high resolution fluorescent and traction force microscopy and enable experiments involving single cell tracking. Modifications to the method can give rise to patterned surfaces and custom stiffness gradients (Tse and Engler, 2010). However, these protocols are not adaptable for routine cell culture or experiments that require large surface areas due to the inherently small sizes of the platforms. Additionally, they are not amenable to conditioned media generation for studying the release of secreted factors, RNA, extracellular vesicles, and other media components (Millar‐Haskell et al., 2022a). We focused on overcoming this size limitation to enable greater versatility in polyacrylamide gel platforms.
We have developed a method of synthesizing polyacrylamide gels inside glass dishes so that they can be used like normal tissue culture dishes (Figure 1). The process is much like casting a gel for polyacrylamide gel electrophoresis, except the gel is polymerized horizontally in a glass cell culture dish. The materials used in this protocol are easy to obtain, autoclave-safe, and re-usable. The polydimethylsiloxane (PDMS) ring and acrylic cover create a seal during polymerization, and the cover is removed after gelation. This format prevents the introduction of excess oxygen during polymerization and creates a flat surface for cell culture. Additionally, this protocol uses 145 mm glass dishes, but it can be adapted for any size, including traditionally used 100 mm glass dishes or, in cases where a custom shape is needed, by changing the dimensions of the PDMS ring. These dishes can be made in bulk and stored for later use as needed. We found that we could culture our cells (PANC-1 pancreatic cancer cells) in these dishes for up to a week with no loss of viability and minimal cell damage upon detaching them (Figure 2). Possible applications of this platform include conditioned media production, supernatant analysis (e.g., extracellular vesicle release or protein secretion), or large cell lysate collection for downstream analysis (Millar‐Haskell et al., 2022b).
Figure 1. Overview for creating polyacrylamide dishes. (1) Polyacrylamide is synthesized in a glass dish using a PDMS ring and acrylic cover. (2) The crosslinker sulfo-SANPAH is activated on the polyacrylamide with UV light and the excess is rinsed off. (3) Extracellular matrix protein is diluted in buffer and incubated on the polyacrylamide to initiate conjugation. (4) After soaking dishes in HBS to remove unconjugated protein, dishes are pre-incubated with media and are ready for cell culture. Created with Biorender.com. Adapted from Millar-Haskell et al. (2022b).
Figure 2. Cell viability assessment using calcein AM dye and flow cytometry. (A) Cartoon representation of cells in a tube after trypsinization with the presence of debris caused by cell death/compromised membranes. Cells were also stained using a live/dead viability assay. (B) Viability of PANC-1 cells cultured on polyacrylamide gel dishes of different elastic modulus and tissue culture plastic. Viability was quantified by percentage of calcein AM+ cells using flow cytometry. (C) Cell debris generation (which corresponds to cell damage and death) was normalized as a percentage of total cell count. Example plot with cell and debris gating is shown.
Materials and reagents
15 mL conical tubes (CELLTREAT, catalog number: 667016B)
Cleanroom wipes (Fisher Scientific, Contec, catalog number: 19-130-5932)
Plastic spoons and clear plastic cups (recommend Amazon or any major retailer)
Feather scalpel blades #11 (Electron Microscopy Sciences, catalog number: 72044-11)
Sylgard 184 silicone elastomer kit (Krayden, catalog number: DC4019862)
145 mm glass Petri dishes (VWR, catalog number: 25354-127)
145 mm plastic Petri dishes (VWR, catalog number: 82050-600)
0.2 μm PES bottle top filter (VWR, catalog number: 10040-436)
Autoclavable sterilization pouches (VWR, Cardinal Health, catalog number: 11213-033)
Sodium hydroxide (0.1 M NaOH)
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9888)
3-aminopropyltriethoxysilane (APTES) (Fisher Scientific, ACROS organics, catalog number: AC430941000); store at 4 °C, avoid contact with air
25% glutaraldehyde (GA) (VWR, Electron Microscopy Sciences, catalog number: 100504-788); store at 4 °C. We have found that this GA can be stored for up to three years without any changes to the end result
40% acrylamide (Sigma-Aldrich, catalog number: A4058); store at 4 °C
2% bis-acrylamide (Sigma-Aldrich, catalog number: M1533); store at 4 °C
Ammonium persulfate powder (APS) (VWR, catalog number: 97064-594), store at room temperature in a desiccator, strong oxidizer
Tetramethylethylenediamine (TEMED) (VWR, catalog number: 97064-684); flammable
125 mm clear acrylic disc (127 mm/5 inch acrylic discs may be found on Amazon, or acrylic sheets may be purchased from McMaster-Carr and laser cut into any size or shape)
Sulfo-SANPAH (Covachem, catalog number: 13414-100); store at 4 °C
HEPES (Sigma-Aldrich, catalog number: H3375-500G)
Protein of interest (e.g., collagen, Corning, catalog number: 354236); store at 4 °C
Hank’s buffered salt solution (HBSS) (ScienCell, catalog number: 0313)
Milli-Q water (or equivalent grade)
Isopropanol (70% solution)
Ethanol (70% solution)
Dimethyl sulfoxide (DMSO) (Sigma-Aldrich, catalog number: 276855)
Ammonium persulfate, 10% w/v solution (see Recipes)
Sulfo-SANPAH, 100 mg/mL solution (see Recipes)
HEPES-buffered saline (HBS) (see Recipes)
Equipment
Vacuum desiccator
Balance/weighing scale
Glass beaker (VWR, catalog number: 13912-284)
Filter forceps (VWR, catalog number: 89259-954)
Scalpel handle #3 (Fine Science Tools, catalog number: 10003-12)
Pipet aid and serological pipets (10, 25, 50 mL)
Set of micropipettes and tips (P10, P200, P1000)
Oven, set at 65 °C
Chemical hood
Biosafety cabinet with vacuum aspirator
Autoclave
UV lamp (e.g., 100-Watt 365 nm UVP lamp)
CO2humidified incubator, set at 37 °C
-20 °C freezer
4 °C refrigerator
Procedure
Creation of polydimethylsiloxane (PDMS) ring molds
Using a balance, weigh Sylgard 184 elastomer base and Sylgard 184 curing agent into a plastic cup at a 9:1 ratio (e.g., 27 g of elastomer base and 3 g of curing agent).
Note: Uncured Sylgard 184 elastomer base is highly viscous.
Mix the PDMS pre-polymer thoroughly with a plastic spoon and pour into plastic 145 mm Petri dishes, making sure the surface is evenly coated. The amount of PDMS pre-polymer can be increased or decreased, but 1–1.5 mm final thickness within the Petri dish is recommended.
Degas Petri dishes in a vacuum chamber to remove air bubbles from pre-polymer mixture.
Cure the PDMS pre-polymer overnight in an oven set at 65 °C.
Note: Make sure the oven surface is leveled for a uniform PDMS thickness. As little as ~1° slope will adversely impact downstream processes.
With gloved hands, remove the cured PDMS discs from the Petri dishes and cut into rings. Using a scalpel, cut out a ring with an outer diameter of 145 mm and an inner diameter of 120 mm to serve as the gasket.
Suggestion: Trace a circle with a diameter of 120 mm on the PDMS disc using a pen or non-solvent resistant marker. The acrylic cover must overlap the PDMS to get a good seal during polymerization.
Clean PDMS rings with 70% isopropanol followed by 70% ethanol. Dry completely.
Seal PDMS rings inside an autoclavable pouch and use a 30 min sterilization cycle with a minimum temperature of 120 °C (see section F for reusability).
Glass dish surface activation
Note: This section should be performed in a chemical fume hood due to hazardous chemical use.
Add enough 0.1 M NaOH to cover the bottom of the 145 mm glass Petri dishes and let sit for 30 min.
Remove excess NaOH and let dishes dry completely.
In a glass beaker, mix APTES into isopropanol to make a 2% solution.
Pour APTES solution into dishes until surfaces are fully covered (20–30 mL/dish) and let sit for 10 min.
Note: APTES is reactive to oxygen. The container from the listed manufacturer has a regenerative rubber seal where the contents are meant to be removed via syringe needle and volumetrically replaced with an inert gas (e.g., N2). APTES loses its reactivity when exposed to molecular oxygen and will therefore become less effective over time. This step extends the shelf life of APTES.
CAUTION: APTES is corrosive; chemical-resistant gloves, lab coat, and goggles should be used.
Following the reaction, remove the APTES solution and add ~30 mL of Milli-Q water to each dish; swirl dishes to mix, let sit for 5 min, and then remove water.
Repeat this step three more times for a total of four exchanges.
For each exchange, dispose in appropriate hazardous waste stream.
Add 20–30 mL of 0.5% glutaraldehyde diluted with Milli-Q water to each of the glass dishes and let sit for 30 min.
Dispose of the glutaraldehyde solution in an appropriate hazardous waste stream and allow dishes to air dry. Proceed to polymerization step (section C) within 24 h of activation for optimal gel attachment.
Polyacrylamide gel polymerization
Note: Parts of this section are recommended to be performed in a biosafety cabinet (BSC) and/or laminar flow hood to minimize contamination. Wipe down all materials with 70% ethanol before placing them in the BSC.
Prepare pre-polymer solutions of acrylamide.
Many polyacrylamide gelrecipesare published in the literature that report the ratios of bis-acrylamide and acrylamide to produce the desired elastic modulus (Aratyn-Schaus et al., 2010;Tse and Engler, 2010; Syed et al., 2015; Charrier et al., 2020).
Using any published reference table, add 40% acrylamide, 2% bis-acrylamide, and Milli-Q water in the desired ratios in a 15 mL conical tube for a total volume of 15 mL of solution per 145 mm glass dish. Optional: Pre-polymer solutions can be made up in larger volume batches, wrapped in parafilm, and stored at 4 °C for 3–4 days. Fresh solutions should be used when possible because even minor evaporation of solution over time can cause noticeable shifts in the storage modulus upon polymerization.
Degas solutions in a vacuum chamber for at least 30 min to remove dissolved oxygen.
In a BSC, use forceps to remove sterile PDMS rings from the autoclave pouch and place them inside the activated glass Petri dishes. Press down firmly on the PDMS ring with the forceps to ensure good contact with the glass.
Spray down acrylic discs with ethanol and dry with cleanroom wipes or compressed air and set inside BSC.
Bring degassed pre-polymer solutions inside BSC. Initiate the reaction by adding ammonium persulfate (1:100 dilution) and TEMED (1:1,000 dilution).
Gently invert tube 2–3 times and pour ~15 mL into the glass Petri dish.
Place the acrylic cover on top of the PDMS ring to create a seal.
Like mounting a coverslip to a slide, grab one side of the acrylic cover with forceps and set onto the PDMS ring at an angle.
Lower the acrylic cover—watch as it makes contacts with the solution and check for any bubbles in the process.
Pipette any excess solution from around the ring, place solution back into the conical tube, and cap the tube. Use the extra solution in the conical tube to gauge when the polyacrylamide has gelled, as it will indicate when the dish is ready.
After the polyacrylamide has fully polymerized (~15 min at room temperature), remove the acrylic cover.
Aspirate unpolymerized solution at the PDMS–acrylic interface.
Note: Unpolymerized solution is expected. The overlap between the PDMS ring and the acrylic cover is designed to prevent uneven gelation at the edges and facilitate removal of the acrylic cover.
Using a P1000 micropipette, add 100% isopropanol into the PDMS–acrylic interface where the unpolymerized solution sat until it penetrates the polyacrylamide–acrylic interface. Gently lift up on the acrylic cover to help the isopropanol enter the gel-cover interface while pipetting.
Once the isopropanol has completely infiltrated the polyacrylamide–acrylic interface, the acrylic cover should pop off very easily.
Wash polyacrylamide gel three times with HBS.
Note: Do not put vacuum aspirator on or directly above the gel as it may get caught in the vacuum. Instead, place the aspirator on the PDMS ring and tip the dish so the fluid falls into the vacuum aspirator.
Polyacrylamide gel dishes can be stored at 4 °C in HBS, sealed with parafilm or plastic wrap.
Matrix protein-polyacrylamide conjugation
Note: This section is recommended to be performed in a BSC with aseptic technique whenever possible, to minimize contamination. The extracellular matrix (ECM) protein of choice (fibronectin, collagen, laminin, etc.) depends on the cell line used and will need to be tested for suitability.
Prepare working solutions of the ECM protein of interest (e.g., collagen at 100 μg/mL) by diluting in HBS.
Remove the storage buffer from the polyacrylamide dishes.
Dilute sulfo-SANPAH stock solution to 0.5 mg/mL in Milli-Q water and pipette directly on the surface of polyacrylamide gels (~5 mL). Sulfo-SANPAH hydrolyzes rapidly in water, so this step needs to be done promptly.
Place the dishes under the UV light for up to 10 min to activate the surface.
Note: Remove the lids to the dishes before placing under UV light.
Sulfo-SANPAH should turn burgundy (dark orange) when it has reacted with the polyacrylamide.
Activation time may change based on UV lamp wavelength, power, and distance of dish from light source. For the lamp listed above, a distance of three inches from the light source for 10 min is sufficient.
Wash the polyacrylamide dishes three times with HBS to remove excess sulfo-SANPAH.
Add the ECM protein solution directly on top of the polyacrylamide and place the dishes in a cell culture incubator at 37 °C overnight.
Roughly 10 mL of solution is sufficient for a uniform conjugation.
We found that an overnight incubation worked well, but less time (1–2 h) may be sufficient.
After incubating with the protein solution, wash once with HBS and replace with fresh HBS.
Soak the gels in HBS overnight (or 2–3 h) at 37 °C.
Repeat this step with one more exchange of fresh HBS to remove excess unconjugated sulfo-SANPAH and/or ECM protein if necessary.
The gel should turn to a very light transparent orange.
Collagen can be visualized under a light microscope equipped with a 10× objective, which can be used to check for any non-uniformity.
Dishes can be stored at 4 °C in HBS sealed with parafilm or plastic wrap for weeks.
Cell culture
Note: This section should be performed in a BSC with aseptic technique whenever possible, to minimize contamination.
Remove HBS and incubate dishes with fresh cell culture media for 1 h at 37 °C prior to seeding cells.
Plate cells by dripping the cell suspension over the entire surface and allow to adhere for ~1 h.
Add 25–30 mL of media once cells have adhered to the surface.
When cells are ready to be passaged, remove the cell media, add warm HBSS or serum-free media to the dishes, and let sit for 15–30 min in a cell culture incubator.
Note: This will wash the cells in preparation for trypsinization.
If cell media contains a high concentration of serum (e.g., 10% fetal bovine serum) or polyacrylamide gels are softer (< 10 kPa), more than one exchange may be necessary to remove excess protein within the gels.
Serum proteins must be adequately removed to prevent inhibition of the trypsin.
If phenol is present in cell media, the polyacrylamide gels should change from a pink color to almost clear, indicating the cells are ready to be trypsinized.
Remove HBSS or serum-free media and proceed with routine passaging.
Cleaning and reusing of the glass dish
With gloved hands, remove PDMS ring, clean with warm, soapy water, and sterilize in the autoclave. PDMS rings can be placed inside a beaker of isopropanol and/or detergent solution before sterilization to remove absorbed material.
Remove polyacrylamide from the dish. Polyacrylamide can be removed by gloved hand or with forceps.
Bleach glass dish for at least 15 min, rinse with copious water, and clean with warm soapy water.
Sterilize dishes in autoclave. Dishes are ready to be re-used (start at section B: Glass dish surface activation).
Data analysis
Cell viability and cell debris measurements were performed in technical and experimental triplicates using an Agilent NovoCyte flow cytometer. An ordinary one-way ANOVA was performed on cell viability data and Brown-Forsythe and Welch’s ANOVA were performed on cell debris data.
Recipes
Ammonium persulfate, 10% w/v solution
Dissolve in deionized water at a concentration of 10% w/v. Ammonium persulfate should crackle as it dissolves in water.
Aliquot and store at -20 °C for up to six months.
Sulfo-SANPAH, 100 mg/mL solution
Dissolve sulfo-SANPAH in DMSO at 100 mg/mL.
Aliquot in 25 μL volumes in microcentrifuge tubes and freeze at -80 °C for up to one year.
HEPES buffered saline (HBS)
Dissolve 50 mM HEPES and 0.9% w/v NaCl in Milli-Q water.
Bring to pH 8.0 with NaOH.
Sterile filter solution using 0.2 μm PES bottle top filter.
Acknowledgments
This work was funded by the National Institutes of Health (R01GM26643, R01HL133163, R01HL145147, U19AI158930). The protocol was adapted from Millar-Haskell et al. (2022). The authors would like to acknowledge John Sperduto and Bryan Ferrick for their assistance in acquiring materials.
Competing interests
The authors declare no competing interests.
Ethics
No animal or human subjects were used in this work.
References
Aratyn-Schaus, Y., Oakes, P. W., Stricker, J., Winter, S. P. and Gardel, M. L. (2010). Preparation of Complaint Matrices for Quantifying Cellular Contraction. J. Vis. Exp.: e3791/2173.
Chandler, E. M., Berglund, C. M., Lee, J. S., Polacheck, W. J., Gleghorn, J. P., Kirby, B. J. and Fischbach, C. (2011). Stiffness of photocrosslinked RGD-alginate gels regulates adipose progenitor cell behavior. Biotechnol. Bioeng. 108(7): 1683–1692.
Charrier, E. E., Pogoda, K., Li, R., Park, C. Y., Fredberg, J. J. and Janmey, P. A. (2020). A novel method to make viscoelastic polyacrylamide gels for cell culture and traction force microscopy. APL Bioeng. 4(3): 036104.
Chen, W. L. K. and Simmons, C. A. (2011). Lessons from (patho)physiological tissue stiffness and their implications for drug screening, drug delivery and regenerative medicine. Adv. Drug Delivery Rev. 63: 269–276.
Engler, A. J., Sen, S., Sweeney, H. L. and Discher, D. E. (2006). Matrix Elasticity Directs Stem Cell Lineage Specification. Cell 126(4): 677–689.
Gleghorn, J. P., Manivannan, S. and Nelson, C. M. (2013). Quantitative approaches to uncover physical mechanisms of tissue morphogenesis. Curr. Opin. Biotechnol. 24(5): 954–961.
Kumai, J., Sasagawa, S., Horie, M. and Yui, Y. (2021). A Novel Method for Polyacrylamide Gel Preparation Using N-hydroxysuccinimide-acrylamide Ester to Study Cell-Extracellular Matrix Mechanical Interactions. Front. Mater. 8: e637278.
Lachowski, D., Cortes, E., Pink, D., Chronopoulos, A., Karim, S. A., P. Morton, J. and del Río Hernández, A. E. (2017). Substrate Rigidity Controls Activation and Durotaxis in Pancreatic Stellate Cells. Sci. Rep. 7(1): e1038/s41598-017-02689-x.
Mih, J. D., Sharif, A. S., Liu, F., Marinkovic, A., Symer, M. M. and Tschumperlin, D. J. (2011). A Multiwell Platform for Studying Stiffness-Dependent Cell Biology. PLoS One 6(5): e19929.
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Millar‐Haskell, C. S., Sperduto, J. L., Slater, J. H., Thorpe, C. and Gleghorn, J. P. (2022a). Secretion of the disulphide bond generating catalyst QSOX1 from pancreatic tumour cells into the extracellular matrix: Association with extracellular vesicles and matrix proteins. J. Extracell. Biol. 1(7): e48.
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13CO2-labelling and Sampling in Algae for Flux Analysis of Photosynthetic and Central Carbon Metabolism
OG Or Geffen *
DA David Achaintre *
HT Haim Treves
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4808 Views: 498
Reviewed by: John P Phelan Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Nature Plants Dec 2021
Abstract
The flux in photosynthesis can be studied by performing 13CO2 pulse labelling and analysing the temporal labelling kinetics of metabolic intermediates using gas or liquid chromatography linked to mass spectrometry. Metabolic flux analysis (MFA) is the primary approach for analysing metabolic network function and quantifying intracellular metabolic fluxes. Different MFA approaches differ based on the metabolic state (steady vs. non-steady state) and the use of stable isotope tracers. The main methodology used to investigate metabolic systems is metabolite steady state associated with stable isotope labelling experiments. Specifically, in biological systems like photoautotrophic organisms, isotopic non-stationary 13C metabolic flux analysis at metabolic steady state with transient isotopic labelling (13C-INST-MFA) is required. The common requirement for metabolic steady state, alongside its very short half-timed reactions, complicates robust MFA of photosynthetic metabolism. While custom gas chambers design has addressed these challenges in various model plants, no similar tools were developed for liquid photosynthetic cultures (e.g., algae, cyanobacteria), where diffusion and equilibration of inorganic carbon species in the medium entails a new dimension of complexity. Recently, a novel tailor-made microfluidics labelling system has been introduced, supplying short 13CO2 pulses at steady state, and resolving fluxes across most photosynthetic metabolic pathways in algae. The system involves injecting algal cultures and medium containing pre-equilibrated inorganic 13C into a microfluidic mixer, followed by rapid metabolic quenching, enabling precise seconds-level label pulses. This was complemented by a 13CO2-bubbling-based open labelling system (photobioreactor), allowing long pulses (minutes–hours) required for investigating fluxes into central C metabolism and major products. This combined labelling procedure provides a comprehensive fluxome cover for most algal photosynthetic and central C metabolism pathways, thus allowing comparative flux analyses across algae and plants.
Keywords: Photosynthesis Algae Isotope labelling Non-stationary 13C-metabolic flux analysis Metabolomics
Background
Over the last two decades, numerous analytical and computational approaches have emerged for deciphering metabolic networks. Understanding metabolic pathways and their regulation is crucial for metabolic engineering, biotechnology, or pharmacology, which can be explored through metabolic flux analysis (MFA) studies. Metabolite levels do not provide reliable information about pathway flux, due to its complex relationship with protein activity and substrate level (Blank and Sauer, 2004). Major progress has been made through the use of isotopic labelling, allowing a better and more accurate assessment of the concentration of metabolites by NMR, gas chromatography (GC), or liquid chromatography (LC) mass spectrometry (MS) and combinations thereof (Niedenführ et al., 2015). Various tools have been developed alongside computational and mathematical modelling methods (e.g., INCA, METRAN, SUMOFLUX), aiming to fit a theoretical model to estimate metabolic fluxes (Heise et al., 2015).
Isotopic non-stationary 13C metabolic flux analysis (13C-INST-MFA) offers a robust framework for flux estimation in single carbon substrate–based systems (e.g., autotrophic, methanotrophic) (Jazmin and Young, 2013). Aiming to resolve differential balance equations for the time-dependent labelling of intermediate metabolites, 13C-INST-MFA is ideally suited to slowly labelled systems (e.g., due to large intermediate pools or pathway bottlenecks). Thus, as the entry point for the single-carbon substrate system of most photoautotrophs, photosynthetic metabolism represents a unique challenge for 13C-INST-MFA, due to its relatively small intermediate pool size and, consequently, very rapid turnover rates (Szecowka et al., 2013; Heise et al., 2014; Abernathy et al., 2017; Allen and Young, 2020). Despite this challenge, this approach has been widely used in the past decade to estimate fluxes in photoautotrophic systems, ranging from cyanobacteria to land plants (Ma et al., 2014; Adebiyi et al., 2015; Cheah and Young, 2018; Wieloch, 2021; Xu et al., 2022).
Supported by the development of targeted analytical methodology (Arrivault et al., 2009 and 2015) and tailored labelling setups (Szecowka et al., 2013; Arrivault et al., 2017), these studies provided robust quantitative data on fluxes through central C metabolism and photosynthesis. The pioneering study of Szecowka and colleagues (Szecowka et al., 2013) has obtained the fluxes in canonical pathways of photosynthetic carbon metabolism in Arabidopsis rosette based on a combined 13CO2-labelling/GC- and LC-tandem MS/MS approach. Similar achievements have been made in Arabidopsis (under different conditions) (Ma et al., 2014), maize (Arrivault et al., 2017), tobacco (Chu et al., 2022), and Camelina sativa (Xu et al., 2021), but only partly in algae (Xiong et al., 2010; Wu et al., 2015) due to the experimentally challenging supply of rapidly labelled inorganic carbon pulses.
Using a tailor-made microfluidics labelling system to supply 13CO2 at steady state, we investigated in vivo labelling kinetics in intermediates of the Calvin Benson cycle and sugar, starch, organic acid, and amino acid synthesis pathways, and flux into protein and lipids, in several model and non-model green algae (Treves et al., 2022). Our system allows sampling at the 0–40 s pulse timescale in a highly reproducible manner, yielding high quality data. Furthermore, when combined with traditional labelling setups such as open systems with bubbling of 13CO2 for longer pulse times, our protocol, which is applicable to other unicellular photoautotrophic systems, largely improves the precision of flux estimates through the application of 13C-INST-MFA.
Materials and reagents
Inorganic 13C (Sigma-Aldrich, catalog number: 364592)
N2 gas (Oxygen and Argon Works Ltd.)
O2 gas (Oxygen and Argon Works Ltd.)
12CO2 gas (Oxygen and Argon Works Ltd.)
13CO2 (isotopic purity 99-atom percentage) gas (Sigma-Aldrich, catalog number: 364592-10L-EU)
Lime soda (Merck Millipore, catalog number: 1068395000)
Chlamydomonas reinhardtii strain CC-124 (UTEX Culture Collection of Algae)
Chlorella sorokiniana UTEX 1663 (UTEX Culture Collection of Algae)
Chlorella ohadii
HEPES (Sigma-Aldrich, catalog number: 54457)
Sodium hydroxide (NaOH) (Sigma-Aldrich, catalog number: 655104)
Calcium chloride (CaCl2) (Sigma-Aldrich, catalog number: C4901)
Ammonium chloride (NH4Cl) (Sigma-Aldrich, catalog number: 213330)
Magnesium sulphate heptahydrate (MgSO4·7H2O) (Sigma-Aldrich, catalog number: 230391)
Dipotassium phosphate (K2HPO4) (Sigma-Aldrich, catalog number: P3786)
Monopotassium phosphate (KH2PO4) (Sigma-Aldrich, catalog number: PX1562)
Ethylenediaminetetraacetic acid (EDTA) (Sigma-Aldrich, catalog number: E9884)
Boric acid (BO3H3) (Sigma-Aldrich, catalog number: B0394)
Zinc sulphate heptahydrate (ZnSO4·7H2O) (Sigma-Aldrich, catalog number: 221376)
Manganese(II) chloride tetrahydrate (MnCl2·4H2O) (Sigma-Aldrich, catalog number: 221279)
Iron(II) sulphate heptahydrate (FeSO4·7H2O) (Sigma-Aldrich, catalog number: 215422)
Cobalt(II) chloride hexahydrate (CoCl2·6H2O) (Sigma-Aldrich, catalog number: 255599)
Copper sulphate pentahydrate (CuSO4·5H2O) (Sigma-Aldrich, catalog number: 209198)
Ammonium molybdate tetrahydrate (Mn7O24(NH4)6·4H2) (Sigma-Aldrich, catalog number: 09878)
Methanol (Sigma-Aldrich, catalog number: 34860)
Chloroform (Sigma-Aldrich, catalog number: 650498)
tert-Butyl methyl ether (Sigma-Aldrich, catalog number: 650560)
Liquid nitrogen (N2)
Dry ice
Ethanol, technical
HP (HEPES-phosphate) medium (pH 7.2) (see Recipes)
Nutrient stock (see Recipes)
Phosphate buffer 1,000× (see Recipes)
Hutner’s solution (see Recipes)
MTBE (methyl tert-butyl ether) solution (see Recipes)
Equipment
Gas (mass-flow) controllers calibrated for N2, O2, and CO2 (Brooks Instruments, model: 5850S) and smart control software (Brooks Instruments)
20 mL syringes
Syringe pump (KF Technology, model: NE-1600 Multi-Channel Syringe Pump)
Transparent tubing (Vici Jour, catalog number: R-T-4007-M10)
Luer adapters (Upchurch Scientific, Union Polypropylene, UPP series)
Flangeless ferrule (Upchurch Scientific, catalog number: UPP-200N)
Union assembly (Upchurch Scientific, catalog number: UPP-360)
Luer adapter (Upchurch Scientific, catalog number: UPP-658)
Gas washing bottle (Robu Glasfilter-Geräte, catalog number: 41101)
Flat glass photobioreactors (PSI, model: FMT-150)
Cool-white LED array (PSI, model: 3500-D)
Centrifuge (Eppendorf, model: 5418 R)
LC-MS (Thermo Fisher Scientific, model: TSQ Quantum Ultra with Excalibur 2.07 SP1 and TSQ Quantum 1.4 software)
GC-MS (LECO Instruments GmbH, model: Pegasus III TOF-MS)
Gas chromatograph (Agilent Technologies, model: 6890N24)
Humidifier:
Laboratory bottles (DURAN, Sigma-Aldrich, catalog number: Z305197)
Head with filter-disc for GL 45 (ROBU Glasfilter-Geraete GmbH, catalog number: ISO 4793-80)
50 mL centrifuge tubes (Sarstedt, catalog number: 62.547.004)
Lyophilizer (Martin Christ, model: Alpha 2-4 LSCbasic)
Microfluidic mixer (Manufactured based on original design from Sivashankar et al., 2016)
Light meter (LI-COR, model: LI-250A)
Air pump (Sera precision, model: air 275 R plus, catalog number: 08814)
Autoclave (Witeg, model: WAC-60 230 V)
Filter 0.22 μm (Sigma-Aldrich, catalog number: SLMP025SS)
Water bath (GFL, model: 1002)
Oxygen sensor (Mettler Toledo, model: InPro 6800)
pH sensor (Mettler Toledo, model: InPro 3250)
Upright microscope (Nikon, model: Eclipse E200)
SpeedVac concentrator (Thermo Fisher Scientific, model: SPD210)
Gas analyser (LI-COR, model: LI850)
Procedure
Algal cultures
Grow algal cultures in flat glass bioreactor vessels, as specified in the instructions below. Further details associated with this technique are described in Treves et al. (2017).
Prepare HP medium (see Recipes) for the bioreactors.
Autoclave (121 °C, saturated steam, 30 min) each bioreactor cuvette (see https://photo-bio-reactors.com/products/photobioreactors/#details) with 1 L of HP medium and both oxygen and pH sensors connected.
On a sterile bench, connect 0.22 μm filters to all air inlets/outlets of the system and add 10 mL of axenic inoculum into the bioreactor medium, to reach an initial cell density of OD735nm = 0.02. The optical density of the culture is measured by the bioreactor following activation (see https://photo-bio-reactors.com/products/photobioreactors/#details).
Initiate the bioreactors at the optimal or target temperature defined for each strain and irradiate the cells at light regimes (in μmol photons m-2·s-1) optimised and designed for the study goals. For reliable estimation of in vivo fluxes, illumination levels used for labelling should be set to reflect the penetrating light levels within the bioreactor, as measured using an integrating sphere inside the running culture in the bioreactors.
Supply air for bubbling using an air-pump at approximately 1 L/min and monitor pH, dissolved oxygen concentration, and optical densities (OD) at 680 and 735 nm every 1–5 min, through integral bioreactor sensors (https://photo-bio-reactors.com/products/photobioreactors/#details).
Perform independent biological replicates of three separate bioreactor runs for each alga or condition.
Determine the weight of organic material within the samples through the measurement of ash-free dry weight per volume for each culture, as previously described in Klassen et al. (2015).
Note: Assess and validate the axenicity of the cultures with light microscopy and Luria-Bertani plating/incubation.
13CO2-labelling and sampling procedures
Two setups are used to provide inorganic 13C to algal liquid cultures (Figure 1).
Figure 1. Simplified labelling procedure scheme. Flowchart presenting the main steps of rapid (left) and slow (right) labelling procedures.
Very short pulses (up to 40 s) and rapid labelling (time points 5, 10, 20, and 40 s):
Fill a glass bottle with 400 mL of fresh HP medium (pH 7.2).
Place the glass bottle in a temperature-controlled water bath kept at the growth temperature of each algal culture, preferably in a separate room to avoid 13CO2 contamination in the bioreactors’ running cultures prior to mixing.
Bubble with the synthetic gas mixture (78% N2, 21% O2, and 400 ppm 13CO2) for 30 min to reach < 1% residual 12CO2 levels (see Figure 2B). Run exhaust flow through lime soda pellets via gas washing bottle to avoid 13CO2 cross contamination of running cultures.
Figure 2. Kinetics of 13CO2 supply via bubbling (A) through algal cultures or (B) HP medium or double-distilled water (DDW). Monitoring of 12CO2 levels at the pulse, as measured on the exhaust of algal cultures/medium/DDW using a gas analyser. (A) Different algal cultures, grown to a density corresponding to T0 for each alga and condition, were bubbled with natural air. Thereafter, the natural air was rapidly changed (red arrow) to 13C-based synthetic labelled air mixture (see Procedure). Reciprocally, approximately 20 min after the first change, bubbling is rapidly replaced again for natural air (green arrow).
Wash a 20 mL syringe three times and fill it with the bubbled solution (keep contact with air to a minimum).
Load the syringe to a syringe pump.
Connect the syringe with transparent tubing by screwing the luer to the screw tip of the syringe.
Turn on the light source above the syringe pump and bioreactor and adjust to illumination levels measured in step A4.
Connect transparent tubing (with length adjusted to pulse time, see Notes) between the mixer and the cooled 50 mL tubes (see Figure 3A).
Figure 3. Rapid algal labelling system scheme. (A) Fresh medium bubbled with 13CO2-based synthetic air mixture (see Procedure) is mixed with fresh algal sample withdrawn from the bioreactor under controlled LED illumination. (B) Solutions are rapidly (< 1 s) mixed in a tailor-made transparent chip and injected through transparent tubing into 70% methanol solution kept at -70 °C. Pulse length (5–40 s) is controlled via tubing length.
Withdraw a fresh algal sample into a 20 mL syringe using a transparent tube under the light. Load the syringe to the pump while avoiding any shading throughout the process.
Connect the syringe to transparent tubing by screwing the luer to the screw tip of the syringe (Figure 3A).
Operate the pump manually to push the liquids in both syringes and remove residual air from the tubes to the microfluidic mixer. Program the syringe pump to the required volume in each of the four tube lengths (see Section B, step 1h, above). Push the cultures and fresh media through the transparent mixer and tubing and collect the 1:1 mixture to a 70% methanol solution in 50 mL tubes cooled to -70 °C. Tubes are cooled in an ethanol bath precooled with dry ice (see scheme in Figure 3A). Inject the mixed sample into the cooled 70% methanol to reach a 1:2 ratio (e.g., 15 mL of sample into 30 mL of 70% methanol) (see Notes).
Keep the cooled 50 mL tubes at -70 °C until centrifugation (3,200× g, 3 min, -9 °C).
Discard the supernatant (see Notes) and freeze the pellet immediately in liquid nitrogen.
Repeat the same procedure for the non-labelled samples for each pulse time series replica by mixing the same algal culture with fresh HP medium pre-bubbled with ambient air at the same temperature as in Section B, step 1b.
Following rapid freezing, samples may be stored in a -80 °C freezer until extraction.
Resuspend the frozen pellets in ice-cold methanol/chloroform (5:1, v/v).
Perform four freeze-thaw cycles (see Notes) of resuspended cells and perform metabolite extraction following the methanol/chloroform procedure (see Mettler et al., 2014).
Notes:
Synthetic gas mixture is mixed using mass flow controllers as described previously (Szecowka et al., 2013).
Light is provided by an upper-positioned cool-white LED array through the entire route from sampling at the bioreactor to the quenching tube, with intensities corresponding to the penetrating light for each culture and avoiding shading.
Pulse duration is implemented by fitting the length of the transparent tubing downstream to the mixer to spray the culture/13CO2 solution mixture into the quenching tube at desired timings.
Metabolites level within the supernatant (see section B, step 1m) should be assessed by LC-MS/MS and should ideally be up to approximately 1% that of equivalent pellets.
Depending on the cell wall rigidity of the algal strains, the number of freeze-thaw cycles should be adjusted to the point where all chlorophyll migrates to the chloroform phase.
Microfluidic mixer (Figure 3B) may be produced by 3D printing, according to the detailed design provided in Sivashankar et al., 2016.
Long 13C incubation times (15–300 min) and slow labelling:
Introduce inorganic 13C via direct bubbling of the cultures with the gas mixture used for rapid labelling.
Bubble algal cultures with 13CO2 gas mixture (approximately 1 L/min) through a humidifier as done for the rapid labelling. Switch the gas mixture from natural air to the same synthetic 13C-labelled air mixture (T0). (Residual 12CO2 in the cultures should be below 2% within a few minutes of pulse. See Figure 2A). Run exhaust flow from labelled culture through lime soda pellets via gas washing bottle to avoid 13CO2 cross contamination of other running cultures.
Collect samples into 50 mL Falcon tubes before switching (T0) and at 15, 30, 60, 120, 180, and 300 min of bubbling with 13C synthetic gas mixture.
Centrifuge immediately (3,200× g, 3 min, -9 °C), remove supernatant, and freeze pellets in liquid nitrogen.
Resuspend frozen pellets in precooled (-20 °C) MTBE solution.
Perform four freeze-thaw cycles of resuspended pellets, followed by a two-phase MTBE extraction to analyse metabolites, protein, and starch, as described in Jüppner et al. (2017).
i. Dry the upper MTBE phase, containing lipids, in a SpeedVac concentrator and store at -80 °C to analyse lipid content and 13C labelling.
ii. Dry the lower phase, containing the polar and semipolar metabolites, in a SpeedVac concentrator and store at -80 °C for metabolite profiling.
iii. Store the solid pellets containing the precipitated protein and starch at -20 °C for further analysis of labelling into starch and protein as previously described (Bradford, 1976; Arrivault et al., 2009; Ishihara et al., 2015; Fernandez et al., 2017).
Notes:
Slow labelling is required, since longer incubation times would lead to Ci depletion in the closed microfluidic system.
Measured residual 12CO2 in the cultures reaches below 2% within a few minutes of pulse (Figure 2A) and H12CO3- is rapidly equilibrated by the cells as previously reported for low CO2-acclimated algae (Tchernov et al., 2003) and demonstrated here by the absence of a slow 12CO2 decay phase in the cultures (Figure 2A and 3B).
In step 2c, centrifugation is necessary as some metabolites can accumulate in the medium over time, thus leading to labelling patterns in the complete suspension not reflecting that in the cells.
Data analysis
Data analysis, peak annotation, and quality control (QC) procedures are thoroughly described in Arrivault et al. (2009) and Ma et al. (2014). Briefly, data analysis includes metabolite peak annotation and correction, followed by correction of raw data for the abundance of stable isotopes using the Corrector software tool (https://www.mpimp-golm.mpg.de/19405/Corrector_package_V1_91.zip), analysis of calibration curves for each metabolite and correction for signal decay along each run using periodic injection of standard mix solution; relative isotopomer abundance (m + n) for each metabolite is calculated as in Szecowka et al. (2013).
Recipes
All recipes regarding the preparation of the HP medium and associated stocks are available in Harris (1989).
HP medium, pH 7.2 (for 1 L)
HEPES 1.19 g
Nutrient stock 25 mL
Phosphate buffer 1,000× 1 mL
Hutner’s solution 1 mL
Make up to 1 L with DDW, set pH at 7.2 using NaOH, and autoclave.
Store in a glass bottle at room temperature (solution is not light sensitive).
Note: Following autoclave, add 25 mL of CaCl2 18 mM.
Nutrient stock (for 1 L)
NH4Cl 8.57 g
MgSO4·7H2O 4 g
Make up to 1 L with DDW and autoclave.
Store in a glass bottle at room temperature (solution is not light sensitive).
Phosphate buffer 1,000× (pH = 7) (for 1 L)
K2HPO4 106 g
KH2PO4 53 g
Make up to 1 L with DDW and autoclave.
Store in a glass bottle at room temperature (solution is not light sensitive).
Hutner’s solution, pH = 6.5–6.8 (for 1 L)
EDTA 50 g
Bo3H3 11.4 g
ZnSO4·7H2O 22 g
MnCl2·4H2O 5.06 g
FeSO4·7H2O 4.99 g
CoCl2·6H2O 1.61 g
CuSO4·5H2O 1.57 g
Mn7O24(NH4)·4H2O 1.1 g
Store in a glass bottle at 4 °C and wrap the glass bottle in aluminium foil (solution is light sensitive).
MTBE solution
Methanol/methyl tert‐butyl‐ether (1/3).
Store in a glass bottle at 4 °C (solution is not light sensitive).
Acknowledgments
We thank Dr. Stephanie Arrivault from MPI-MP for providing support for the optimization of algal harvest and extraction procedures through iterative LC-MS/MS measurements. We thank Prof. Dr. Mark Stitt for constructive discussions regarding experimental setup and controls design. We thank Dr. Hirofumi Ishihara for his support of integrating the labelling mixer with existing 13C-labelling gas mixer setup. This work was supported by grant 1697/22 from the Israeli Science Foundation and by the Alon fund from the Israel Council of Higher Education. This protocol was adapted from Treves et al. (2022).
Competing interests
The authors declare no competing interests.
References
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The authors used this protocol in mBio Dec 2021
Abstract
Magnaporthe oryzae is a filamentous fungus responsible for the detrimental rice blast disease afflicting rice crops worldwide. For years, M. oryzae has served as an excellent model organism to study plant pathogen interactions due to its sequenced genome, its amenability to functional genetics, and its capacity to be tracked in laboratory settings. As such, techniques to genetically manipulate M. oryzae for gene deletion range from genome editing via CRISPR-Cas9 to gene replacement through homologous recombination. This protocol focuses on detailing how to perform gene replacement in the model organism, M. oryzae, through a split marker method. This technique relies on replacing the open reading frame of a gene of interest with a gene conferring resistance to a specific selectable chemical, disrupting the transcription of the gene of interest and generating a knockout mutant M. oryzae strain.
Key features
• Comprehensive overview of primer design, PEG-mediated protoplast transformation, and fungal DNA extraction for screening.
Graphical overview
Keywords: Magnaporthe oryzae Primer design Protoplast transformation Gene deletion Split marker Homologous recombination ILV1 Hygromycin DNA extraction
Background
The fungal plant pathogen Magnaporthe oryzae is the causal agent of rice blast disease. Rice blast afflicts all parts of rice, including leaves, necks, and panicles. Due to its severity, rice blast continues to annihilate rice crops worldwide every year (Fernandez and Orth, 2018; Asibi et al., 2019). As such, dissecting the intricacies of M. oryzae biology is vital to further our understanding of this devastating plant pathogen. As an organism that can be cultivated in the lab as well as genetically manipulated, M. oryzae serves as a model organism to elucidate the molecular basis of rice blast disease and plant pathogen interactions. M. oryzae genetic transformations enable researchers to characterize protein functionality through gene deletions or visualize protein localization through fluorescent tagging. Through editing of the M. oryzae genome, methods to prevent the spread of this disease may be developed. For years, the split marker method has been a key approach for gene deletions in M. oryzae (Sweigard et al., 1995; Fernandez et al., 2014a, 2014b, and 2021). Relying on the homologous recombination that occurs naturally in M. oryzae, the split marker method entails generating flanking regions of a gene of interest through two rounds of PCR and fusing each of them to half of a selectable marker, without requiring any subcloning. This method relies on three homologous recombination events occurring, with two events fusing the flank regions to each half of the selectable marker, and one fusing the selectable marker cassette together. Combining these flanks with M. oryzae protoplasts will generate mutants that have replaced the gene of interest with the selectable marker. In more recent times, various methods for gene editing have been reported. For instance, CRISPR-Cas9-based approaches have been applied for gene editing in M. oryzae (Foster et al., 2018). However, the split marker method has been widely utilized throughout the years due to its convenient, effective, and inexpensive features that facilitate the generation of mutant strains in M. oryzae.
Materials and reagents
Biological materials
Magnaporthe oryzae entry strain of choice, such as Guy11 as used in this protocol (Guy11 strain can be obtained from the Fungal Genetics Stock Center).
Reagents
Yeast nitrogen base w/o amino acids (BD Difco, catalog number: 233520)
Ammonium nitrate 98%, ACS reagent (Thermo Fisher, catalog number: 423350250)
L-Asparagine, 99% (Thermo Fisher, catalog number: B21473.36)
Dextrose/D-(+)-glucose anhydrous ACS reagent grade (MP Biomedicals, catalog number: 152527)
Sucrose (Thermo Fisher, catalog number: J65148.36)
Agar (Sigma-Aldrich, catalog number: A1296-100G)
Peptone (Thermo Fisher, catalog number: 211677)
Yeast extract (Thermo Fisher, catalog number: 212750)
Casamino acids (Thermo Fisher, catalog number: 228820)
NaOH (Sigma-Aldrich, catalog number: S5881-500G)
Na2HPO4 (Sigma-Aldrich, catalog number: S9763-100G)
100% ethanol (Thermo Fisher, catalog number: T038181000)
100% isopropanol (Thermo Fisher, catalog number: T036181000)
RNase (Sigma-Aldrich, catalog number: 10109142001)
Sodium nitrate (Sigma-Aldrich, catalog number: 221341-500G)
Potassium chloride (Thermo Fisher, catalog number: A11662.0B)
Magnesium sulfate heptahydrate (Thermo Fisher, catalog number: A14491.0B)
Potassium phosphate, dibasic (Sigma-Aldrich, catalog number: P3786-100G)
Zinc sulphate heptahydrate (Sigma-Aldrich, catalog number: Z0635-100G)
Boric acid (VWR, catalog number: BDH9222-500G)
Manganese chloride (Thermo Fisher, catalog number: 271410250)
Iron sulfate heptahydrate, ACS, 99+% (Alta Aesar, catalog number: 14498-30)
Cobalt chloride hexahydrate (Thermo Fisher, catalog number: 011344.30)
Copper sulfate pentahydrate (Thermo Fisher, catalog number: 197722500)
Sodium molybdate (Thermo Fisher, catalog number: 206371000)
Na4EDTA (Thermo Fisher, catalog number: J15700.A1)
Biotin (Sigma-Aldrich, catalog number: B4501-100MG)
Pyridoxin (Sigma-Aldrich, catalog number: P5669-25G)
Thiamine (Thermo Fisher, catalog number: 148991000)
Riboflavin (Sigma-Aldrich, catalog number: 47861)
PABA (Sigma-Aldrich, catalog number: A9878-5G)
Nicotinic acid (Sigma-Aldrich, catalog number: N0761-100G)
1 M Tris-HCl, pH 8.0 (Thermo Fisher, catalog number: 15568025)
0.5 M EDTA, pH 8.0 (Sigma-Aldrich, catalog number: 324506-100ML)
Sorbitol (Thermo Fisher, catalog number: 036404.A3)
Lysing enzymes from Trichoderma (Sigma-Aldrich, catalog number: L1412)
1 M Tris-HCl, pH 7.5 (Thermo Fisher, catalog number: 15567027)
1 M CaCl2 (Thermo Fisher, catalog number: J63122.AD)
PEG 4000 (Polysciences, catalog number: 16861-250)
Chlorimuron ethyl (Sulfonylurea) (Thermo Fisher, catalog number: J66605.03)
Hygromycin B (Thermo Fisher, catalog number: 10687010)
Penicillin/Streptomycin/Neomycin (PSN) antibiotic 100× solution (Thermo Fisher, catalog number: 15640055)
Q5 DNA polymerase (NEB, catalog number: M0491S)
PowerPol 2× PCR polymerase mix (Abclonal, catalog number: RK20718)
Solutions
20× (+) NO3 salts (see Recipes)
BDCM bottom media (see Recipes)
BDCM top media (see Recipes)
Liquid CM (see Recipes)
CM bottom media (see Recipes)
CM top media (see Recipes)
Trace elements (see Recipes)
Vitamin solution (see Recipes)
DNA extraction buffer (see Recipes)
70% ethanol (see Recipes)
Osmotic (OM) buffer (see Recipes)
ST buffer (see Recipes)
STC buffer (see Recipes)
PTC buffer (see Recipes)
1 M NaPO4 (pH 5.8) (see Recipes)
Recipes
20× (+) NO3 salts (1 L)
Reagent Quantity
Sodium nitrate 120.0 g
Potassium chloride 10.4 g
Magnesium sulfate 10.4 g
Potassium phosphate 30.4 g
H2O Up to 1 L
Autoclave at 121 °C for 25 min, cool to room temperature, then store at 4 °C.
BDCM bottom (1 L)
Reagent Quantity
Yeast nitrogen base w/o amino acids 1.7 g
Ammonium nitrate 2.0 g
L-Asparagine 1.0 g
Dextrose 10.0 g
Sucrose 273.83 g
H2O Up to 1 L
Agar 1.5 g/100 mL
Adjust pH to 6.0 with 1 M Na2HPO4 and autoclave at 121 °C for 25 min.
BDCM top (1 L)
Reagent Quantity
Yeast nitrogen base w/o amino acids 1.7 g
Ammonium nitrate 2.0 g
L-Asparagine 1.0 g
Dextrose 10.0 g
H2O Up to 1 L
Agar 1.0 g/100 mL
Adjust pH to 6.0 with 1 M Na2HPO4 and autoclave at 121 °C for 25 min.
Liquid CM (1 L)
Reagent Quantity
20× (+) NO3 salts 50.0 mL
Trace elements 1.0 mL
Vitamin solution 1.0 mL
Peptone 2.0 g
Yeast extract 1.0 g
Casamino acids 1.0 g
Dextrose 10.0 g
H2O Up to 1 L
Adjust pH to 6.5 with 1 M NaOH and autoclave at 121 °C for 25 min.
CM bottom (1 L)
Reagent Quantity
20× (+) NO3 salts 50.0 mL
Trace elements 1.0 mL
Vitamin solution 1.0 mL
Peptone 2.0 g
Yeast extract 1.0 g
Casamino acids 1.0 g
Dextrose 10.0 g
Sucrose 273.83 g
H2O Up to 1 L
Agar 1.5 g/100 mL
Adjust pH to 6.5 with 1 M NaOH and autoclave at 121 °C for 25 min.
CM top (1 L)
Reagent Quantity
20× (+) NO3 salts 50.0 mL
Trace elements 1.0 mL
Vitamin solution 1.0 mL
Peptone 2.0 g
Yeast extract 1.0 g
Casamino acids 1.0 g
Dextrose 10.0 g
H2O Up to 1 L
Agar 1.0 g/100 mL
Adjust pH to 6.5 with 1 M NaOH and autoclave at 121 °C for 25 min.
Trace elements (100 mL)
Reagent Quantity
Sterile, autoclaved H2O 80 mL
Zinc sulfate 2.2 g
Boric acid 1.1 g
Manganese chloride 0.5 g
Iron sulfate 0.5 g
Cobalt chloride hexahydrate 0.17 g
Copper sulfate pentahydrate 0.16 g
Sodium molybdate 0.15 g
Na4EDTA 5.0 g
Add all reagents in order. Using a hot plate, stir the solution until it comes to a boil. Cool to 60 °C and adjust the pH to 6.5 with 1 M KOH. Cool to room temperature. Adjust volume to 100 mL with ddH2O. Place in autoclaved bottles. Store at 4 °C. The solution should be a deep violet color before use.
Vitamin solution (100 mL)
Reagent Quantity
Biotin 0.01 g
Pyridoxin 0.01 g
Thiamine 0.01 g
Riboflavin 0.01 g
PABA (p-aminobenzoic acid) 0.01 g
Nicotinic acid 0.01 g
Sterile, autoclaved H2O Up to 100 mL
Place in autoclaved bottles. Store in the dark at 4 °C.
DNA extraction buffer (100 mL)
Reagent Quantity
Potassium chloride 7.45 g
1 M Tris-HCl (pH 8.0) 10 mL
0.5 M EDTA (pH 8.0) 2 mL
H2O Up to 100 mL
Filter sterilize for long-term storage.
70% ethanol (1 L)
Reagent Final concentration Quantity
Ethanol (absolute) 70% 700 mL
ddH2O n/a 300 mL
Total n/a 1000 mL
Osmotic (OM) buffer (100 mL)
Reagent Quantity
Magnesium sulfate heptahydrate 29.0 g
1 M NaPO4 (pH 5.8) 1 mL
Lysing enzymes from Trichoderma 0.75 g
H2O Up to 100 mL
Adjust pH to 5.5 with 1 M Na2HPO4. Filter sterilize using a 0.45 μm syringe filter into 50 mL centrifuge tubes. Two 50 mL tubes should have 40 mL of OM buffer, while a third 50 mL tube should contain the remaining 20 mL. OM should be prepared the day before using and placed at 4 °C overnight.
ST buffer (500 mL)
Reagent Quantity
Sorbitol 54.66 g
1 M Tris-HCl (pH 7.0) 50 mL
H2O Up to 500 mL
Autoclave for 25 min at 121 °C and store at 4 °C.
STC buffer (500 mL)
Reagent Quantity
Sorbitol 109.32 g
1 M Tris-HCl (pH 7.5) 5.0 mL
1 M CaCl2 5.0 mL
H2O Up to 500 mL
Autoclave for 25 min at 121 °C and store at 4 °C.
PTC buffer (20 mL)
Reagent Quantity
PEG 4000 8.0 g
Sorbitol 3.64 g
1 M Tris-HCl (pH 7.5) 200 μL
1 M CaCl2 200 μL
H2O Up to 20 mL
Autoclave for 25 min at 121 °C and store at room temperature.
1 M NaPO4 (pH 5.8)
Reagent Amount
1 M NaH2PO4 92.1 mL
H2O Up to 100 mL
Adjust pH to 5.8 with 1 M Na2HPO4. Bring volume up to 100 mL and autoclave for 25 min at 121 °C.
Laboratory supplies
Pipettes 1,000 (Sartorius, catalog number: 14532053), 10 (Sartorius, catalog number: 11014473), and 200 μL (Sartorius, catalog number: 10195873)
Serological pipette (Scilogex SCI-Pet, Amazon)
25 mL serological pipette (Genesee Scientific, catalog number: 12-106)
Hemocytometer (Fisher Scientific, catalog number: 02-671-5)
Petri dishes (Fisher Scientific, catalog number: FB0875713)
Scalpel (Feather, catalog number: 02058370)
Miracloth (Calbiochem, catalog number: B36658)
Oakridge tube (Thermo Fisher, catalog number: 3119-0050)
50 mL conical tubes (Corning, catalog number: 430291)
500 mL flasks (Pyrex, catalog number: 4980)
Sterile paper towels (Scott multifold paper towels, Amazon)
Sterile funnel (Thermo Fisher, catalog number: 4252-0065PK)
100 mL bottles (Kimax, catalog number: 14397)
Dark trays (Mr. Pen plant trays, Amazon)
Aluminum foil (Reynolds Wrap, Amazon)
Incubator (Precision, catalog number: 31483)
1.5 mL microcentrifuge tubes (Fisher Scientific, catalog number: 05-408-130)
0.45 μm syringe filters (Fisher Scientific, catalog number: 09-720-514)
Monarch PCR purification kit (NEB, catalog number: T1030S)
Equipment
Incubator (Percival, model: I36VL)
Centrifuge (Eppendorf, model: Centrifuge 5810 R with S-4-104 rotor)
Water bath (Thermo Fisher, model: 180 Water Bath Series)
Two-speed Waring laboratory blender (Fisher Scientific, catalog number: 7010S)
Eberbach container for Waring blenders (Fisher Scientific, catalog number: 14-509-25)
Incu-shaker (Benchmark, model: 10LR)
Thermal cycler (Bio-Rad, model: C1000 Touch)
Vacufuge plus (Eppendorf, model: 022820109)
Centrifuge for Eppendorf tubes (Eppendorf, model: Centrifuge 5418)
Homogenizer (Amazon, SP Bel-Art ProCulture F65100-0000)
Hot plate and stirrer (Thermo Fisher, catalog number: SP88854100)
Procedure
Primer design and PCR reactions
Generate upstream and downstream regions outside of gene of interest’s open reading frame and selectable marker pieces.
Design primers that amplify 1 kb upstream of the beginning of your gene’s open reading frame (L1, L2; Figure 1).
Figure 1. Split marker primer design
Design primers that amplify 1 kb downstream following the end of your gene’s open reading frame (L3, L4).
Refer to Table 1 to see the primers to amplify the two pieces of the ILV1 gene that confers resistance to sulfonylurea (see Note 1).
Table 1. List of oligonucleotides used in the ILV1 transformation (see Note 2)
ID 5′----3′ sequences
ILV1 gene
ILV1 M13F CGCCAGGGTTTTCCCAGTCACGAC GTCGACGTGCCAACGCCACAG
IL Split R AAGCATGTGCAGTGCCTTC
LV1-M13R AGCGGATAACAATTTCACACAGGA TAGGTCGACGTGAGAGCATGC
LV1Split F GAGGCCGACGTCATAGGCATC
M13 sequences for the selectable marker
M13F CGCCAGGGGTTTTCCCAGTCACGAC
M13R AGCGGATAACAATTTCACACAGGA
M13 for the gene of interest
M13F TCCTGTGTGAAATTGTTATCCGCT
M13R GTCGTGACTGGGAAAACCCTGGCG
Amplify flanks using PCR.
Amplify the upstream and downstream region of your gene of interest using M. oryzae genomic DNA as the template.
Amplify the two pieces of your selectable marker using an ILV1 containing plasmid as the template.
Purify the amplified fragments using a PCR purification kit.
Fuse the upstream region of your gene of interest with the 5′ end of the selectable marker together using the 5′ products generated from the first round of PCR. Final DNA concentration should be approximately 5 μg (see Note 3).
Fuse the downstream region of your gene of interest with the 3′ end of the selectable marker together using the 3′ products generated from the first round of PCR. Final DNA concentration should be approximately 5 μg.
Purify the resulting fused fragments using a PCR purification kit.
Combine the fused fragment containing the upstream region of the gene and the 5′ end of the selectable marker along with the downstream region of the gene and 3′ end of the selectable marker in one tube.
Dry each tube until only ~10 μL of DNA are present using a vacuum concentrator.
Add 50 μL of STC buffer to each tube (see Note 4).
Protoplast generation
Grow M. oryzae mycelia on liquid complete medium (CM) (Figure 2).
Figure 2. Materials needed for generation of liquid CM mycelial culture
Plate fungal strain on three CM plates each containing three colonies. M. oryzae was grown at 24 °C with a 12 h photoperiod using a I36VL Percival incubator with a IncuWhite Linear LED lighting system, with an intensity of 120 μmol/m2/s (Figure 3).
Figure 3. CM plate containing three 5-day-old colonies
After five days, cut the growing circumference of mycelia from each colony using a sterile scalpel. (Video 1).
Video 1. Cutting growing circumference of mycelia. Using a blade, the growing edge of mycelia is cut from CM plates and placed in liquid CM to blend.
Combine the cut edges of mycelium in 250 mL of liquid CM in a sterile blender. Blend on high for 10 s, then on low for 10 s.
Transfer blended mycelium into a 500 mL flask and add 250 μL of antibiotic solution to the culture. The PSN antibiotic solution from Thermo Fisher is used at a 1× concentration.
Incubate culture at 24 °C with a 12 h photoperiod for approximately 48 h with shaking at 150 rpm (see Note 5).
After 24 h, add an additional 100 mL of liquid CM to the culture along with 100 μL of antibiotic solution and leave in shaking incubator at 200 rpm for the remainder of the 48 h.
Lyse mycelia to generate protoplast.
Filter mycelia from liquid culture using a sterile funnel and sterile Miracloth (Figures 4 and 5).
Figure 4. Materials needed to filter fungal culture
Figure 5. Filtered mycelia
Rinse filtered mycelia with sterile distilled water.
Using sterile paper towels, dry the filtered mycelia by squeezing the mycelia between paper towels to remove as much moisture as possible.
Divide dry mycelia into two 50 mL falcon tubes containing 40 mL of OM buffer.
Incubate tubes at 29 °C with shaking at 75 rpm for 3 h in a shaker incubator (see Note 6).
Harvest protoplast; do all of the following on ice.
Divide each falcon tube among two sterile clear polycarbonate Oakridge tubes.
Slowly add cold ST buffer drop by drop until each Oakridge tube is filled to the neck (see Note 7) (Video 2).
Video 2. Slow addition of ST buffer to Oakridge tube. Following mycelia lysis, the OM buffer and mycelia mixture is divided between two Oakridge tubes. Using a serological pipette, cold ST buffer is added slowly until the tube is filled to the neck.
Centrifuge tubes at 3,200× g for 25 min at 4 °C using a swinging bucket centrifuge with brakes disabled to ensure proper generation of protoplast layer.
Using a 1 mL pipette, slowly draw up protoplasts from each tube into new Oakridge tubes. Protoplasts from two Oakridge tubes can be combined into one new Oakridge tube (see Note 8) (Figure 6).
Figure 6. Protoplast appearance in Oakridge tube after centrifuging
Add STC buffer to the neck of each new Oakridge tube containing protoplasts.
Centrifuge tubes at 1,900× g for 10 min at 4 °C, with breaks half engaged.
A protoplast pellet should now be seen.
Washing protoplast.
Decant all STC buffer from each Oakridge tube.
Gently resuspend pellet with 5 mL of cold STC buffer.
Two resuspended pellets can be combined into one Oakridge tube.
Add STC buffer to the neck of each Oakridge tube.
Centrifuge tubes at 1,900× g for 10 min at 4 °C, with breaks half engaged.
Repeat steps B4a–B4e 2 additional times for three total protoplast washes.
Protoplast transformation.
Resuspend final protoplast pellet in 1 mL of STC buffer.
Using a hemocytometer, calculate protoplast concentration. A proper yield should be 2 × 108–5 × 108 protoplasts. Concentrate or dilute protoplasts with STC accordingly (Figure 7).
Figure 7. Protoplast after harvest. Visualization is at 40× magnification.
For each transformation, in a 1.5 mL tube, add 100 μL of protoplast to DNA (5 μg of each flank) that has been combined and resuspended in 50 μL of STC (see Note 9).
Incubate tubes at room temperature for 20 min (see Note 10).
*Please note that this is a time critical step. Avoid leaving the protoplasts in PTC for longer than the allotted time. Add 500 μL of PTC buffer to each tube. After adding PTC to the final tube, begin a 15 min timer (see Note 11).
Repeat the addition of 500 μL of PTC buffer to each tube until all tubes have a total of 1 mL of PTC.
Invert the tubes periodically during the 15 min incubation period.
Divide and label Petri plates. Four plates should be used for each strain (Figure 8).
Figure 8. Four Petri plates set aside per strain
Protoplast plating.
Add protoplasts to 100 mL of bottom media. *Please ensure that the bottom media is at approximately 45 °C as hotter temperatures will destroy the protoplast. Gently swirl to mix (see Notes 12 and 13).
Pour and divide media between four Petri dishes (see Note 14) (Video 3).
Video 3. Division of media. Following PTC incubation, the protoplasts are added to barely warm bottom media, swirled, and plated between four Petri dishes.
Once solidified, transfer plates to clean, dark trays and cover with aluminum foil.
Incubate plates at 24 °C for 16 h.
The next day, add top media with corresponding antibiotics to transformation plates (see Note 15).
Place plates back in dark trays and incubate for at least five days until colonies appear (Figure 9).
Figure 9. Colonies after fungal transformation
Screening fungal colonies
DNA extraction.
When colonies appear on the top layer of transformation plates, re-plate colonies in a grid on new media with corresponding antibiotics.
After 3–6 days, transfer a small amount of mycelia using a sterile toothpick or pipette tip from each colony into a sterile 1.7 mL tube and label each colony accordingly.
To each tube, add 500 μL of DNA extraction buffer.
Using an electronic homogenizer, pulverize mycelia for ~10 s (see Note 16).
Spin tubes at 2,100× g for 10 min at room temperature.
Following centrifugation, transfer the supernatant to a new tube containing 300 μL of 100% isopropanol. Mix tube (see Note 17).
Centrifuge tubes at 12,000× g for 10 min at room temperature.
Discard as much supernatant as possible.
Wash tube with 300 μL of 70% ethanol.
Centrifuge tubes at 12,000× g for 10 min at room temperature.
Discard as much supernatant as possible.
Air dry tubes by placing inverted tubes on a tissue for ~15 min.
After all ethanol has evaporated, add 30 μL of sterile water and 1 μL of RNase to each tube.
Incubate tube for 20 min at room temperature.
Incubate tube at 55 °C for 15 min to inactivate RNase.
Following incubation, add 150 μL of sterile water to each tube. Mix.
Use 1 μL of DNA extraction for screening.
PCR verification.
Using primers that fall outside the L1 and L4 region, run a PCR reaction for each of the potential mutants using the DNA extraction as a template. Use the wild-type Guy11 as a negative control (see Notes 18 and 19).
Run PCR amplification on a gel.
Verify band size for confirmation of positive transformation (see Note 20) (Figure 10).
Figure 10. Schematic of DNA gel depicting arbitrary amplifications following transformation. This arbitrary gene of interest is proposed to be 1 kb (although sizes will vary based on the gene of interest selected). A wild-type band would include the 1 kb upstream region, the 1 kb gene, and the 1 kb downstream region, yielding a size of 3 kb. A positive mutant would carry a higher band size at 4.8 kb, including the 1 kb upstream region, the 2.8 kb ILV1 gene, and the 1 kb downstream region. Please note that these sizes will vary based on the size difference between the gene of interest and the selectable marker.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
Fernandez et al. (2021). Role of two metacaspases in development and pathogenicity of the rice blast fungus Magnaporthe oryzae. mBio (Figure S1, panel c). Mutant strains for M. oryzae metacaspases were generated using this protocol.
General notes and troubleshooting
General notes
This protocol details the use of the ILV1 gene that confers resistance to sulfonylurea (gene size: 2.8 kb). However, other selectable markers can be used, such as hygromycin. For these PCR reactions, it is recommended to use a high-fidelity DNA polymerase, such as Q5 and Phusion.
Please note that certain primers require the addition of M13 primer sequences. The M13 sequences promote the fusion of fragments generated during the first round of PCR.
Typically, a PCR reaction volume of 100–200 μL is required to generate 5 μg of PCR product.
For transforming plasmids, 2–5 μg of plasmid should be linearized with a restriction enzyme before fungal transformation, PCR purified, dried to ~10 μL, and resuspended in 50 μL of STC buffer.
Please note that a healthy growth following this 48-h incubation period will produce a light brown media color. An overgrown culture will produce dark brown/black media.
During this incubation period, place bottles containing bottom medium in 55 °C water bath. Also, ensure centrifuge is at 4 °C.
Ensure that the addition of ST buffer to the Oakridge tubes is done drop by drop to ensure proper protoplast harvest.
The protoplasts are located at the hazy interface towards the middle of the tube. Ensure that when pipetting up protoplasts, the mycelium on the bottom is undisturbed.
A control should be included that has only 50 μL of STC buffer with no DNA and combined with 100 μL of protoplasts to check for protoplast integrity.
At this point, begin cooling the bottom medium bottles from 55 °C. By the end of the PTC buffer incubation, the media should be cool enough to touch with a gloved hand and should be around 45 °C.
Begin by pipetting the PTC buffer around the side of the tube until all PTC has been added. Slowly pipette the PTC buffer and protoplast mixture up twice. After mixing, gently invert the tubes twice.
Ensure that the media corresponds to the specific selection marker used. CM media is used for hygromycin selection; BDCM media is used for sulfonylurea selection; BASTA media is used for BASTA selection. Hygromycin is used as a 50 mg/mL stock concentration to a final concentration of 100 μg/μL. Sulfonylurea is used as a 100 mg/mL stock concentration to a final concentration of 50 μg/μL. BASTA is used as a 50 mg/mL stock concentration to a final concentration of 100 μg/μL.
Bottom media should be aliquoted in 100 mL volumes per transformation before autoclaving.
Ensure that each plate is 1/3 filled with media. Adding too much media to each plate will prevent properly adding top media the next day.
Bottom media should be used to plate protoplasts initially. Make sure that the top media is used for selection the following day.
The ground mycelia should not be left in the extraction buffer for a prolonged period of time. Additionally, if extracting more than one sample, the homogenizing pestle can be disinfected between samples by dipping in sterile water, followed by 100% ethanol.
Following this step, the samples can be left at -20 °C overnight and the DNA extraction can continue the next day.
Confirmation primers that fall outside the L1 and L4 region will ensure that the gene replacement has an endogenous incorporation of the selectable marker. Additionally, confirmation primers can be paired with internal split marker primers to yield smaller PCR fragments that can also be validated based on fragment size.
For this PCR screen, DNA polymerases such as DreamTaq from Thermo Fisher or PowerPol from ABclonal may be used for a more inexpensive reaction.
Amplicons of potential positive transformants can be confirmed using a sequencing service. Additionally, positive transformants can be further validated using Southern blot analysis.
Acknowledgments
This work was supported by the Early Career Grant and Undergraduate Research Internship program from UF/IFAS research. Graphical overview and Figures 1 and 10 were created with BioRender.com.
Competing interests
There are no conflicts of interest or competing interests.
References
Asibi, A. E., Chai, Q. and Coulter, J. A. (2019). Rice blast: A disease with implications for global food security. Agronomy 9(8): 451.
Fernandez, J., Lopez, V., Kinch, L., Pfeifer, M. A., Gray, H., Garcia, N., Grishin, N. V., Khang, C. H. and Orth, K. (2021). Role of two metacaspases in development and pathogenicity of the fungus Magnaporthe oryzae. mBio 12(1): e03471-20.
Fernandez, J., Marroquin-Guzman, M., Nandakumar, R., Shijo, S., Cornwell, K. M., Li, G. and Wilson, R. A. (2014a). Plant defence suppression is mediated by a fungal sirtuin during rice infection by Magnaporthe oryzae. Mol. Microbiol. 94(1): 70–88.
Fernandez, J., Marroquin-Guzman, M. and Wilson, R. A. (2014b). Evidence for a transketolase-mediated metabolic checkpoint governing biotrophic growth in rice cells by the blast fungus Magnaporthe oryzae. PLoS Pathog. 10(9): e1004354.
Fernandez, J. and Orth, K. (2018). Rise of a cereal killer: The biology of Magnaporthe oryzae biotrophic growth. Trends Microbiol. 26(7): 582–597.
Foster, A. J., Martin-Urdiroz, M., Yan, X., Wright, H. S., Soanes, D. M. and Talbot, N. J. (2018). CRISPR-Cas9 ribonucleoprotein-mediated co-editing and counterselection in the rice blast fungus. Sci. Rep. 8(1): e1038/s41598-018-32702-w.
Sweigard, J. A., Carroll, A. M., Kang, S., Farrall, L., Chumley, F. G. and Valent, B. (1995). Identification, cloning, and characterization of PWL2, a gene for host species specificity in the rice blast fungus. Plant Cell 7(8): 1221–1233.
Article Information
Copyright
© 2023 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 > Microbe-host interactions > Fungus
Molecular Biology > DNA > Transformation
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4,810 | https://bio-protocol.org/en/bpdetail?id=4810&type=0 | # Bio-Protocol Content
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An Optimized Protocol for Detecting Guard Cell–specific Gene Expression by in situ RT-PCR in Brassica rapa
YS Yingying Song *
XG Xinlei Guo
JW Jian Wu
JL Jianli Liang
RL Runmao Lin
ZY Zifu Yan
XW Xiaowu Wang
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4810 Views: 443
Reviewed by: Alessandro DidonnaSailendra Singh Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Horticultural Plant Journal May 2022
Abstract
Since the genetic transformation of Chinese cabbage (Brassica rapa) has not been well developed, in situ RT-PCR is a valuable option for detecting guard cell–specific genes. We reported an optimized protocol of in situ RT-PCR by using a FAMA homologous gene Bra001929 in Brassica rapa. FAMA in Arabidopsis has been verified to be especially expressed in guard cells. We designed specific RT-PCR primers and optimized the protocol in terms of the (a) reverse transcription time, (b) blocking time, (c) antigen-antibody incubation time, and (d) washing temperature. Our approach provides a sensitive and effective in situ RT-PCR method that can detect low-abundance transcripts in cells by elevating their levels by RT-PCR in the guard cells in Brassica rapa.
Keywords: Brassica rapa In situ RT-PCR Guard cells Low abundance Quantitative analysis
Background
In situ RT-PCR combines RT-PCR and in situ hybridization technologies to amplify specific nucleic acid sequences in cells or tissue sections, enabling the localization of specific low-copy number sequences to be detected by immunohistochemistry (Nuovo, 2001). This technique identifies mRNA in the tissues; it was initially used to locate and detect viral gene expression in human and animal cells (Martinez, 1995; Bates et al., 1997; Hoyland et al., 1997; Kher and Bacallao, 2001; Cubas-Nuñez et al., 2017) and was then introduced into plant research. Thus far, this technology has been successfully applied in many plants including pea, Arabidopsis (Deeken and Kaldenhoff, 1997), cucumber (Urbańczyk-Wochniak et al., 2002), tomato (Portillo et al., 2013), barley (Ferdous et al., 2017), and barley and Arabidopsis (Athman et al., 2014); the protocol from Athman et al. (2014) has been widely used in plant research (Hocking et al., 2017; Olsen and Krause, 2019). Bra001929 is a homolog in Brassica rapa to the guard cell–specific gene FAMA in Arabidopsis, which has been identified and analyzed in a wide range of plant species. The Bra001929 gene can also be used as a marker gene for Chinese cabbage guard cells.
Materials and reagents
Leaf epidermis
Double-edged blade (Gillette)
RNaseZap® solution (Ambion, catalog number: AM9786)
Formalin (Sigma, catalog number: F8775-25ML)
Acetic acid (Sigma, catalog number: 695092-500ML)
Ethanol absolute (Sangon Biotech, catalog number: A500737-0500)
DEPC (Sigma, catalog number: D5758-25ML)
Nuclease-free 10× PBS (Solarbio, catalog number: P1022)
RNaseOUTTM recombinant ribonuclease inhibitor (Invitrogen, catalog number: 10777-019)
10× TURBO DNaseTM buffer (Invitrogen, TURBO DNAfreeTM kit, catalog number: AM1907)
DNase I, RNase-free (Invitrogen, catalog number: EN0521)
0.5 M EDTA, pH 8.0, RNase-free (Ambion, catalog number: AM9260G)
EasyScript One-Step gDNA Removal and cDNA Synthesis SuperMix (Transgen, catalog number: AE311-02)
DIG-11-dUTP (Roche, catalog number: 11093088910)
Bovine serum albumin (BSA) (Sigma, catalog number: A6003-100G)
BM Purple AP Substrate (Roche, catalog number: 11442074001)
Tris (Biotopped, catalog number: T6061)
NaCl (Solarbio, catalog number: S8210)
Glycerol (Sigma, catalog number: 56-81-5)
Dilute Anti-DIG-AP in 1% BSA blocking solution (see Recipes)
DNase/RNase free water (see Recipes)
FAA fixative (see Recipes)
Washing buffer I (see Recipes)
1× PBS (see Recipes)
1% BSA blocking solution (see Recipes)
40% glycerol solution (see Recipes)
Equipment
Pointed tweezers (AsOne, catalog number: CC-9820-01)
Super frost plus slides, 76 mm × 25 mm (Fisher Scientific, catalog number: 12-550-15)
Cover glass (CITOGLAS, catalog number: BZ1164)
Frame-SealTM incubation chambers (Bio-Rad, catalog number: SLF-0601)
15 mL centrifuge tube (Corning, catalog number: 433052)
2 mL centrifuge tube (Axygen, catalog number: MCT-060-A)
0.2 mL centrifuge tube (Axygen, catalog number: PCR-02-C)
Metal constant temperature incubator (Coyotebio, catalog number: H2O3-100C)
Water bath thermal cycler, an automated high throughput water bath thermal cycler with flexible modular design for rapid temperature change and precise control. Key technical parameters: three tanks, temperature controlled 40–97 °C (HC Scientific, catalog number: ID 156 B03)
Optical microscope (Zeiss, catalog number: 3150012307)
Software
ImageJ (https://imagej.net/software/imagej/)
Microsoft Excel (Microsoft Corp., Albuquerque, NM, USA)
Procedure
Tissue fixation
First make 20 mL of FAA fixative (see Recipes): 2% formalin, 5% acetic acid, and 60% ethanol well mixed; place in refrigerator at 4 °C.
Fix the Frame-SealTM incubation chambers on the glass slides and add 150 μL of pre-cooled FAA fixative to each glass slide.
Rinse the blade with sterile water before tissue fixation, quickly tear off the epidermis with pointed tweezers, and immediately place it in an ice bath in FAA fixative.
Place the glass slide under the optical microscope to detect whether the epidermal tissue is stuck with mesophyll tissue. If so, then peel the epidermis again; if not, proceed to the next step.
The sample needs to be fixed 3–4 times, 15 min each. Try to remove it with a pipette every time the fixative is changed. During fixation, make washing buffer 1:5% acetic acid and 60% ethanol, mix well, and place in the refrigerator at 4 °C.
After fixation, add 150 μL of washing buffer 1 to each slide to wash the tissue samples on ice three times for 8–10 min each time. To ensure sufficient washing, the slide can be rotated slightly.
Add 150 μL of 1× PBS to each slide and continue washing three times for 3 min each on ice to maintain the ion concentration; finally, add the premix to each slide: 102 μL of RNase-free water + 3 μL of RNaseOUTTM ribonuclease (RNase) inhibitor (see Figure 1).
Figure 1. Schematic of in situ RT-PCR procedure
Removal of genomic DNA
Configuration of master mix: 12 μL of 10× TURBO DNaseTM buffer + 3 μL of 1UμL -1 DNase I. After mixing, add to the sample in a final volume of 120 μL and cover with a cover glass.
Adjust the temperature of the metal bath to 37 °C in advance and place the glass slide in the metal bath for 1 h.
Adjust the temperature of the metal bath to 70 °C, carefully remove the coverslip with tweezers, add 4 μL of 0.5 M EDTA pH 8.0 to the slide, cover the coverslip to inactivate for 15 min, and store at 4 °C in the refrigerator for approximately 5 min.
Use tweezers to carefully remove the cover glass. Since the epidermis may stick to the cover glass during high-temperature incubation, you can use a sterile scalpel to move it into the solution and a pipette to remove as much DNase solution as possible.
Pipette 200 μL of pre-cooled DEPC water into the slide to wash the sample on ice. To ensure sufficient washing, the slide can be rotated slightly.
In order not to affect subsequent experiments, discard as much RNase-free water as possible with a pipette after washing (see Figure 1).
Reverse transcription
In order to obtain higher synthesis efficiency, first configure the primer and RNase-free water master mix: 3 μL of 10 μM gene-specific reverse primer + 84.6 μL of RNase-free water.
Immediately add the premix to the glass slide that has removed RNase-free water and cover it with a cover glass.
Adjust the temperature of the metal bath to 65 °C in advance, incubate the slides at 65 °C for 8 min, and prepare the RT reaction solution during the incubation.
After the incubation, adjust the temperature of the metal bath to 42 °C; then, use the pointed tweezers to carefully remove the cover glass and add the RT premix (Table 1) after the slide is ice-bathed for 2 min. To avoid false positives during the in situ RT-PCR process, we used the control in which the reverse transcription enzyme was not used in the RT reaction.
Table 1. RT reaction premix
Component Volume
2× ES Reaction Mix 30 μL
EasyScript® RT Enzyme Mix 1.2 μL
RNaseOUTTM ribonuclease (RNase) inhibitor 1.2 μL
Bubbles may be present after adding the RT premix; use a 10 μL pipette to carefully pipette them out and then cover the cover glass.
Place the slides in a metal bath and incubate at 42 °C for 1.5 h. Then, adjust the temperature of the metal bath to 85 °C and incubate for 1 min to inactivate the reverse transcriptase. After incubation, remove the slides and place in the refrigerator at 4 °C for 5 min (see Figure 1).
In situ PCR
Because the temperature of the water bath PCR thermal cycler is low, it is necessary to turn on the thermal cycler to preheat at the beginning of the experiment. The PCR reaction is premixed (Table 2). Configure the in situ PCR mix when the slide is placed at 4 °C.
Table 2. Primer design
Primer Primer sequence
Bra001929F 5′-GAA CAA GTC GTG CTT GGC TG-3′
Bra001929R 5′-CAC GTG TTT ATA CCT ACT TGC CT-3′
Vortex and briefly centrifuge the configured in situ PCR mix and place it in a refrigerator at 4 °C.
Use tweezers to carefully remove the cover glass and remove the RT reaction solution.
Take 200 μL of pre-cooled RNase-free water to wash the tissue three times on ice. To ensure sufficient washing, the slide can be rotated slightly.
In order not to affect the subsequent PCR, use a pipette to discard as much RNase-free water as possible after washing.
Add 90 μL of pre-cooled in situ PCR mix (in Table 3) to each sample. If there are bubbles, use a 10 μL pipette to carefully suck them out. Finally, carefully glue the polyester covers and press them firmly along the edge of the frame. Avoid squeezing out the PCR mix.
The primers for Bra001929 were designed according to the following guidelines: primer length of 18–24 bp, GC content of 40%–60%, with the last five bases of the 3' end being less rich in GC, which resulted in the pairs of primers for Bra001929.
The suggested PCR cycle setting is in Table 4. However, annealing temperature and extension time are adjusted according to primer and product size (see Figure 1).
Table 3. PCR reaction
Component Volume
H2O 64.94 μL
5× Phusion HF Buffer* 18 μL
10 mM dNTPs 1.8 μL
Forward primer 4.5 μL
Reverse primer 4.5 μL
Phusion DNA Polymerase 0.9 μL
DIG-11-dUTP 0.36 μL
Table 4. PCR cycling instructions
Cycle step 3-step protocol Cycles
Temperature Time
Initial denaturation 98 °C 30 s 1
Denaturation 98 °C 10 s 35
Denaturation 55 °C 25 s
Extension 72 °C 5 s
Final extension 72 °C 7 min 1
Immunoassay
The frame and the polyester covers are very tightly bonded. If you tear it directly, the frame will also be torn off the glass slide. So Incubate the slide on ice and use a sterile scalpel to cut off the polyester covers along the frame.
After cutting off the polyester covers, carefully remove the PCR mix.
Pipette 200 μL of pre-cooled 1× PBS to wash the tissue on ice three times for 3 min each. To ensure sufficient washing, the slide can be rotated slightly.
Remove as much 1× PBS as possible after each wash.
Adjust the temperature of the metal bath to 37 °C in advance; add 100 μL of 1% BSA blocking solution to the tissue sample, rotate the slide slightly to mix, and incubate at room temperature for 30 min.
Dilute the alkaline phosphatase–labeled anti-Digoxigenin antibody at 1:500 in fresh 1% BSA blocking solution. Each sample needs to be configured with 80 μL: 0.16 μL of Anti-Digoxigenin-AP + 79.84 μL of 1% BSA blocking solution. Store in the refrigerator at 4 °C.
After blocking at room temperature, carefully remove the 1% BSA blocking solution and pipette 100 μL of alkaline phosphatase–labeled anti-digoxigenin antibody binding solution to the tissue sample and incubate at room temperature for 1 h.
After the antigen and antibody incubation, carefully remove the antibody binding solution.
Add 150 μL of elution buffer II to carefully rinse the tissue sample three times, each for 10 min.
After elution, add 70 μL of alkaline phosphatase substrate BM Purple to the tissue sample in the dark (turn the alkaline phosphatase substrate upside down and mix well before use) and leave it at room temperature for 30 min without a cover glass. Then, observe under the optical microscope every 15 min (10× magnification).
Once the purple-blue amplification signal is detected, find the best place to take pictures and archive. Then, remove the alkaline phosphatase substrate, pipette 150 μL of elution buffer II into the slide to wash the sample on ice three times for 2 min each, and then wash once with sterile water on ice. Finally, add 40 μL of 40% glycerol for mounting and cover with a coverslip (see Figure 1).
Data analysis
The positive guard cells exhibited significant hybridization signals using the optimized protocol based on three duplicates (Figure 2). Here, we used the optical density value to assess the hybridization signal intensity. First, ImageJ (1.53c; Wayne Rasband software; National Institutes of Health; USA) was used for image analysis to obtain the optical density of the region of interest (ROI) and the Area (Area of the selected range) and IntDen (IOD of the selected range) of positive guard cells, positive epidermal cells, negative guard cells, and negative epidermal cells in the results. The ratio of IntDen to Area is the average optical density value of a single cell. Data were analyzed using Microsoft Excel (Microsoft Corp., Albuquerque, NM, USA). The results were expressed as mean ± S.E.M. (standard error of the mean, see Figure 3).
Figure 2. Three duplicates of in situ RT-PCR in B. rapa guard cells. A. First duplicate with visualization of the transcripts of the target genes in the guard cells. B. Second duplicate. C. Third duplicate. Red letters represent guard cells and blue letters represent epidermal cells. The control without RT showed no visualization of transcripts in the guard cells. The 8-bit grayscale images were converted by ImageJ software. Scale bars represent 100 μm.
Figure 3. Mean optical density values of three duplicates for different cells in the in situ RT-PCR. A. Mean optical density values of the positive guard cells correspond to the grayscale image of the positive cells marked in red letters (Figure 2b, 2f, 2j). B. Mean optical density values of the control epidermal cells in the positive slides correspond to the grayscale image of the positive cells marked with blue letters (Figure 2b, 2f, 2j). C. Mean optical density values of the control guard cells correspond to the grayscale image of the control cells marked with red letters (Figure 2d, 2h, 2l). D. Mean optical density values of the control epidermal cells correspond to the grayscale image of the control cells marked with blue letters (Figure 2d, 2h, 2l). The error was calculated from the standard deviation of the cell average and different letters on top of the bars indicate significant differences.
Recipes
DNase/RNase free water
Add 1 mL of DEPC to 1,000 mL of deionized water to prepare 0.1% DEPC water. Put it in a 1,000 mL blue cap bottle, shake and mix thoroughly, let it stand for 24 h, and then high pressure for 40 min to remove DEPC.
FAA fixative
2% formalin, 5% acetic acid, 60% ethanol
Washing buffer I
5% acetic acid, 60% ethanol
DNase/RNase free water
Add 1 mL of DEPC to 1,000 mL of deionized water to prepare 0.1% DEPC water. Put it in a 1,000 mL blue cap bottle, shake and mix thoroughly, let it stand for 24 h and then high pressure for 40 min to remove DEPC.
1× PBS
1 mL of 10× PBS + 9 mL of H2O
1% BSA blocking solution
Dilute Anti-DIG-AP in 1% BSA blocking solution: 0.16 μL of Anti-Digoxigenin-AP + 79.84 μL of 1% BSA blocking solution.
Washing buffer I
100 mmol/L Tris-HCl, pH 7.5, 150 mmol/L NaCl
40% glycerol solution
4 mL of glycerol solution + 6 mL of H2O
Acknowledgments
This work is supported by the National Natural Science Foundation of China (Grant No. 31630068), the Key Laboratory of Biology and Genetic Improvement of Horticultural Crops, Ministry of Agriculture, China. This protocol was modified based on the previous work from Athman et al. (2014).
Competing interests
The authors declare no conflicts of interest.
References
Athman, A., Tanz, S. K., Conn, V. M., Jordans, C., Mayo, G. M., Ng, W. W., Burton, R. A., Conn, S. J. and Gilliham, M. (2014). Protocol: a fast and simple in situ PCR method for localising gene expression in plant tissue. Plant Methods 10: 29.
Bates, P. J., Sanderson, G., Holgate, S. T. and Johnston, S. L. (1997). A comparison of RT-PCR, in-situ hybridisation and in-situ RT-PCR for the detection of rhinovirus infection in paraffin sections. J. Virol. Methods 67(2): 153–160.
Ferdous, J., Sanchez-Ferrero, J. C., Langridge, P., Milne, L., Chowdhury, J., Brien, C. and Tricker, P. J. (2017). Differential expression of microRNAs and potential targets under drought stress in barley. Plant Cell Environ. 40(1): 11–24.
Hoyland, J. A., Mee, A. P., Baird, P., Braidman, I. P., Mawer, E. B. and Freemont, A. J. (1997). Demonstration of estrogen receptor mRNA in bone using in situ reverse-transcriptase polymerase chain reaction. Bone 20(2): 87–92.
Hocking, B., Conn, S. J., Manohar, M., Xu, B., Athman, A., Stancombe, M. A., Webb, A. R., Hirschi, K. D. and Gilliham, M. (2017). Heterodimerization of Arabidopsis calcium/proton exchangers contributes to regulation of guard cell dynamics and plant defense responses. J. Exp. Bot. 68(15): 4171–4183.
Kher, R. and Bacallao, R. (2001). Direct in situ reverse transcriptase-polymerase chain reaction. Am. J. Physiol. Cell Physiol. 281(2): C726–732.
Deeken, R. and Kaldenhoff, R. (1997). Light-repressible receptor protein kinase: a novel photo-regulated gene from Arabidopsis thaliana. Planta 202(4): 479–486.
Cubas-Nuñez, L., Duran-Moreno, M., Castillo-Villalba, J., Fuentes-Maestre, J., Casanova, B., Garcia-Verdugo, J. M. and Gil-Perotin, S. (2017). In situ RT-PCR Optimized for Electron Microscopy Allows Description of Subcellular Morphology of Target mRNA-Expressing Cells in the Brain. Front. Cell Neurosci. 11: 141.
Martinez, A., Miller, M. J., Quinn, K., Unsworth, E. J., Ebina, M. and Cuttitta, F. (1995). Non-radioactive localization of nucleic acids by direct in situ PCR and in situ RT-PCR in paraffin-embedded sections. J. Histochem. Cytochem. 43(8): 739–747.
Portillo, M., Cabrera, J., Lindsey, K., Topping, J., Andres, M. F., Emiliozzi, M., Oliveros, J. C., Garcia-Casado, G., Solano, R., Koltai, H., et al. (2013). Distinct and conserved transcriptomic changes during nematode-induced giant cell development in tomato compared with Arabidopsis: a functional role for gene repression. New Phytol. 197(4): 1276–1290.
Nuovo, G. J. (2001). Co-labeling using in situ pcr: A review. J. Histochem. Cytochem. 49(11): 1329–1339.
Olsen, S. and Krause, K. (2019). A rapid preparation procedure for laser microdissection-mediated harvest of plant tissues for gene expression analysis. Plant Methods 15: 88.
Urbańczyk-Wochniak, E., Filipecki, M. and Przybecki, Z. (2002). A useful protocol for in situ RT-PCR on plant tissues. Cell Mol. Biol. Lett. 7(1): 7–18.
Article Information
Copyright
© 2023 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 > Cell imaging
Molecular Biology > RNA > RNA detection
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4,811 | https://bio-protocol.org/en/bpdetail?id=4811&type=0 | # Bio-Protocol Content
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Peer-reviewed
Absolute Quantification of mRNA Isoforms in Adult Stem Cells Using Microfluidic Digital PCR
SB Shubhangi Das Barman
ZF Zofija Frimand
AM Antoine de Morree
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4811 Views: 428
Reviewed by: Philipp WörsdörferSébastien GillotinHeng Sun
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Original Research Article:
The authors used this protocol in Science Nov 2019
Abstract
Adult stem cells play key roles in homeostasis and tissue repair. These cells are regulated by a tight control of transcriptional programs. For example, muscle stem cells (MuSCs), located beneath the basal lamina, exist in the quiescent state but can transition to an activated, proliferative state upon injury. The control of MuSC state depends on the expression levels of myogenic transcription factors. Recent studies revealed the presence of different mRNA isoforms, with distinct biological regulation. Quantifying the exact expression levels of the mRNA isoforms encoding these myogenic transcription factors is therefore key to understanding how MuSCs switch between cell states. Previously, quantitative real-time polymerase chain reaction (qRT-PCR) has been used to quantify RNA expression levels. However, qRT-PCR depends on large amounts of RNA input and only measures relative abundance. Here, we present a protocol for the absolute quantification of mRNA isoforms using microfluidic digital PCR (mdPCR). Primary MuSCs isolated from individual skeletal muscles (gastrocnemius and masseter) are lysed, and their RNA is reverse-transcribed into cDNA and copied into double-stranded DNA. Following exonuclease I digestion to remove remaining single-stranded DNA, the samples are loaded onto a mdPCR chip with TaqMan probes targeting the mRNA isoforms of interest, whereupon target molecules are amplified in nanoliter chambers. We demonstrate that mdPCR can give exact molecule counts per cell for mRNA isoforms encoding the myogenic transcription factor Pax3. This protocol enables the absolute quantification of low abundant mRNA isoforms in a fast, precise, and reliable way.
Graphical overview
Schematic overview of the workflow. (A) Isolation of individual muscles (gastrocnemius and masseter) from C57/BL6 mice followed by digestion using collagenase II and dispase. (B) Sorting of 500 cells directly into PCR tubes using fluorescence-activated cell sorting (FACS). (C) Reverse transcription of mRNA to cDNA. (D) Polymerase reaction to generate a duplicated cDNA product. (E) Exonuclease I digestion to remove remaining single-stranded DNA and the non-hybridized primers. (F) Denaturation step to inactivate exonuclease I. (G) Loading the samples into the microfluidic chip. (H) Running the TaqMan Digital PCR assay in the Fluidigm Biomark HD real-time PCR machine. (I) Data analysis using the Digital PCR software.
Keywords: Digital PCR Microfluidics Muscle stem cells mRNA isoforms Pax3 Alternative polyadenylation Skeletal muscle
Background
Muscle stem cells (MuSCs) reside beneath the basal lamina of skeletal muscle fibers and maintain a state of quiescence (de Morree and Rando, 2023). Upon injury, the MuSCs switch from quiescence to activation, giving rise to proliferating progenitors that will eventually differentiate and fuse with myofibers (Yin et al., 2013;Relaix et al., 2021). To understand the exit from quiescence and how the myogenic fate is determined, it is important to understand the factors that maintain or break the quiescent state The transcription factor Pax3 plays a key role in myogenesis. In the absence of Pax3, embryonic MuSCs fail to expand and migrate, and no muscle is formed, resulting in embryonic lethality (Relaix et al., 2005). Recent work revealed that thePax3gene generates multiple mRNA isoforms with different 3′UTRs in MuSCs from adult mice (Boutet et al., 2012; de Morree et al., 2019). One of the isoforms of Pax3 mRNA has a long 3′UTR containing microRNA binding sites, is translationally repressed by the microRNA 206, and is linked to quiescence (Figure 1). The other has a short 3′UTR lacking the microRNA binding sites, is not translationally repressed, and is linked to activation and cell cycle entry of MuSCs (de Morree et al., 2019). Therefore, it is important to understand Pax3 isoform expression patterns in MuSCs, as quantifying the two isoforms of Pax3 will contribute to understanding its role in MuSCs quiescence and activation.
Figure 1. Schematic figure depicting how the TaqMan probes, Pax3total and Pax3long, target their respective target sites on the Pax3 3′ terminal exon. Top: a schematic of thePax3gene with boxes denoting exons. The 3′ terminal exon is enhanced, and the open reading frame (ORF) coding part is highlighted, as well as the three polyadenylation sites (PASs) present in the 3′ untranslated region (3′UTR). Bottom: three major Pax3 mRNA isoforms are represented. All contain the complete ORF (not depicted) but differ in the length of the 3′UTR due to the selection of different PASs. These isoforms can be detected with TaqMan probes. The Pax3total TaqMan probes hybridize to sequences that are present in both long and short isoforms of Pax3. The Pax3long TaqMan probes hybridize to sequences that are present only in long isoform of Pax3. Each TaqMan probe contains a reporter dye and a quencher. During the polymerase reaction, the dye and the quencher are separated from the target sequence; as a result, fluorescence increases.
Quantitative real-time polymerase chain reaction (qRT-PCR) is often considered as the gold standard for measuring mRNA expression levels, despite many factors of variability such as RNA templates, inconsistent data analysis, and data normalization (Nolan et al., 2006). This method uses intercalating fluorescent dyes to assess transcript abundance during a PCR amplification. Though effective, it cannot measure exact copy numbers and requires a reference gene like GAPDH or HPRT to calculate relative expression levels. These reference genes are used because they are considered stably expressed in all cells. However, they may not be (Kozera and Rapacz, 2013). MuSCs undergo a 6-fold increase in size during activation (Rodgers et al., 2014;Brett et al., 2020). Thus, the change in cell volume may affect the interpretation of relative quantification by qRT-PCR, complicating the interpretation of how much Pax3 is expressed in cells. Instead, several adaptations have been made that enable single molecule quantification using a technique called digital PCR (Mao et al., 2019). In digital PCR, the total reaction volume is partitioned into small reaction chambers at a dilution so that 5%–65% of the small reaction chambers contain a single molecule of target cDNA. Then, a PCR amplification is performed to amplify those molecules, enabling the researcher to simply score for each small reaction chamber whether a PCR reaction took place or not. This digital assessment can then be calculated into an exact number of molecules present in the original reaction volume using Poisson statistics. Recent technologies that enable the formation of small droplets (droplet digital PCR) made digital PCR more routinely accessible (Hindson et al., 2013;Mao et al., 2019).
However, while droplet digital PCR enables the low-cost quantification of absolute mRNA levels, there is a variability between droplets that lowers the precision (Emslie et al., 2019), which is key when measuring low-abundant transcripts. Here, we use a parallel technology, microfluidic digital PCR (mdPCR), in which the sample is partitioned in microfluidic nanochambers (see Graphical overview). These nanochambers are uniform and enable highly reproducible amplification reactions, to enable the absolute quantification of low abundant transcripts with high accuracy. We previously applied this protocol to find that MuSCs isolated from lower hindlimb muscle expressed on average three molecules of Pax3 mRNA, two thirds of which were long isoforms. In contrast, MuSCs isolated from the diaphragm muscle expressed on average 10 molecules of Pax3 mRNA, one third of which were long isoforms (de Morree et al., 2019).
This protocol can also be used to quantify mRNA isoforms in other stem cell types, such as muscle-resident fibroadipogenic progenitors, which play important roles in homeostasis and repair of multiple tissues (Joe et al., 2010;Uezumi et al., 2010). The levels of mRNA isoforms can be quantified accurately across different cell states. Expression of disease genes along the progression of a disease can also be studied using mdPCR. Finally, the efficacy of therapies that change relative isoform abundance, such as exon-skipping for Duchenne Muscular Dystrophy, can be assessed with mdPCR (Verheul et al., 2016). Hence, this protocol can be used both in basic biology and translational studies.
Materials and reagents
GE 48.48 Dynamic Array Reagent kit with control line fluid, 10 IFCs (Fluidigm, catalog number: 85000800) (contains the control line fluid, 20× GE Sample Loading Reagent)
TE buffer, pH 8.0 (Teknova, catalog number: T0224)
CellsDirect One Step qRT-PCR kit (Life Technologies Invitrogen, Thermo Fisher Scientific, catalog number: 46-7201)
TaqMan Fast Advanced Master Mix (Applied Biosystems, Thermo Fisher Scientific, catalog number: 444455)
Pipettes (p10, p20, p200, p1000) (Mettler Toledo, Rainin Pipet-Lite XLS+)
Cells of interest (we used MuSCs for the protocol)
Ethanol 70% (v/v) (TechniSolve®, VWR Chemicals)
Exonuclease I and buffer (New England Biolabs, M0658S)
PCR strip of 8,200 μL, PCR performance tested, transparent, PP, flat cap (Sarstedt, catalog number: 72.991.002)
12.765 Digital ArrayTMIFC (Standard Biotools, Fluidigm, BMK-M-12.765)
TaqMan probe Pax3total (Boutet et al., 2012; de Morree et al., 2019): Pax3total_Forward: 5′ GTCAGAGACTGGAACATATGAAGAATGT 3′ Pax3total_Reverse: 5′ GCCTGCGGTGCTATAGGT 3′ Pax3total_Middle: 5′ CTGCCCACATCTCAGC 3′
TaqMan probe Pax3long (Boutet et al., 2012; de Morree et al., 2019): Pax3long_Forward: 5′ TGTGTTAGCAGGACTAGACATAGAACT 3′ Pax3long_Reverse: 5′ TTGAGGCTGCAACACAAAGATACTA 3′ Pax3long_Middle: 5′ CACCCTGCCCAACGTA 3′
RNaseZap™ RNase Decontamination solution (ThermoFisher Scientific, catalog number: AM9780)
Equipment
BioMark HD PCR machine (Fluidigm)
Integrated Fluidic Circuit (IFC) controller MX (Fluidigm)
Mini centrifuge (Fisherbrand, catalog number: 12006901)
ScanR High Content Screening microscope (Evident, model: OLYMPUS)
BD FACSAriaTMIII Sorter (BD Biosciences, catalog number: 648282-24)
Master Cycler Gradient Thermal Cycler (Eppendorf, catalog number: 5331)
Vortex-Genie 2 (Scientific Industries, SKU: SI-0236)
Software
Digital PCR Analysis Software (Biomark and EP1 software v4.8.1)
GraphPad PRISM (GraphPad Software Inc.)
Fiji image processing package (ImageJ2)
Procedure
Preparation
Program the thermal cycler with protocols for the reverse transcriptase and polymerase step, as well as the exonuclease step.
Set the program for reverse transcription: heat the lid to 105 °C, incubate the sample for 15 min at 50 °C (step to reverse transcribe the RNA to cDNA), then incubate the sample for 2 min at 95 °C (step to inactivate the reverse transcriptase and activate the Taq polymerase), incubate the sample for 15 s at 95 °C, and then incubate the sample for 4 min at 60 °C (specific target amplification, to make double-stranded DNA). Finally, incubate the sample at 4 °C to preserve the DNA.
Set the program for the exonuclease reaction: heat the lid to 105 °C and incubate the sample for 10 min at 37 °C (to digest all single-stranded DNA and the non-hybridized primers). Then, incubate the sample for 10 min at 80 °C (to denature and inactivate the exonuclease). Finally, incubate the sample at 4 °C to preserve the DNA.
Cell sorting
Prepare a single-cell suspension of your cell type of interest. Here, we digested the gastrocnemius (calf muscle) and masseter (jaw muscle) and generated single-cell suspensions (Frimand et al., 2022).
In our experiment, we were interested in MuSCs, which can be isolated based on the cell surface markers CD31, CD45, Sca1 (all negative markers), and Vcam1 (positive marker). We therefore stained our single-cell suspension with antibodies against CD31, CD45, Sca1, and Vcam1 in order to isolate the MuSCs as the Vcam1+/Sca1-/CD31-/CD45- cells, as described previously (Frimand et al., 2022).
Note: If other cell types shall be purified, alternative markers need to be determined.
Start up the FACS ARIA III and perform the CS&T and Accudrop controls following the BD Biosciences Manual. Next, calibrate the droplet position of the FACS. To this end, install the adapter for plate sorting and bring out the plate sorting arm (seeFigure 2).
Figure 2. FACS ARIA III cell sorter setup. (A) Setup of the BD Aria 3 sorting arm with 96-well plate holder and a PCR strip inserted into the 96-well plate holder. (B) The sorting arm with 96-well plate holder is moved to the home position. Arrows highlight the plate sort adapter (1), the plate sorting arm (2), and a PCR strip after test sort, showing the test droplet in the center of each lid (3 and inset). (C) PCR strip with lids open inserted into the 96-well plate holder for sorting cells. (D) Sorting 500 cells into each tube of the PCR strip.
Insert a PCR strip with lids closed into a 96-well plate holder (Figure 2A), install the plate holder onto the plate sorting arm, and bring the arm to the home position (Figure 2B). Test sort a droplet and check its position on top of the lid; it should be dead center (Figure 2B). Adjust the position of the plate if necessary and save the new home position.
Add 5 μL of 2× Reaction Mix from the Cells Direct PCR kit per tube of the PCR strip. Prepare the number of tubes required for your experiment (in our case, six), cut off the remaining tubes from the PCR strip with scissors, and discard them. Position the PCR strip (with open lid!) into the holder on the plate sorting arm (Figure 2C).
Sort 500 cells of the first single-cell suspension into the first tube, making the volume in that sample tube 6 μL (5 μL of 2× Reaction Mix + 1 μL of sorted cells) (Figure 2D). Unload the sample, flush the sample- line, and load the next sample. Sort 500 cells into the next tube. Proceed until all tubes have received 500 cells. Here, we sorted 500 MuSCs from the gastrocnemius and masseter muscles from three mice.
Notes:
The CellsDirect Reaction Mix kit can be used for a maximum of 1,000 cells. Depending on the mRNA content of the cell, more or fewer cells can be used as input with a maximum of 1,000 cells. We recommend confirming sorting efficiency by sorting 500 cells into a coated well of a 96-well plate and counting the cell number using microscopy.
The 2× Reaction Mix is one of the components of the CellsDirect One Step qRT-PCR kit. It consists of dNTPs, reaction buffer, and magnesium sulphate, which supports the activity of the polymerase enzyme.
Close the lids on the PCR strip, vortex, and spin down immediately in a mini centrifuge. The samples can be kept on ice to be used immediately or stored at -20 °C for short term (1–2 weeks) or at -80 °C for long term (6–8 months).
Note: The lower the temperature, the higher the stability of the mRNA collected in the sample tubes (Jaquet et al., 2022).
CRITICAL STEP: Vortex the sample tubes thoroughly to mix the sample components and spin them down in a mini centrifuge before storage.
Reverse transcriptase-polymerase reaction
CRITICAL STEP: To prevent RNase contamination, this section of the protocol is carried out under sterile and RNase-free conditions. Wear gloves and spray down the workspace with 70% EtOH. Further, all the steps are performed on ice to improve RNA stability. The chemical RNaseZapis recommended to clean the bench to prevent RNase contamination.
CRITICAL STEP: The contents of the CellsDirect Reaction mix need to be thawed on ice before use and all the steps need to be performed in ice.
Thaw the TaqMan probes and the remaining additives (DEPC-treated water, TaqMan probes, resuspension buffer, SuperScriptTMIII RT/PlatinumTMTaq Mix, and the ROX reference dye) of the CellsDirect kit.
Prepare a master mix (CellsDirect Reaction mix) for all your sample tubes (in our experiment, six tubes: three sample tubes with MuSCs sorted from gastrocnemius muscle and three sample tubes with MuSCs sorted from masseter muscle).
For six sample tubes, the mix is prepared by adding 24 μL of resuspension buffer, 1.2 μL of SuperScriptTMIII RT/PlatinumTMTaq Mix, 1.2 μL of TaqMan probe sets (for forward and reverse primer, respectively, making the final volume of each set 2.4 μL), 2.3 μL of DEPC-treated water, and 0.1 μL of ROX reference dye to a RNase-free 1.5 mL Eppendorf tube. Here, we use TaqMan probes for Pax3total and Pax3long (each targeting individual sequences in the Pax3 mRNA).
Notes:
The Pax3long TaqMan probe set hybridizes to a site that is present only in long isoform of Pax3, while the Pax3total TaqMan probe set hybridizes to a site that is present in both long and short isoforms of Pax3 (Figure 2), each generating different amplicons of Pax3 reflecting different sites of the transcript.
We recommend always preparing the master mix for n samples + 1 to ensure that there is sufficient volume for pipetting.
Retrieve the PCR strip containing the lysed cells in CellsDirect 2× on ice. If the cells were stored and frozen previously, then thaw on ice.
Add 4 μL of the CellsDirect Reaction mix (master mix), making the final volume in each sample tube 10 μL. Vortex and spin the tubes down in a mini centrifuge and immediately transfer the tubes to ice after they have been spun down. CRITICAL STEP: Vortexing the sample tubes helps to evenly distribute the molecules in the tube. This is an important step to ensure that the molecules in the reaction volume are homogeneously distributed. However, it must be followed with a centrifugation step to pool all the reaction volume at the bottom of the tube.
Place the sample tubes in the thermal cycler and close the machine.
Run the reverse transcriptase-polymerase reaction (see step A1 for the program).
After the reverse transcriptase program is completed, place the tubes containing the cDNA (10 μL of total volume) on ice. Briefly spin down the samples in a mini centrifuge and add 1.2 μL of 10× ExoI buffer and 1 μL of exonuclease I into each of the tubes for a total volume of 12.2 μL per tube (in our case, six sample tubes). Vortex and spin the tubes down in a mini centrifuge and immediately transfer the tubes to ice after they have been spun down.
Note: The exonuclease buffer facilitates the activity of the exonuclease, which digests the single-stranded DNA, including all the non-hybridized TaqMan probes. This step ensures that these non-hybridized probes do not interfere with the downstream TaqMan detections: Pax3total and Pax3long during mdPCR.
Place the sample tubes back in the thermal cycler, close the machine, and run the exonuclease program (see step A2 for the program)
Once the exonuclease program has completed, immediately place the tubes containing the double-stranded cDNA back on ice.
The samples can be used immediately or stored at -20 °C for short term (1–2 weeks) or at -80 °C for long term (6–8 months) storage.
Priming the 12.765 Digital ArrayTMIntegrated Fluidic Circuit (IFC)
The following steps adhere to the manufacturer’s protocol.
CRITICAL STEP: This section of the protocol is performed under sterile conditions. Wear gloves and avoid working above the microfluidic chip to prevent any fluid from dropping on top of the chip, which might affect the accurate detection of gene expression.
CRITICAL STEP: Switch on the IFC Controller MX machine. It takes 10 min to warm up.
Unwrap the microfluidic chip. Avoid touching the center of the chip. Any pressure will damage the microfluidic channels. An unwrapped chip has to be used within 24 h.
Note: This protocol can be performed using other digital PCR chips by Fluidigm.
Inject the control line fluid in each of the accumulators on the chip by pushing down the black O-ring. Beware not to splash any of the oily control line fluid.
Note: In order to facilitate easy flow of the liquid and to avoid air bubbles, inject the control line fluid into each of the accumulators at an angle of 45 degrees. The fluid prevents evaporation of the sample and helps to seal the chip.
Insert the chip into the IFC Controller.
Run the Prime (115×) script to prime the chip by dispersing the oil. It takes approximately 20 min to run the script.CRITICAL STEP: Once primed, the chip must be loaded within an hour to prevent the microfluidic chambers from collapsing.
Preparing the sample pre-mix for Digital PCR
CRITICAL STEP: This section of the protocol is carried out under sterile and RNase-free conditions. Wear gloves and spray down the workspace with 70% EtOH.
Prepare a master mix for each target mRNA isoform. Prepare a master mix A for six samples, consisting of 18 μL of DNA-free water, 30 μL of TaqMan Gene Expression Master Mix, 3 μL of 20× GE Sample Loading Reagent, and 1.5 μL of the Pax3long TaqMan probe in a 1.5 mL Eppendorf tube. Each TaqMan probe consists of the forward, reverse, and middle primer.
Prepare a master mix B for six samples, consisting of 18 μL of DNA-free water, 30 μL of TaqMan Gene Expression Master Mix, 3 μL of 20× GE Sample Loading Reagent, and 1.5 μL of Pax3total TaqMan probe in a 1.5 mL Eppendorf tube.
Prepare two PCR tube strips. Add 9 μL of master mix A to the first six sample tubes of the first PCR tube strip and 9 μL of master mix B into the first six sample tubes of the second PCR tube strip.
Add 1 μL of the cDNA from step C10 into each of the PCR-sample tubes containing master mix A and master mix B, bringing the final volume to 10 μL.
Vortex the PCR sample tubes and spin them down in a mini centrifuge. Place them immediately on ice.CRITICAL STEP: Vortexing the sample tubes helps to evenly distribute the molecules in the tube. This is an important step to ensure that the samples in the chip are homogeneous.
Loading the samples onto the chip
CRITICAL STEP: Wear gloves and spray down the workspace with 70% EtOH to avoid contamination.
Remove the primed chip from the IFC Controller MX once the priming program has finished.
Add 8 μL of DNA-free water into each of the H inlets. Pipette 8 μL from each sample tube to the individual wells.CRITICAL STEP: Load the samples directly into the wells and avoid going above the chip to prevent cross-contamination. To prevent bubbles while loading the sample, only pipette until the first resistance on the pipette.
Return the chip to the IFC Controller MX and select the Load (115×) script to load the samples into the chip. It takes approximately 45 min to load the samples onto the chip.
Note: Always make sure that the correct script is selected before running the software.
Switch on the Fluidigm Biomark HD Real-Time PCR machine while the chip load program is running.
Note: The Fluidigm Biomark HD Real-Time PCR machine needs to be switched on prior to use so that the CCD camera can cool down to the optimum temperature (-5 °C).
Real-time PCR
After the samples are loaded, remove the chip from the IFC Controller MX.
Insert the chip into the Fluidigm Biomark HD Real-Time PCR in a direction in which the barcode is facing outside and click Load.
CRITICAL STEP: When putting the chip into the drawer of the BioMark HD, slowly remove the protective sticker from underneath the chip. Removal is required for any light to pass through the chip during the amplification reaction.
Once the samples are loaded onto the chip, open the data collection software icon on the desktop connected to the Fluidigm Biomark HD Real-Time PCR.
Click on the option Start a New Run.
CRITICAL STEP: Double-check if the status bar on the software shows that both the camera and lamp are green before loading the chip into the machine. The icons will both be green if they are ready. Otherwise, wait until the temperature calibration is completed.
Once the system reads the barcode on the chip, verify the barcode and the chip type on the software.
Next, select the chip run file to run the program. Enter the chip information and save it before proceeding to the next step.
Select the specific application type (Digital PCR), reference dye (ROX), and the assay probes. Select the probe types and click next.
Click on the dPCR Standard v1.pcl protocol and start the run. A standard run takes approximately 1.5 h.
Result analysis using the Digital PCR Analysis Software
Select the Digital PCR Analysis software icon on the software and click on it once the run is completed.
To open the specific chip run file, click on the Open a Chip Run option and double-click on the chiprun.bml file to open it in the software.
Under the Sample and Detector setup option in the Chip explorer plane, highlight and annotate your wells (we named our samples as GA1, GA2, and GA3 and Mass1, Mass2, and Mass3) under the Sample Type option from the drop-down menu in the Editor.
Next, choose the Detector Type from the same menu and name them as Pax3total and Pax3long.
Click on the Update option to finalize the changes made. For the data analysis to begin, click on the Analyze option.
Note: The software has three ways of baseline correction. It is recommended to test all three methods and evaluate the appearance of the signal curves, which should be S-shaped. We recommend using the Constant baseline correction.
Click Analyze.
Once the analysis is done, click on the Panel Summary or Panel Details to view the result sheet in the form of table, image, or a heatmap.
Export all the files with the .csv extension.
Data analysis
We isolated MuSCs from the gastrocnemius and masseter muscles from three wildtype C57/BL6 mice that were discontinued from a local breeding program. The muscles were digested using collagenase II and dispase, followed by staining and preparation for sorting (Frimand et al., 2022). From the stained single-cell suspensions, 500 MuSCs were then sorted using a BD ARIA III FACS directly into sample tubes in a PCR tube strip containing 2× reaction mix (a total of six tubes: three each of gastrocnemius and masseter). The accuracy of the sort was confirmed by sorting an additional 500 cells into a collagen-coated well in a 96-wells plate and counting the cells by microscopy after staining with Hoechst (Figure 3A). Following the extraction and single step amplification of cDNA using reverse transcriptase polymerase reaction, mdPCR was performed using TaqMan probes for Pax3total and Pax3long (Figure 3B–3D). The numbers of Pax3total and Pax3long transcripts on average per MuSC can be calculated using the equation 1 below. The numbers of Pax3total transcripts were 3.87 (± 1.93) and 17.24 (± 0.57) on average per MuSC for gastrocnemius and masseter, respectively. The numbers of Pax3long transcripts were 3.6 (± 0.97) and 2.8 (± 1.81) on average per MuSC for gastrocnemius and masseter (Figure 3E). The numbers of Pax3short transcripts as calculated from equation 2 were 0.25 (± 0.25) and 14.5 (± 2.21) on average per MuSC for gastrocnemius and masseter, respectively. The results from mdPCR can be validated using single molecule RNA fluorescent in situ hybridization (Gaspar and Ephrussi, 2015; de Morrée et al., 2017; de Morree et al., 2019).
Figure 3. Absolute quantification of Pax3 mRNA isoforms in muscle stem cells (MuSCs) using microfluidic digital PCR (mdPCR). (A) Hoechst-stained cells (500 cells) sorted into a collagen-coated well of a 96-well half bottom plate to verify sorting efficiency. One 20× field is shown. (B) PCR amplification graph of Pax3total mRNA isoforms on one sample of gastrocnemius MuSC cDNA. (C) Heatmap of the mdPCR results of Pax3total detection of gastrocnemius MuSC cDNA. The red boxes denote nanochambers in which a signal was amplified. (D) Table representing the relative abundance of Pax3long and Pax3short isoforms in MuSCs isolated from gastrocnemius and masseter muscles (n = 3). (E) Graph depicting the relative amount of Pax3long isoforms as a percentage of total Pax3 mRNA transcripts in MuSCs isolated from gastrocnemius and masseter muscles (n = 3, *= p < 0.05, unpaired two-tailed Student’s t-test).
Equation 1: Calculating the mean number of molecules per cell
(Vloading× Vcellsdirect)/(Vchamber× Nchambers)/Ncells= X molecules per cell
Vloadingis the loading volume for the chip (in this protocol, 10 μL, see step C4);
Vcellsdirectis the volume of the initial cDNA reaction (in this protocol, 12.2 μL, see step C7);
Vchamberis the volume of one nanochamber (for this chip, 0.85 nL);
Nchambersis the total number of nanochambers used for a sample (for this chip, 765);
Ncellsis the number of cells added to the cDNA synthesis reaction (in our experiment, 500);
X is the number of positive nanochambers.
(10 μL × 12.2 μL)/(0.85 nL × 765)/500 = 0.375X
Each chamber has a 0.85 nL volume. Each sample is spread over 765 chambers (in the case of a 12.765 chip) with a combined volume of 0.65 μL. In that volume, we detected X number of positive chambers that contained one molecule. X number of positive chambers multiplied by (10 μL/0.65 μL = 12.3) is 15.4X molecules per loading mix and thus per 1 μL of sample. Multiply 15.4X by 12.2 μL to get the total molecules per sample at 187.7X. Divide 150.1X by 500 cells to get 0.375 times X molecules per cell. Fill in the number of positive nanochambers for X to get the number of transcripts per cell.
Equation 2: Calculating isoform abundance
Calculation:
Xpax3total - Xpax3long = Xpax3short molecules per cell
Xpax3total is the number of detected Pax3total molecules per cell. Xpax3long is the number of detected Pax3long molecules per cell. Xpax3short is the number of calculated Pax3short molecules per cell.
General notes and troubleshooting
General notes
This protocol allows for the absolute quantification of mRNA isoforms, as demonstrated by our measurements of isoforms in quiescent MuSCs. The TaqMan probes selected are designed to specifically anneal to the site of interest (Pax3total targeting both short and long isoforms and Pax3long targeting the Pax3long isoforms). During the extension phase of each polymerase reaction in the mdPCR chip, the 5′ nuclease activity of the enzyme will cleave the 5′ reporter dye from the perfectly hybridized probe, resulting in increased fluorescence or signal (Liu et al., 2013). This protocol allows a precise calculation of the number of Pax3long and Pax3short transcripts per MuSC. It can also be used to detect other types of mRNA isoforms, including alternative splicing isoforms and alternative 5′ UTR isoforms.
TaqMan Digital PCR works on the TaqMan Probe-based Assay chemistry. There are alternative ways of detecting amplified PCR products that are based on EVA Green Dye based chemistry (McDermott et al., 2013). However, this dye might target non-specific sequences of double-stranded DNA (Mao et al., 2007). In contrast, the TaqMan-based assay is highly specific because it requires specific hybridization between the three probes and the target to initiate fluorescent signals (Liu et al., 2018).
This protocol is cost effective due to the low amount of reagents required to run the protocol (Mao et al., 2019). However, mdPCR chips are expensive and for single-time use. The cellular limit of the CellsDirect Reaction Mix is also restricted, which means that when RNA content exceeds the limit, not all can be converted into cDNA. This could lead to an incorrect assessment of the number of molecules detected. In order to prevent this, use a maximum of 500 cells as input for the CellsDirect kit. To ensure that the number of molecules detected is accurate, it is also important to vortex all the sample tubes thoroughly in order to homogenize the samples before loading it on the chip. This can also be ensured by increasing the volume of cDNA used for the sample mix.
Acknowledgments
Cell sorting was performed at the FACS Core Facility, Aarhus University, Denmark. Figures were created using Biorender.com. We thank Dr. Line Reinert, Aarhus University, for assistance with the BioMark HD. This work was supported by a Start Package grant (0071116) and a Project Grant (0080556) from NovoNordiskFonden to A.D.M. This protocol was applied and validated in a previous publication (de Morree et al., 2019).
Competing interests
The authors have no competing financial interests and no conflicts of interest.
Ethics considerations
The protocol was performed in accordance with animal care guidelines at Aarhus University and local ethics regulations.
References
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Spatial Centrosome Proteomic Profiling of Human iPSC-derived Neural Cells
FU Fatma Uzbas *
AO Adam C. O’Neill *
(*contributed equally to this work)
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4812 Views: 789
Reviewed by: Alessandro DidonnaAbraam YakoubYiqun Yu
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Original Research Article:
The authors used this protocol in Science Jun 2022
Abstract
The centrosome governs many pan-cellular processes including cell division, migration, and cilium formation. However, very little is known about its cell type-specific protein composition and the sub-organellar domains where these protein interactions take place. Here, we outline a protocol for the spatial interrogation of the centrosome proteome in human cells, such as those differentiated from induced pluripotent stem cells (iPSCs), through co-immunoprecipitation of protein complexes around selected baits that are known to reside at different structural parts of the centrosome, followed by mass spectrometry. The protocol describes expansion and differentiation of human iPSCs to dorsal forebrain neural progenitors and cortical projection neurons, harvesting and lysis of cells for protein isolation, co-immunoprecipitation with antibodies against selected bait proteins, preparation for mass spectrometry, processing the mass spectrometry output files using MaxQuant software, and statistical analysis using Perseus software to identify the enriched proteins by each bait. Given the large number of cells needed for the isolation of centrosome proteins, this protocol can be scaled up or down by modifying the number of bait proteins and can also be carried out in batches. It can potentially be adapted for other cell types, organelles, and species as well.
Graphical overview
An overview of the protocol for analyzing the spatial protein composition of the centrosome in human induced pluripotent stem cell (iPSC)-derived neural cells. ① Human iPSCs are expanded, which serve as the starting cell population for the neural induction (Sections A, B, and C in Procedure). ② Neurons are induced and differentiated for 40 days (Section D in Procedure), in at least four biological replicates. ③ Total protein is isolated either at 15th or 40th day of differentiation, for neural stem cells and neurons, respectively (Sections E and F in Procedure). ④ Selected bait proteins are immunoprecipitated using the respective antibodies (Sections G and H in Procedure). ⑤ Co-immunoprecipitated samples are analyzed with mass spectrometry (Section I in Procedure). ⑥ Mass spectrometry output (.RAW) files are processed using MaxQuant software to calculate intensities (Section A in Data analysis). ⑦ The resulting data are pre-processed, filtered, and statistically analyzed using Perseus and R software (Sections B and C in Data analysis) ⑧ Further analysis is done using software or web tools such as Cytoscape or STRING to gain biological insights (Sections D and E in Data analysis).
Keywords: Centrosome Spatial proteomics Interactome Microtubule organization Mass spectrometry Human iPSC-derived neurons Perseus MaxQuant
Background
The centrosome is a multifunctional organelle that is known primarily for its microtubule organizing functions, as well as roles related to cell cycle, cell polarity, and migration (Bornens, 2021). Embedded in the pericentriolar material, the centrosome is a highly ordered structure whose core is made of a pair of orthogonally oriented centrioles, each with nine triplets of microtubules with differing maturity—named the mother and daughter centrioles (Fu et al., 2015).
The centrioles exhibit structural differences such as the presence or absence of subdistal and distal appendages, which also facilitate their functional distinction. For example, the more mature mother centriole has a higher capacity to anchor microtubules via these appendages, which are also used in membrane docking—a prerequisite in cilium formation (Bornens, 2002; Brugués et al., 2012; Paridaen et al., 2013; Tanos et al., 2013). The pericentriolar material surrounding the centrosome also serves important functions such as microtubule nucleation, and also contains important signaling proteins (Woodruff et al., 2014; Loukil et al., 2017; Lin et al., 2022).
Although traditionally assumed to be largely composed of a set of homogenous proteins across cell types that support its generic functions, several centrosomal proteins have been identified based on candidate studies that reveal the presence of cell type–specific dynamic relationships (Camargo Ortega and Götz, 2022). For example, the centrosomal protein formerly named AT-hook-containing transcription factor (AKNA) differentially localizes to the subdistal appendages of the centrosomes in neural stem cells and neurons to govern specific processes associated with brain development as well as cell fate determination (Camargo Ortega et al., 2019). NINEIN is another dynamically associated centrosome protein, whose loss from the subdistal appendages in neurons leads to the loss of centrosomal microtubule organizing functions (Wang et al., 2009; Shinohara et al., 2013; Zhang et al., 2016).
Although centrosome proteomes have been cataloged for cancer cells and Drosophila (Andersen et al., 2003; Sauer et al., 2005; Müller et al., 2010; Gheiratmand et al., 2019), these few dynamically identified proteins call for more widespread analysis of interacting partners among a range of cell types. The large amount of cellular material needed for centrosome isolation via fractionation and its inconsistent fractionation behavior prohibits, however, large-scale analyses and does not inform on the location of the newly identified interacting partners on the centrosome. Bio-ID facilitates such proximity labeling, but is compounded by overexpressing a tagged fusion protein, potentially impacting endogenous centrosome functions that might be sensitive to its levels within a cell (Sears et al., 2019). Proteins identified through Bio-ID protocols may also not biochemically interact but rather just localize near the protein/structure of interest (Rattray and Foster, 2019).
We recently applied co-immunoprecipitation of core endogenous centrosomal proteins coupled with mass-spectrometry to outline an effective alternative to both fractionation and tagging strategies (O’Neill et al., 2022). Using core centrosomal proteins with overlapping but non-redundant positions within this organelle, we could resolve spatial interaction networks across the cell types investigated (human dorsal neural stem cells and neurons), identifying the centrosome as a hub for RNA-binding proteins that also explain how ubiquitous proteins can cause organ-specific disease phenotypes when mutated. The use of the microtubule depolymerizing agent, nocodazole, offered an additional mechanism to identify the microtubule-dependent and -independent interacting partners within the cell types of interest (O’Neill et al., 2022).
Here, we outline in detail this method for interrogating the spatial centrosome proteome in human cells. We begin by describing the specific culture of the neural cells starting from human induced pluripotent stem cells (iPSCs), followed by the co-immunoprecipitation. We end with a detailed description of the analysis of proteomic data resulting from the previous steps. Adopting this strategy across a range of cell types can potentially offer further insights into the spatial organization and heterogeneity of the centrosome, extending the functional repertoire of this organelle. The approach can be further adapted to different sub-cellular targets or tissues, as well as other cell types including primary cells isolated from different species.
Materials and reagents
Human induced pluripotent stem cell lines, HMGU1 (hPSCreg: ISFi001-A) and HMGU12 (hPSCreg: ISFi002-A), obtained from the Helmholtz Center Munich iPSC Core Facility. The source of the cells is BJ fibroblasts (ATCC CRL-2522)
mTeSRTM 1 culture media (basal medium and 5× supplement) (StemCell Technologies, catalog number: 05850)
DMEM/F-12, GlutaMAXTM supplement (GibcoTM, catalog number: 10565018)
Neurobasal medium (1×) [-] L-Glutamine (GibcoTM, catalog number: 21103-049)
B-27 supplement with vitamin A (50×) (GibcoTM, catalog number: 17504044)
GlutaMAXTM supplement (100×) (GibcoTM, catalog number: 35050061)
Penicillin-Streptomycin (GibcoTM, catalog number: 15140122)
N2 supplement (100×) (GibcoTM, catalog number: 17502048)
Non-essential amino acids (100×) (GibcoTM, catalog number: 11140050)
2-mercaptoethanol (50 mM) (GibcoTM, catalog number: 31350010)
Insulin, human (10 mg/mL) (Sigma-Aldrich, catalog number: I9278-5ML)
Collagenase type IV (1 mg/mL) (StemCell Technologies, catalog number: 07909)
StemProTM AccutaseTM (GibcoTM, catalog number: A1110501)
Geltrex Matrix, reduced growth factor (GibcoTM, catalog number: A1413202)
Matrigel® growth factor reduced (Corning®, catalog number: 354230)
Laminin from Engelbreth-Holm-Swarm murine sarcoma basement membrane (Sigma-Aldrich, catalog number: L2020-1MG)
Poly-L-Ornithine hydrobromide (Sigma-Aldrich, catalog number: P3655-500MG)
Dorsomorphin (Sigma-Aldrich, catalog number: P5499-5MG)
SB431542 hydrate (Sigma-Aldrich, catalog number: S4317-5MG)
Recombinant human FGF-basic (Peprotech, catalog number: 100-18B)
ROCK inhibitor (Y-27632), 5 mg (StemCell Technologies, catalog number: 72304)
Nocodazole (Sigma-Aldrich, catalog number: M1404)
Dimethyl sulfoxide (DMSO), sterile-filtered (Sigma-Aldrich, catalog number: D2438)
Phosphate buffered saline (PBS) (10×) (GibcoTM, catalog number: 70011044)
Sterile water (Braun, catalog number: 0082423E)
MilliQ water (Merck) or distilled water (any brand)
Media filter: Filtropur V25, 250 mL, 0.2 μm (Sarstedt, catalog number: 83.3940.001)
Media filter: Filtropur V50, 500 mL, 0.2 μm (Sarstedt, catalog number: 83.3941.001)
Tris-base [NH2C(CH2OH)3] (Millipore, catalog number: 648310)
Sodium chloride (NaCl) (Sigma, catalog number: S3014)
Ethylenediaminetetraacetic acid tetrasodium salt (EDTA) (Sigma, catalog number: E6758)
Ethylene glycol-bis(2-aminoethylether)-tetraacetic acid (EGTA) (Sigma, catalog number: E4378)
Triton X-100 (Bio-Rad, catalog number: 1610407)
Sodium dodecyl sulfate (SDS) (Sigma, catalog number: L3771)
2-mercaptoethanol, molecular biology grade (Sigma-Aldrich, catalog number: M3148-25ML)
cOmplete, Mini Protease Inhibitor Cocktail (Roche, catalog number: 11836153001)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A2153)
Protein assay reagent A (Bio-Rad, catalog number: 5000113)
Protein assay reagent B (Bio-Rad, catalog number: 5000114)
Protein assay reagent S (Bio-Rad, catalog number: 5000115)
DynabeadsTM protein A (Invitrogen, catalog number: 10002D)
DynabeadsTM protein G (Invitrogen, catalog number: 10004D)
Micropipette tips (any brand)
6-well cell culture dishes (Falcon®, catalog number: 353046)
10 cm cell culture dishes (Falcon®, catalog number: 353003)
Cell scrapers (TPPTM, catalog number: 99002) (Sarstedt, catalog number: 83.3951)
50 mL Falcon tubes, 15 mL Falcon tubes (any brand)
Protein LoBind® Tubes, 1.5 and 2 mL (Eppendorf, catalog numbers: 0030108116 and 0030108132)
Protein LoBind® Tubes, 50 mL (Eppendorf, catalog number: 0030122240)
5, 10, and 25 mL serological pipettes (any brand)
Rabbit polyclonal anti-CDK5RAP2 (Sigma-Aldrich, catalog number: 06-1398)
Mouse polyclonal anti-CNTROB (Abcam, catalog number: ab70448)
Rabbit polyclonal anti-CP110 (Abcam, catalog number: ab99338)
Mouse monoclonal anti-CEP170 (InvitrogenTM, catalog number: 72-413-1/41-3200)
Rabbit polyclonal anti-CEP192 (Novus Biological, catalog number: NBP1-28718)
Rabbit polyclonal anti-CEP152 (Merck Millipore, catalog number: ABE1856)
Rabbit polyclonal anti-CEP63 (Merck Millipore, catalog number: 06-1292)
Rabbit polyclonal anti-CEP135 (Antibodies online, catalog number: ABIN2801434)
Rabbit polyclonal anti-POC5 (Novus Biological, catalog number: NBP1-78741)
Rabbit polyclonal anti-ODF2 (Abcam, catalog number: ab43840)
ROCK inhibitor (10 mM) (see Recipes)
Dorsomorphin (5 mM) (see Recipes)
SB431542 (10 mM) (see Recipes)
Poly-L-Ornithine, 10 mg/mL (see Recipes)
Neural maintenance medium (N3) (see Recipes)
Neural induction medium (N3+SMADi) (see Recipes)
Nocodazole (3.3 mM) (see Recipes)
2 M NaCl (see Recipes)
0.5 M EDTA, pH 7.5 (see Recipes)
0.5 M EGTA, pH 7.6 (see Recipes)
1 M Tris-HCl, pH 7.5 (see Recipes)
0.5 M Tris-HCl, pH 6.8 (see Recipes)
10% Triton X-100 (v/v) (see Recipes)
10% SDS (w/v) (see Recipes)
IP/wash buffer A (see Recipes)
Wash buffer B (see Recipes)
Protein standard (BSA) (64 mg/mL) (see Recipes)
1× Laemmli buffer (see Recipes)
Equipment
Inverted microscope (Leica, DM IL LED)
Neubauer improved cell counting chamber (Assistent, catalog number: 40442702)
Humidified cell culture incubator (any brand)
Brand Accu-jet pro pipette controller (Pro-lab, catalog number: 26330)
Microliter pipettes (Gilson®, P1000, catalog number: 10387322; P200, catalog number: 10327282; P20, catalog number: 10082012; P2, catalog number: 10635313)
Ice bucket (any brand)
Spectrophotometer or microplate reader (any brand with 750 nm wavelength)
DynaMag-2 magnet (Thermo Fisher Scientific, catalog number: 12321D)
1.5 mL vertical tube rotator (Stuart, catalog number: SB3)
Microcentrifuge with refrigeration (Thermo Fisher Scientific, catalog number: 75002446)
Microcentrifuge with refrigeration (Thermo Fisher Scientific, catalog number: 75004510)
Chemical fume hood (any brand)
Heating block (any brand with 95 °C capacity)
QExactive HF mass spectrometer (Thermo Fisher Scientific, Waltham, Massachusetts, USA)
Software
MaxQuant: https://www.maxquant.org/
Perseus: https://www.maxquant.org/perseus/
R Statistical Software: https://www.r-project.org/
Microsoft Excel: https://www.microsoft.com/de-de/microsoft-365/excel
Web resources
Ensembl Genome Browser: http://www.ensembl.org/index.html
Proteomics Identification Database (PRIDE): https://www.ebi.ac.uk/pride/
STRING Database: https://string-db.org/
Perseus documentation: http://www.coxdocs.org/doku.php?id=:perseus:start
Procedure
Table of contents and timeline:
Part I: Cell culture
A. Induced pluripotent stem cell culture media and plate preparation (0.5–1 day)
B. Induced pluripotent stem cell culture (3–5 days)
C. Induced pluripotent stem cell expansion (4–6 days × n)
D. Neural induction (40–42 days)
E. (Optional) Nocodazole treatment of the cells (4–5 h)
Part II: Protein isolation
F. Cell lysis for total protein isolation (2–4 h)
G. Assay protein concentration (0.5–1 h)
Part III: Immunoprecipitation
H. Immunoprecipitation (4–5 h)
I. Mass spectrometry (variable)
*The timeline may differ from the estimates we provide here depending on the researcher or availability of the resources.
General notes on cell culture and immunoprecipitation
Cell confluency is a critical element affecting proper neural induction and differentiation from iPSCs. The splitting ratios outlined in this protocol can be cell line dependent. In addition to the ratios mentioned, Table 1 outlines retrospective statistics on the range of seeding densities from different replicates of the successful differentiations using these splitting ratios. We recommend carrying out a small-scale pilot using 6-well plates in the initial instance using the splitting ratios and seeding counts outlined here as well as variations to this, such as 1:2 and 1:3, after day 10 of neural induction (Figure 1). CRITICAL: It is critical to keep the cell density above a certain level while the cells are still progenitors, i.e., still forming rosettes, to avoid their mis-direction towards other lineages.
Table 1. Example range of cell seeding densities used across different experiments that resulted in successful neural differentiation
* Harvested (per cm2) Seeded (per cm2)
Day 0 - 350,000–550,000
Day 10 1.2 Mio–1.7 Mio 630,000–730,000
Day 15 690,000–1.6 Mio 300,000
Day 21 820,000–1.4 Mio 420,000
Day 28 480,000–850,000 190,000–250,000
*Days since day 1 of neural induction (Mio: million)
It is advised to evaluate the cultures at different timepoints via immunostaining for marker proteins such as PAX6 (for neural stem cells); MAP2, CTIP2, SATB2, CUX1 (for neurons); and FOXG1 (forebrain).
Figure 1. Images of cells a few hours after splitting at different stages of neural differentiation. Scale bars: 200 μm.
Although a large amount of material is required to interrogate the full proteomic profile of the 10 baits, an advantage of this method is that the samples can be created in batches. That is, as long as all the proteomic samples across all replicates are run together on the mass spectrometer, not all of the 10 baits plus the negative controls need to be immunoprecipitated at the same time. For example, it is possible to carry out four immunoprecipitations of different baits plus a negative control in one batch, and the rest of the six immunoprecipitations plus a negative control in another batch for that cell type. The highly reproducible neural differentiation plus proteomic analysis across the four replicates makes this possible. However, if microtubule-dependent and microtubule-independent proteomic interactions are being interrogated for a particular bait (e.g., CEP120 in neural progenitors), the cells from the same batch of differentiation should be divided into nocodazole and DMSO treatments, and immunoprecipitated using the same antibody(s).
The protocol can easily be adapted to other cultured cell types, as well as primary cells isolated from different tissues or from different species, given the sufficient starting amount of isolated total protein for the co-immunoprecipitations can be obtained and relevant optimizations of the steps are done.
Unless specified otherwise, 1× PBS at room temperature is used for washing the cells.
It is advised to use pre-warmed media (37 °C) when splitting the cells and when changing the media daily.
Part I: Cell culture
Induced pluripotent stem cell culture media and plate preparation (TIMING: 0.5–1 day)
Prepare the medium:
When required, remove the mTeSR supplement (5×) from the freezer and thaw overnight at 2–8 °C prior to use.
Aseptically add 100 mL of mTeSR supplement (5×) to 400 mL of cold (2–8 °C) mTeSR basal medium. This mixture is named mTeSR medium hereafter.
Coat the plates:
Thaw Geltrex on ice and dilute 1:100 in pre-cooled DMEM/F-12 medium (at 4 °C) to avoid gelling the product; mix by turning the Falcon tube upside down a few times. Add 1 mL to each well of a 6-well plate, swirl the plate to distribute the solution evenly, and incubate in a humidified cell culture incubator for at least 2 h or overnight.
After incubation, remove the coating solution before seeding the cells (Optional: Wash the wells once with 1–2 mL of sterile 1× PBS). PAUSE POINT: If not using straight away, add 2 mL of 1× PBS per coated well, wrap the plate with parafilm, and store at 2–8 °C for up to one week.
Induced pluripotent stem cell culture (TIMING: 3–5 days)
Note: Before thawing the cells, have the Geltrex-coated plate(s), medium, ROCK inhibitor ready, and warm the water bath to 37 °C.
Prepare mTeSR medium supplemented with 10 μM ROCK inhibitor (1:1,000 dilution of 10 mM stock solution) sufficient for thawing and seeding the iPSCs as described below (hereafter referred to as mTeSR+ROCKi).
Pre-warm 3–4 mL of mTeSR+ROCKi to 37 °C in a 15 mL Falcon.
Rapidly thaw a vial of viable iPSCs frozen at a density of 2 × 105–1 × 106 (or one well of a 6-well plate) by placing the cryovial in a water bath at 37 °C for 1–2 min (until a tiny ice particle remains).
Using a 1 mL sterile pipette, transfer the cells into the pre-warmed 15 mL Falcon tube containing mTeSR+ROCKi. TIP: Disinfect the cryovial with 70% ethanol before opening.
Centrifuge at 300× g for 4 min at room temperature and remove the supernatant.
Resuspend the cells in 1.5–2 mL of mTeSR+ROCKi. Remove the Geltrex/PBS from the Geltrex-coated plate and add the cell suspension mixture to a single well of the 6-well plate (or up to three wells, with at least 1.5 mL final media in each).
Rock the plate side-to-side in both directions to evenly distribute cells in the well and incubate at 37 °C, 5% CO2.
The next day, remove the medium, wash the cells once with 1× PBS, add 1.5 mL of fresh mTeSR without ROCK inhibitor per well, and incubate at 37 °C, 5% CO2.
Replace the medium with 1.5 mL of fresh mTeSR every day until the cells are 80% confluent. Optional: When the cells are more than ~70% confluent, 2 mL of medium can be used instead.
It is good practice to observe iPSC lines under a phase contrast microscope daily to check for iPSC-like morphology, presence of differentiated cells, and confluency (Figure 2).
Figure 2. Induced pluripotent stem cells (iPSCs) at different growth phases. A) A few days after splitting. Cells might look larger, with loose cell-to-cell contacts. B) Semi-confluent stage. Center of the colonies has round cells with tight contacts, while the growing edges contain looser cells. C) With high confluency, most of the iPSCs become round and compact with tight cell-to-cell contacts. Scale bars: 200 μm.
Induced pluripotent stem cell expansion (TIMING: 4–6 days × n)
Notes:
Here, we describe splitting the cells from one well of a 6-well plate to six wells (1:6 splitting ratio). Size of the plates, volumes, and the ratio can be up- or down-scaled.
Have a 6-well plate coated with Geltrex ready on the day of splitting.
We describe here the passaging of the iPSCs using a combination of enzymatic and mechanical approaches. First, the wells are treated with Collagenase IV to soften the basement membrane, and then the cell clumps are collected using a cell scraper and further dissociated mechanically.
Dilute the Collagenase IV 1/4 in 1× PBS and keep aliquots in Falcons at -20 °C. Once thawed, working solution can be kept at 2–8 °C for a few weeks and should not be re-frozen.
For splitting the cells, pre-warm at least 1 mL of Collagenase and 9 mL of mTeSR to 37 °C. Volumes can be slightly increased to account for pipetting loss.
Aspirate the media from the well and add 2 mL of sterile 1× PBS. Swirl the plate a few times and remove the PBS.
Add 1 mL of Collagenase (1/4 in 1× PBS) to the well, distribute equally by swirling, and incubate the plate at 37 °C for 5 min (no more than 8 min, as the cells should not detach at this stage yet).
Aspirate Collagenase and add 1 mL of 1× PBS or mTeSR.
Scratch the well first with a 200 μL pipette tip at different directions to break cell colonies. Then, use a cell scraper to detach the cells from the plate completely.
Using a 1 mL pipette (or a serological pipette, depending on the number of wells) transfer the cells into a 15 mL Falcon and centrifuge at 300× g for 4 min at room temperature. Alternatively, keep the falcon at room temperature for 5-8 minutes for the cell clumps to precipitate to the bottom of the Falcon tube.
While cells are spinning, add 1 mL of mTeSR (pre-warmed to 37 °C) to each of the six Geltrex-coated wells.
Once cells have finished spinning, aspirate the supernatant and resuspend the cell pellet in 3 mL of fresh mTeSR, by pipetting the media up and down quickly 5–8 times. CRITICAL: Make sure to have neither single cells nor very large cell clumps. Clumps of 10–30 cells can be ideal. If the cells are too much dissociated by mistake, add ROCK inhibitor (10 μM final) for one day to help their survival. If the clumps are too large, iPSCs tend to differentiate in the middle of the colonies before the plate becomes confluent. To avoid large clumps, cells can be pipetted up and down additionally a few times before seeding to break them up further.
Add 0.5 mL of resuspended cell mixture to each of the six wells. Rock the plate side to side in both directions a few times and incubate the cells at 37 °C, 5% CO2. TIP: Avoid swirling the plate, as it will cause the cells to accumulate in the center of the well and start differentiating after a few days due to high density.
The next day, replace the media with 1.5 mL of fresh mTeSR, and repeat this step every day. Optional: When the cells are more than ~70% confluent, 2 mL of medium can be used instead.
Once at 80% confluency, split the cells again.
Continue expansion until having enough cells for neural induction.
Neural induction (TIMING: 40–42 days)
Note: Here, we describe a protocol for generating 60–100 × 10 cm dishes with neurons at day 40 of differentiation (Figure 3). This can account for 11 immunoprecipitations (IPs) for each of DMSO and nocodazole treatments (22 IPs in total, including negative controls) (Figure 4). Number of plates and IPs can be up- or down-scaled. For each 10 cm dish on day 0, grow 3–5 wells of 80% confluent iPSCs (Table 1). Along the differentiation, adjust the number of plates to proceed in each splitting according to the final number of plates desired. For neurons, calculate some additional plates to account for potential partial detachment of the cells, especially after day 35 of the differentiation.
Figure 3. Schematic representation of expansion and differentiation of induced pluripotent stem cells (iPSCs) towards neurons for co-immunoprecipitation of centrosome proteins with their interactors
Figure 4. Statistics (A) on the number of plates with day 40 neurons vs. total protein (milligrams) obtained in different sparse (B) or normal density (C) batches. Scale bars: 200 μm.
Dilute Matrigel 1:300 from an 8–12 mg/mL stock in DMEM/F-12 medium and add 4–4.5 mL per each of the two 10 cm dishes. Leave at 37 °C, 5% CO2 from at least 4 h to overnight.
CRITICAL: Do not let the coating dry on any part of the plate, as this alters the attachment of the cells and success of the induction. If part of the plate dries out, this might generate a heterogenous population of cells, which is impossible to sort out later. Therefore, adjust the coating volume carefully.
Before seeding the cells, remove the coating solution (Optional: Wash with 3 mL of 1× PBS).
PAUSE POINT: If the coated plates will not be used immediately, store at 4 °C after adding 6 mL of fresh 1× PBS up to two weeks.
Pre-warm Accutase and mTeSR medium to 37 °C.
Remove the media from 6-well plates containing iPSCs and wash once with 1× PBS.
Add 0.5–0.7 mL Accutase to 7–10 wells containing iPSCs (leave one well for re-splitting as per section C1) and incubate at 37 °C, 5% CO2 for 5 min (no longer than 8 min) (Optional: Swirl the plate once in the middle of the incubation). At the end of the incubation, cells should be suspended in solution mostly in single cell form.
After 5 min, add 1 mL of mTeSR or 1× PBS to each well containing Accutase and carefully collect and pool all the cells in a Falcon tube. Centrifuge at 300× g for 4 min at room temperature.
Remove the supernatant, being careful of the cell pellet. Resuspend the cells with 6 mL of mTeSR at 37 °C supplemented with 10 μM ROCK inhibitor (mTeSR+ROCKi).
Add 3 mL of cells to each of the 10 cm dishes pre-filled with mTeSR+ROCKi, to have a final volume of 10 mL.
From our experience, 3–4 confluent wells of a 6-well plate suffice for seeding a 10 cm dish.
Alternatively, count cells and seed 350,000–550,000 cells per cm2 of the 10 cm dish (Table 1) and adjust the final volume to 10 mL.
CRITICAL: It is important to have a dense cell layer of iPSCs on day 1 for proper neuroectoderm induction (Figure 5, Day1). With lower densities, the cells might assume neural crest-like or other lineages (Liu et al., 2015; Hindley et al., 2016).
Figure 5. Example images from different stages of successful differentiations. Cells were split on days 10, 15, 21, and 28. Days 11, 16, 22, and 29 represent the days after splitting. Day 15 and 40 are the timepoints used for the proteomic analysis in O’Neill et al. (2022). Scale bars: 200 μm.
After seeding, rock the plates from side to side in both directions a few times to homogenously distribute the cells after placing into the incubator (avoid swirling) and keep at 37 °C, 5% CO2 overnight.
The next day, if cells completely cover the entire plate (Figure 5, Day 1), begin neural induction. If not, replace with fresh mTeSR and incubate for another day. Do not keep the seeded cells more than two days before the induction.
The next day, remove the media, wash the cells once with 1× PBS, and replace with 10 mL of neural induction medium (N3+SMADi). This represents day 1 of neural induction.
Neural induction medium (N3+SMADi) is changed every day until day 10 of differentiation when a uniform neuroepithelial sheet should have formed.
On/before day 10, have the poly-L-ornithine/Laminin-coated plates ready, as follows:
Thaw a 10 mg/mL aliquot of poly-L-ornithine (PO). TIP: Keep the PO aliquot at 4 °C after thawing.
Coat four 10 cm dishes with poly-L-ornithine diluted 1:500 in sterile water. Incubate at 37 °C, 5% CO2 for at least 4 h. To be on the safe side, add at least 4.5 mL of coating solution per 10 cm dish to prevent partial drying out.
Following coating, remove the PO solution (optional: wash the plates once with 1× PBS) and coat the plates with Laminin diluted 1:200 in 1× PBS. Incubate at 37 °C, 5% CO2 for at least 4 h (or overnight). CRITICAL: Do not let the Laminin coating dry on any part of the plate, as this influences the density of attached cells, which is critical for the neural differentiation.
The same PO+Laminin coating procedure can be applied to sterile glass coverslips in 24-well plates to seed the cells for immunostaining.
PAUSE POINT: If the PO+Laminin-coated plates will not be used immediately, they can be stored after adding 6 mL of 1× PBS and sealed with parafilm at 2–8 °C for up to four weeks. Nonetheless, we recommend coating freshly before each splitting.
At day 10 of neural induction, split the cells 1:3. The exact splitting ratio can vary across cell lines but should be aimed at reaching 100% confluency on day 15 (Table 1, Figure 5).
Wash the cells with 1× PBS after removing the media, add 2 mL of Accutase per 10 cm dish, and incubate at 37 °C, 5% CO2 for 5 min.
Add 3 mL of neural induction medium (N3+SMADi) or 1× PBS to each plate, mechanically dissociate cells with a rubber (TPE) cell scraper, and then collect in a 50 mL Falcon.
Spin at 300× g for 4 min.
Remove the supernatant being careful of the cell pellet and re-suspend the cells with 18 mL of N3+SMADi supplemented with ROCK inhibitor (10 μM) (N3+SMADi+ROCKi) (assuming the cells were collected from 2 × 10 cm dishes).
Dispense cells according to a 1:3 ratio (in this case, 3 mL of cell suspension per 10 cm dish pre-filled with 5 mL of N3+SMADi+ROCKi) and incubate at 37 °C, 5% CO2. Optional: Count cells and seed according to the ranges provided on Table 1.
The next day (day 11), replace media with N3 medium without ROCK inhibitor. Replace the media again on day 12.
If the growing rate of cells do not forecast reaching to 100% confluency at day 15, supplement the media with 4 ng/ml FGF (1:2,500 dilution of 10 µg/ml stock) on day 11 or day 12, and keep with this medium for only one day. Avoid high doses or extended periods of culture with FGF, as it might caudalize the cells.
From day 12 on, replace the medium with fresh N3 every other day (unless FGF is added; if so, adjust the media changing days accordingly).
On day 15, cells should be 100% confluent with the appearance of rosettes (Figure 5).
TIP: If the rosettes are not readily visible, try to observe them while washing the cells with PBS or after changing the media.
If the cells were also seeded on the coverslips, fix them with PFA (to stain for cell identity markers, such as PAX6) or methanol (to stain for most centrosome proteins).
On/before day 15, have the poly-L-ornithine/Laminin-coated plates ready and split the cells as described for day 10 from section D12 onwards.
From this stage on, replace the media every second day (8 mL of N3 per 10 cm dish). Always replace the media the day after splitting the cells, preferably after washing the cells once with 1× PBS.
Split the cells also on days 21 and 27/28 (for approximate cell counts, refer to Table 1) and proceed with media changes until day 40 (Figure 3). TIP: From approximately day 35 on, be extra careful when changing the media, as the cells tend to detach locally or as an entire sheet from the plate. To avoid this, either add the medium very slowly using a 25 mL pipette dropwise or change only half of the media, but daily.
Observe the morphology of the cells along the differentiation to see if they assume morphologies typical of each stage (Figures 5 and 6).
Figure 6. Example images from failed differentiations. Cells might assume cobblestone or spiny/fibroblast-like morphology, have large and/or darker nuclei, or enlarged cytoplasm. Cultures should be monitored frequently (e.g., after each media change) to watch out for such aberrant morphologies, which might signal potential failure of the differentiation. Scale bars: 200 μm.
(Optional) Nocodazole treatment of the cells (TIMING: 4–5 h)
Notes:
Nocodazole is a drug that blocks polymerization of the microtubules and can be used to assess microtubule-independent centrosome interactions of the proteins. To this end, nocodazole and DMSO (solvent, as control) are added to their respective culture plates 4 h before harvesting the cells for protein isolation and immunoprecipitation (Figure 7).
Figure 7. Day 40 neurons after 4 h of nocodazole treatment. A) Nocodazole-treated cells B) DMSO-treated control cells. Scale bars: 200 μm.
Because nocodazole increases cell death and reduces protein yield, plates can be divided into nocodazole and DMSO treatments with approximately 60:40 ratio to account for this loss.
For microtubule depolymerization, alternative agents such as vinorelbine, vincristine, colchicine (Bates and Eastman, 2017), or cold exposure (Li and Moore, 2020) can also be considered and tested to see if they result in reduced cell death.
Add nocodazole (3.3 mM stock concentration, dissolved in DMSO) to a final concentration of 3.3 μM to the N3 medium (1:1,000). Swirl plate to distribute it evenly.
As a control, DMSO is added to its respective plates using the same ratio added to the nocodazole plates (i.e., 1:1,000).
Incubate the plates at 37 °C, 5% CO2 for 4 h, confirm the de-polymerization under the microscope, and begin collecting the cells for lysis and protein isolation.
Part II. Protein isolation
Cell lysis for total protein isolation (TIMING: 2–4 h)
Notes:
On the day of immunoprecipitation (IP), freshly make up IP/wash buffer A and wash buffer B and store on ice for the duration of the procedure.
Cool centrifuge for spinning 50 mL Falcons to 4 °C.
Pre-cool 1× PBS to 4 °C and keep in the ice bucket (in case of nocodazole treatment).
Obtain plates from incubator and aspirate the media. Wash cells with 3 mL of 1× PBS per plate. Aspirate PBS and add 3 mL of fresh 1× PBS.
TIP: If handling a lot of plates, you can split them to batches, keeping the rest in the incubator.
For nocodazole-treated plates, wash with ice-cold PBS, keep plates on ice in bucket while scraping, and process only a couple of plates at a time acting quickly to impede repolymerization of microtubules. TIP: To keep the conditions as similar as possible, DMSO-treated plates can be treated the same way.
Using a cell scraper, scrape the cells off the plate, tilting the plate sideways and pushing all the cells into the PBS. Keep the plate tilted and, using a 10 mL pipette, transfer cells in PBS to a 50 mL Falcon placed on ice.
Continue to scrape cells and transfer to 50 mL Falcon tube(s), pooling the cells within their respective treatments.
Once all plates are scraped (or the 50 mL Falcon is filled), spin at 300× g for 5 min at 4 °C to pellet cells.
Aspirate off the PBS (ensure the pellet is not removed).
PAUSE POINT: If cell lysis will not be done on the same day, cell pellets can be snap-frozen in liquid nitrogen and kept at -80 °C. If this is the case, add the IP/wash buffer A directly on the frozen pellet without thawing it on the day of lysis.
Assess the volume of the pellet in milliliters and add double that volume of IP/wash buffer A.
Incubate on ice for 30 min.
It is important not to dilute the sample too much, as this will affect the protein concentration and subsequent downstream steps. For example, 5 mg of total lysate in 1 mL will be used for immunoprecipitation; therefore, the concentration should not be lower than this.
During the cell lysis, cool a micro-centrifuge capable of spinning 1.5 mL tubes at 18,000× g to 4 °C.
After the 30 min incubation, once the cells are lysed, distribute lysate into 1.5 or 2 mL protein low bind tubes and spin at 18,000× g for 10 min at 4 °C to pellet the cellular debris.
During this time, set up the protein standards for the standard curve as described below.
After the 10 min centrifugation and the cellular debris is pelleted, carefully remove the clear supernatant from the 1.5 mL tubes and pool into a (Optional: protein low bind) 15 mL Falcon (CRITICAL: do not mix different samples), mix well, and proceed with the next step. TIP: It can be practical to calculate the total volume while pooling the lysates for a more accurate estimation of total protein later.
PAUSE POINT: If the cell lysates will not be used immediately, they can be kept at -80 °C.
Assay protein concentration (TIMING: 0.5–1 h)
Notes:
Prepare the dilutions and protein standards on ice.
Adding the reagents to the microplate and subsequent steps can be done at room temperature.
To set up the protein standards, add 20 μL of IP/wash buffer A to six individual tubes (labeled as 8, 4, 2, 1, 0.5, 0.25). Add 20 μL of the 16 mg/mL BSA protein standard (see Recipes) to the first tube (8 mg/mL), performing a 1:2 dilution series with the remaining tubes, down to 0.25 mg/mL.
Example: mix [20 μL of buffer A + 20 μL of standard], transfer 20 μL of this mixture to the next tube that already contains 20 μL of buffer A, mix well, transfer 20 μL of this mixture to the next tube, and so on.
In a fresh tube, make a master mix with a 49:1 ratio of Reagent A and Reagent S (A+S) of the Bio-Rad Protein Assay:
Calculate two replicate wells per four standards (2 mg/mL, 1 mg/mL, 0.5 mg/mL, and 0.25 mg/mL), one for control, two replicates per sample to be measured, and two extras (Figure 8). Example:
A+S mix = [11 + (2 × Nsamples)] × (24.5 μL Reagent A + 0.5 μL Reagent S)
Figure 8. Example layout of a microplate section with protein standards and diluted samples for determination of protein concentration with DC Protein Assay
Add 25 μL of the A+S mix to the wells of the protein reader plate.
Add 5 μL of each protein standard to two corresponding wells each, as well as the 5 μL of IP/wash buffer-only control to its respective well (Figure 8).
Add 5 μL of the cell lysate per sample to their respective wells containing the A+S mix.
If the protein concentration is too high, the measurement will fall above the linear range. In this case, two different dilutions of the samples can be used for measurement, e.g., 1/4 and 1/10, or else, to ensure accurate determination of concentration. Try to make sure at least one of your sample dilutions fall within the range of 2–0.25 mg/mL.
Aliquot 200 μL of Reagent B to each well to be measured, going from the lowest-to-highest protein concentration wells, and incubate at room temperature for 10–15 min.
Measure the absorbance of the samples at 750 nm within one hour and use the protein standard measurements to produce a standard curve using linear regression (y = mx + n) where y is absorbance and x is protein concentration. Then, calculate the amount of protein within your sample(s) of interest based on its absorbance using this formula.
TIP: It is not recommended to exceed 2 mg/mL for the standards, as the linear range can be lost for higher values than this. Calculate the R2 value of the fitted line to check its linearity.
PAUSE POINT: If the cell lysates will not be used immediately, they can be kept at -80 °C.
Part III: Immunoprecipitation
Immunoprecipitation (TIMING: 4–5 h)
Note: At least four replicate immunoprecipitations performed using different biological replicates are recommended for each bait of interest.
Calculate how many immunoprecipitations can be performed with the total lysate obtained from cells (Figure 4).
Each immunoprecipitation for mass spectrometry requires 4–5 mg of starting total protein, while samples for assessing the immunoprecipitation by Western blot can be as little as 500 μg.
Each immunoprecipitation is performed in 1 mL of volume.
Dilute the protein lysate to 5 mg/mL in IP/wash buffer A and aliquot 1 mL of lysate into each respective protein low bind tube for immunoprecipitation. Keep the samples on ice.
For every set of immunoprecipitations, one of the samples should be reserved for no-antibody control.
As an alternative to the no-antibody control, it is encouraged to use isotype-specific control antibody(s) as negative control that matches the class and type of the primary antibodies used in the experiment.
For each immunoprecipitation, add 2 μg of the desired antibody to 1 mL of lysate at 5 mg/mL. For the negative control, add no antibody to the tube or add the isotype control. CRITICAL: From this point on, be very careful about cross-contamination of the samples. Never use the same tips for different samples during washes and avoid splashes across the tubes, as mass spectrometry has a very-high detection sensitivity.
Incubate tubes on an end-to-end rotator in a 4 °C cold room for 1 h. Fifteen minutes before the end of this incubation, prepare the Protein A and Protein G Dynabeads solution:
Mix 10 μL of Protein A and 10 μL of Protein G per immunoprecipitation (including the negative control) in a 1.5 mL tube. Example:
A+G mix = (1 + NIPs) × (10 μL Protein A + 10 μL Protein G)
Place tube on a magnetic strip and remove the liquid without disturbing the beads with the pipette tip.
Resuspend the beads with IP/wash buffer A using the same volume aspirated from the tube, place the tube on the magnetic strip, and remove the liquid.
Repeat this wash two more times and finally re-suspend the beads with IP/wash Buffer A using the original volume.
After 1 h incubation of the lysate with the antibodies, add 20 μL of Protein A+G mix to each IP tube as well as to the negative control. Incubate on the rotator for an additional 2 h at 4 °C.
After this incubation period, place all the IP tubes on the magnetic strip. Remove the wash buffer, leaving the beads coupled to the antibodies and protein attached to the wall of the tube. Add 1 mL of IP/wash buffer A.
CRITICAL: Use a clean tip for each sample to remove the supernatant.
CRITICAL: Work very quickly or in batches to avoid drying of the beads if there are too many samples.
Incubate the samples on a rotator at 4 °C for 15 min and repeat the above step two more times (each with a 15 min incubation).
During this process, pre-heat a heating block to 95 °C.
Under a chemical hood, mix 1× Laemmli buffer and 2-mercaptoethanol in a 39:1 ratio (e.g., 975 μL of 1× Laemmli buffer + 25 μL of 2-mercaptoethanol). Approximately 30–40 μL is required per IP.
After the third wash, place the samples on the magnetic strip and remove the wash buffer. Add 1 mL of wash buffer B to each sample before incubating on a rotator at 4 °C for a couple of minutes. Repeat this step once again.
After these washes, place the samples on the magnetic strip and remove the wash buffer. TIP: 10/20 μL tip can be used following the 1,000 μL tip to remove the supernatant as much as possible. Place the tubes with the beads immediately on ice and transfer to a fume hood.
Add 30–40 μL of fresh Laemmli buffer containing 2-mercaptoethanol to each sample and ensure the beads are well re-suspended. The volume can be decided on a case-by-case basis, but it is important to keep it consistent across different samples and batches.
Incubate the tubes for 10 min at 95 °C. After this time, place the tubes on the magnetic strip in the fume hood and transfer the supernatant (Laemmli buffer now also containing the immunoprecipitated protein of interest and its interactors) to a new protein low bind tube without disturbing the beads.
Store samples at -80 °C until all immunoprecipitations are completed. CRITICAL: Run all samples together on the mass spectrometer, as run-to-run variations are common even if the same samples are re-analyzed at a different timepoint.
Mass spectrometry
LC-MS/MS analysis of the eluates from the co-immunoprecipitations was performed on a QExactive HF mass spectrometer, as described in O’Neill et al. (2022).
Proceed to the MaxQuant section once the .RAW files from mass spectrometry are available.
Data analysis
Table of contents and timeline:
Part I: MaxQuant analysis
A. Processing the mass spectrometry. RAW files using MaxQuant (3–10 days)
Part II: Perseus analysis
B. Pre-processing, annotation, and quality control (0.5–2 days)
C. Statistical analysis (2–7 days)
Part III: Further analysis
D. Combine and summarize the data (0.5 day)
E. Further analysis suggestions (variable)
*The timeline may differ from the estimates we provide here depending on the researcher or availability of the resources.
General notes on data analysis
The data analysis section contains the following main parts (Figure 9):
Processing the mass spectrometry .RAW files to generate intensities (MaxQuant)
Pre-processing, annotation, and quality control (Perseus & R statistical software)
Importing the MaxQuant output into Perseus
Annotating the samples
Quality filtering
[Optional: Adding pathway annotations (e.g., KEGG pathway, GO terms)]
Cluster analysis for quality control (Perseus & R statistical software)
Statistical analysis (Perseus & Excel)
t-test (volcano plot)
Ranking the proteins for statistical significance
Combining and summarizing the data (R statistical software)
Network analyses
Intermediate files generated along this protocol and R scripts are provided in the Supplementary information.
Figure 9. Overview of the analysis pipeline
Part I: MaxQuant analysis
Processing the mass spectrometry .RAW files using MaxQuant (TIMING: 3–10 days)
Notes:
Timing might differ depending on the processing power of the computer, number of samples, and the parameters.
Resulting .RAWfiles from the mass spectrometer were processed using MaxQuant software to generate intensities (Tyanova et al., 2016). Please note that MaxQuant can currently run only on Windows operating systems.
Proteins were identified by searching against the Uniprot (SwissProt, reviewed) database for Homo sapiens (taxon identifier: 9606, 15th June 2021, canonical sequences and isoforms), which is provided in the Supplementary information: MaxQuant Analysis Files.
The most important MaxQuant parameters that differ here from the default values are as follows:
i. Matching between runs was enabled, but only among the replicate samples.
ii. iBAQ and LFQ Intensities were calculated.
iii. Deamidation (NQ) was included in the modifications that are used for protein quantification.
iv.TIP: mzTab is written (Tables section), which is required to do a complete upload of the MaxQuant results to the PRIDE server.
v. Other parameters can be checked in more details from the mqpar.xml file provided in the MaxQuant Analysis Files.
Place all the .RAW files into the same folder.
For this protocol, a subset of the .RAW files from O’Neill et al. (2022), deposited to the ProteomeXchange Consortium through the Proteomics Identification Database (PRIDE) with the accession number PXD031936, were used (https://doi.org/10.6019/PXD031936).
i. Go to the page: https://www.ebi.ac.uk/pride/archive/projects/PXD031936.
ii. Download the following .RAW files navigating to the bottom of the page, either using the search box for individual files or using the FTP link:
1) Negative controls:
a) UZF13667X004__7_CD7_DMSO_ctrl_92-229116.raw
b) UZF13667X003__5_CD9_DMSO_ctrl_92-229115.raw
c) UZF13667X002__1_CD11_DMSO_ctrl_92-229114.raw
d) UZF13667X006__29_CD12_DMSO_ctrl_92-229118.raw
e) UZF13667X007__40_CD13_DMSO_ctrl_92-229119.raw
f) UZF13667X005__18_CD14_DMSO_ctrl_92-229117.raw
2) CEP63 IPs
a) UZF13667X010__8_CD7_DMSO_Cep63_92-229122.raw
b) UZF13667X009__2_CD11_DMSO_Cep63_92-229121.raw
c) UZF13667X012__30_CD12_DMSO_Cep63_92-229124.raw
d) UZF13667X011__19_CD14_DMSO_Cep63_92-229123.raw
3) CEP170 IPs:
a) UZF13667X025__11_CD7_DMSO_Cep170_92-229137.raw
b) UZF13667X024__6_CD9_DMSO_Cep170_92-229136.raw
c) UZF13667X027__33_CD12_DMSO_Cep170_92-229504.raw
d) UZF13667X028__41_CD13_DMSO_Cep170_92-229505.raw
e) UZF13667X026__22_CD14_DMSO_Cep170_92-229503.raw
CRITICAL: Make sure that there is at least the same amount of empty disc space as the total size of the .RAW files to be analyzed before starting the run.
Download the latest version of the MaxQuant software from https://www.maxquant.org/download_asset/maxquant/latest, place in the same folder with the .RAW files, and unzip. MaxQuant version 1.6.17.0 used for this publication is provided in the Supplementary information.
Initiate the software by double-clicking on MaxQuant.exe (no installation is required).
File > Load parameters… > select the mqpar.xml (MaxQuant Analysis Files) that was placed in the same folder as the .RAW files.
When working on another dataset, remove the current files in the template and replace by the new set of .RAW files:
Select all the existing files by right-clicking on the data > Select all.
Raw data > Input data > Remove.
Raw data > Input data > Load.
Select all the .RAW files for the current analysis.
Experimental design parameters (Figure 10) can be assigned either manually using the menu items above [e.g., Set experiment, Set fractions (see BOX1)] or using Excel as follows:
Figure 10. Example experimental design for a MaxQuant run
BOX1: Matching between runs
In cases where some of the peptide sequences could not be identified from the MS/MS spectrum due to insufficient information, missing values can be compensated by matching the run with another, to still get identified features (Tyanova et al., 2016). Matching should be done only between similar samples and the MaxQuant algorithm matches only between the same or adjacent fractions.
TIP: Because it is not expected to have a very similar set of proteins pulled down by different antibodies in co-IPs, unlike the total proteome, we aimed to match only replicate samples. This is done by a trick to assign the biological replicate IPs to the same fraction and skipping a number for the next set of IPs, to minimize false positives, while facilitating recovery of unidentified peaks using replicate IPs (Figure 10).
Manual setup:
i. Assign fractions.
1) Select the replicate IPs by Ctrl+left-click.
2) Raw data > Edit experimental design > Set fractions (Figure 11).
Figure 11. Assigning fractions to a sample (write the same number to both fields)
ii. Assign experiment names:
Select the sample > Raw data > Edit experimental design > Set experiment > and give a unique experiment name to each sample.
Alternatively, assign parameters in bulk using Excel:
i. Raw data > Experimental design file > Write template.
ii. Find the file: experimentalDesignTemplate.txt in the >combined folder and open with Excel.
iii. Assign a unique experiment name to each sample (e.g., Text-to-column function can be used to generate them from sample names). Assign the same fraction number to replicate samples and skip at least one number for the next set of replicates (e.g., all controls: 1, all CEP63 IPs: 3) (see “Fraction” in Figure 10).
iv. Save the file (as .txt) and upload via Raw data > Experimental design file > Read from file > locate the experimentalDesignTemplate.txt (MaxQuant Analysis Files).
Provide the sequence database to be used for peptide identification.
Global parameters > Parameter section > Sequences.
Fasta files: Add > Locate the .fasta file to be used:
The reviewed human protein reference can be retrieved from UniProt database as follows:
i. Go to https://www.uniprot.org > Proteins.
ii. Select Human and Status: “Reviewed” (Swiss-Prot).
iii. Download (all) > Format: FASTA (canonical & isoform).
Taxon identifier: 9606 (Homo sapiens), canonical sequences and isoforms (reviewed), 15th June 2021 was used for the current protocol (MaxQuant Analysis Files: 2021_02_Human_canonical_and_isoforms.fasta).
Select the uploaded file on the software (Fasta file path) and select > Identifier rule > UniProt identifier: >.*\|(.*)\|
Click Start at the bottom of the software.
Depending on the experiment size and the computing power, the run can take from a few days to more than a week.
The results can be found in the “combined” folder that is placed in the same folder with .RAW files. combined > txt > proteinGroups.txt will be used for the subsequent Perseus analysis.
Part II: Perseus analysis
Pre-processing, annotation, and quality control (TIMING: 0.5–2 days)
Notes:
Protein interactors of the baits were determined by processing the MaxQuant result table ProteinGroups.txt in Perseus software (Tyanova et al., 2016). Tutorial pages and videos can be found at http://coxdocs.org/doku.php?id=perseus:start. Please note that Perseus can currently run only on Windows operating systems.
Perseus software generates a new matrix for each step; therefore, always apply the next operation to the latest matrix generated.
It is possible to re-color or re-name a matrix by right-clicking on it, which would be helpful especially for long workflows.
Matrices can be deleted using the red cross mark at the top of the section where the workflow tree is.
The workspace can be saved at any point and can be resumed later.
CRITICAL: The .sps files generated in one version of Perseus cannot be processed using another version. Therefore, we strongly recommend to always record the version number and keep a copy of the software together with the analysis files.
TIP: It is advised to save the Perseus file frequently to avoid data loss due to occasional software crash.
Download the Perseus software.
Latest version can be found under: https://maxquant.net/download_asset/perseus/latest. Perseus version 1.6.14.0 (Supplementary information) was used in the original research described in O’Neill et al. (2022) and for the current protocol.
Initiate the software by double-clicking on Perseus.exe (no installation is required) [Perseus Analysis Files: 1_ExampleAnalysis_Part1.sps]
Import the MaxQuant results table into Perseus:
For this protocol, the following example dataset was used:
i. Negative controls:
1) UZF13667X004__7_CD7_DMSO_ctrl_92-229116.raw
2) UZF13667X003__5_CD9_DMSO_ctrl_92-229115.raw
3) UZF13667X002__1_CD11_DMSO_ctrl_92-229114.raw
4) UZF13667X006__29_CD12_DMSO_ctrl_92-229118.raw
5) UZF13667X007__40_CD13_DMSO_ctrl_92-229119.raw
6) UZF13667X005__18_CD14_DMSO_ctrl_92-229117.raw
ii. CEP63 IPs
1) UZF13667X010__8_CD7_DMSO_Cep63_92-229122.raw
2) UZF13667X009__2_CD11_DMSO_Cep63_92-229121.raw
3) UZF13667X012__30_CD12_DMSO_Cep63_92-229124.raw
4) UZF13667X011__19_CD14_DMSO_Cep63_92-229123.raw
iii. CEP170 IPs:
1) UZF13667X025__11_CD7_DMSO_Cep170_92-229137.raw
2) UZF13667X024__6_CD9_DMSO_Cep170_92-229136.raw
3) UZF13667X027__33_CD12_DMSO_Cep170_92-229504.raw
4) UZF13667X028__41_CD13_DMSO_Cep170_92-229505.raw
5) UZF13667X026__22_CD14_DMSO_Cep170_92-229503.raw
Click on Matrix > Load > Generic matrix upload (green arrow pointing upper left).
File: Select > locate the MaxQuant results folder: combined > txt > proteinGroups.txt [Perseus Analysis Files: 2_proteinGroups.txt]
Include columns by selecting the items on the left menu and transferring to the respective category on the right using arrows. TIP: Holding down Ctrl button and/or left clicking and pulling down allows multiple selection. The order of items within the category can be changed using up and down arrows on the right side. We included columns we marked as “optional” in the initial upload, and then removed, in case some parameters must be retrospectively checked during the analysis, to avoid starting all over again. [ExampleAnalysis_Part1: Full Matrix]
i. Main (experimental quantitative results)
1) LFQ intensity (for each sample)
2) iBAQ (for each sample)
3) Intensity (for each sample)
ii. Numerical
1) Peptides
2) Razor + unique peptides
3) Unique peptides
4) Optional: Sequence coverage [%]
5) Optional: Unique + razor sequence coverage [%]
6) Optional: Unique sequence coverage [%]
7) Mol. weight [kDa]
8) Q-value
9) Score
10) Optional: Intensity
11) Optional: iBAQ peptides
12) MS/MS count
13) Number of proteins
14) Optional: Unique peptides (for each sample)
15) Optional: Peptides (for each sample)
16) Optional: Razor + unique peptides (for each sample)
17) Optional: MS/MS count (for each sample)
iii. Categorical
1) Only identified by site
2) Reverse
3) Potential contaminant
4) Optional: Identification type (for each sample)
iv. Text
1) Protein IDs
2) Majority protein IDs
3) Protein names
4) Gene names
5) Fasta headers
v. Multi-numerical
1) Peptide counts (all)
2) Peptide counts (razor+unique)
3) Peptide counts (unique)
Remove optional columns to avoid large file size during subsequent analyses [ExampleAnalysis_Part1: Smaller Matrix].
i. Left-click on the matrix and select on the menu: Matrix > Processing > Rearrange > Reorder/remove columns.
ii. Select the items on the right menu and transfer the respective category back to the left.
Add different categories to the samples to be able to group them for statistical analysis.
It is possible to add annotation rows manually:
i. Matrix > Processing > Annot. rows > Categorical annotation rows.
ii. Action: Create.
iii. Row name: Assign a name to the category.
iv. Add categories to each sample.
Alternatively, add all annotations at once for a large number of samples and multiple categories, editing the template file in Excel:
i. First, create a mock annotation row following the steps above (this is required for exporting the template file) [ExampleAnalysis_Part1: Placeholder Category].
ii. Then, export to a matrix:
1) Matrix > Processing > Annot. rows > Extract to matrix [ExampleAnalysis_Part1: Placeholder Annotation].
2) Select the new matrix and Matrix > Generic matrix export (upper rightmost, floppy disc icon) > click on the Select button to choose the folder, assign a file name, and click OK.
iii. Open the file in Excel and create a new column (letter “C” should be assigned as the column type, meaning “Categorical”) for each category to be added (TIP: Original column names can be chopped using Data Tools > Text to Columns and combined in different ways to generate categories) (Table 2). Here, we generated the following categories:
1) Experiment: DMSO_ctrl_CD7, DMSO_CEP63_CD11, …
2) Measurement: LFQ, iBAQ, Intensity
3) IP: ctrl, CEP63, CEP170
4) Replicate: CD7, CD9, CD11, CD12, CD13, CD14
5) IP_Rep: ctrl_CD7, ctrl_CD9, CEP63_CD7, …
Table 2. An example annotation file
Experiment Measurement IP IP_Rep Name
#!{Type}C C C C T
DMSO_ctrl_CD7 LFQ ctrl ctrl_CD7 LFQ intensity CD7_DMSO_ctrl
DMSO_CEP63_CD7 LFQ CEP63 CEP63_CD7 LFQ intensity CD7_DMSO_Cep63
DMSO_CEP170_CD12 LFQ CEP170 CEP170_CD12 LFQ intensity CD12_DMSO_Cep170
DMSO_ctrl_CD7 iBAQ ctrl ctrl_CD7 iBAQ CD7_DMSO_ctrl
DMSO_CEP63_CD7 iBAQ CEP63 CEP63_CD7 iBAQ CD7_DMSO_Cep63
DMSO_CEP170_CD12 iBAQ CEP170 CEP170_CD12 iBAQ CD12_DMSO_Cep170
DMSO_ctrl_CD7 Intensity ctrl ctrl_CD7 Intensity CD7_DMSO_ctrl
DMSO_CEP63_CD7 Intensity CEP63 CEP63_CD7 Intensity CD7_DMSO_Cep63
DMSO_CEP170_CD12 Intensity CEP170 CEP170_CD12 Intensity CD12_DMSO_Cep170
iv. Rename the cell “Column name” as “Name” (otherwise Perseus generates an error) and save as a .txt file [Perseus Analysis Files: 3_Categories.txt].
v. Go back to Perseus, select the last matrix before exporting annotations (Smaller Matrix), and go to Matrix > Processing > Annot. rows > Categorical annotation rows > Action: Read from file > Input file: locate the saved annotation file using the Select button [ExampleAnalysis_Part1: Added: Categories].
Filter the annotated matrix based on MaxQuant quality parameters.
Matrix > Processing > Filter rows > Filter rows based on categorical column (Figure 12).
Figure 12. Filtering rows based on a categorical column
i. Column: Only identified by site (these proteins did not go through proper FDR, because they were identified by only peptides with modified amino acids).
1) Values: + (on the right side).
2) Mode: remove matching rows.
3) Filter mode: Reduce matrix.
4) Click OK.
ii. Column: Reverse (sequences found in reverses decoy database, which is used to calculate the FDR, i.e., statistical cutoff for acceptable spectral matches).
1) Values: + (on the right side).
2) Mode: remove matching rows.
3) Filter mode: Reduce matrix.
4) Click OK.
iii. Column: Potential contaminant (proteins that are commonly occurring contaminants in the mass spectrometry data).
1) Values: + (on the right side).
2) Mode: remove matching rows.
3) Filter mode: Reduce matrix.
4) Click OK.
Optional: Check whether anything filtered out looks important using “Split matrix” instead.
Filter out immunoglobulins (these can come from the antibodies used for co-immunoprecipitation):
Matrix > Processing > Filter rows > Filter rows based on text column > Column: Fasta headers
i. Search string: “immunoglobulin.”
ii. Deselect Match whole word.
iii. Mode: Remove matching rows.
iv. Filter mode: Reduce matrix.
v. Click OK.
Optional: Check whether anything filtered out looks important using “Split matrix” instead.
Optional: Add pathway annotations (e.g., KEGG pathway, GO terms); this can also be done later.
Download the annotations:
i. Tools > Annotation download >http://annotations.perseus-framework.org> PerseusAnnotation
1) > FrequentlyUsed > mainAnnot.homo_sapiens.txt.gz.
2) Can also be found under: > OrganismSpecific > h > scroll down to find Homo sapiens.
ii. Download and place the file (no need to extract) into the folder location: Perseus 1.6.14.0 > bin > conf > annotations.
Add the annotations:
Matrix > Processing > Annot. columns > Add annotation.
i. Source: select mainAnnot.homo_sapiens.txt.gz (it should now be automatically listed).
ii. UniProt column: Protein IDs.
iii. Add the annotations of your interest, e.g.:
1) GOBP name
2) GOMF name
3) GOCC name
4) KEGG name
5) Corum
Export the matrix as a backup before the next step of the analysis: Matrix > Export > Generic Matrix Export > AfterBasicFilters.txt.
Correct the missing protein names or gene symbols that might be converted to date because of opening the file in Excel (e.g., SEPTINs, MARCs) (Figure 13).
Figure 13. Pay attention to the protein names; these might be converted to dates when the files are processed in Excel
Further potentially problematic protein names include: CALM3 (Calmodulin-3, P0DP25), PBX3, CV015, TMM70, CC201, CRCC2, TPM1, TPM3, MIC60, MACF1, ZN683, HNF4A, ASCC1, and SREK1 (e.g., missing protein names or gene names).
After correcting, save as .txt file [Perseus Analysis Files: 4_AfterBasicFilters.txt].
Cluster analysis (using Perseus and R software):
Perseus
i. Upload the corrected AfterBasicFilters.txt file: Matrix > Load > Generic matrix upload.
ii. Pre-processing:
1) Transform the data [ExampleAnalysis_Part1: log2 (x+1)]: Log2-transformation: Matrix > Processing > Basic > Transform
a) Transformation: log2(x+1).
b) Columns: select all the main columns.
2) Subset the LFQ Intensities: Matrix > Processing > Rearrange > Reorder/remove columns.
3) Transfer the iBAQ and Intensity columns back to left side and keep as the main columns only LFQ Intensities [ExampleAnalysis_Part1: LFQ]
iii. Filter out the proteins detected in less than two samples in total:
1) Matrix > Processing > Filter rows > Filter rows based on valid values.
2) Min. valids > Number: 2.
3) Mode: In total.
4) Values should be > Greater than: 0.
5) Filter mode: Reduce matrix.
iv. PCA Analysis (Figure 14): Matrix > Analysis > Principal component analysis (the symbol with red and blue dots).
Figure 14. PCA analysis (LFQ Intensities)
1) Labels can be made visible from: Points tab > Right-click on the data > Select all > Show labels (gray label sign on top). Label text can be changed using the menu item on top of the plot.
2) Replicates can be colored using the menu in the Points and Categories tabs of the PCA.
3) Symbol type, size, and color can be modified.
v. Hierarchical clustering (Figure 15): Matrix > Analysis > Hierarchical clustering.
1) Default settings can be used.
2) Appearance of the heatmap can be adjusted using the menu items on the Heat Map tab (size, labels to show, colors etc.) (Figure 16).
Figure 15. Hierarchical clustering of the samples in the current analysis (LFQ Intensities)
Figure 16. Appearance of the heatmap can be adjusted using menu items in the menu
Dimensionality reduction analysis using R statistical software:
i. Export the matrix filtered for PCA analysis: Matrix > Export > Generic Matrix Export > Select: LFQ.txt [Perseus Analysis Files: 5_LFQ.txt].
ii. Refer to the sample R script 6_Cluster_Analysis.R (Perseus Analysis Files).
iii. Run tSNE and UMAP analysis using R (Figure 17).
Figure 17. tSNE and UMAP visualization (LFQ Intensities)
Statistical analysis (TIMING: 2–7 days, depending on the number of samples)
Note: Here, we provide one approach among many possibilities for statistical analysis; alternative tests or methods can also be used for evaluation of the enriched proteins.
Create a separate .sps file per bait, starting with the AfterBasicFilters matrix. In this example, we generated the following two files:
Perseus Analysis Files: 8_ExampleAnalysis_Part2_CEP63.sps.
Perseus Analysis Files: 9_ExampleAnalysis_Part2_CEP170.sps.
Run t-test in Perseus (volcano plot):
Import filtered and corrected data [Perseus Analysis Files: 4_AfterBasicFilters.txt]: Matrix > Load > Generic matrix upload >
Transform the data [ExampleAnalysis_Part2_CEP63: log2 (x+1)]
i. Log2-transformation: Matrix > Processing > Basic > Transform
ii. Transformation: log2(x+1)
Subset the LFQ intensities as a new matrix to run the statistical test [ExampleAnalysis_Part2_CEP63: CEP63 LFQ]. Matrix > Processing > Rearrange > Reorder/remove columns > keep only the LFQ Intensities of the bait of interest and the negative controls and transfer the rest of the main columns to the left. For example, the new matrix may contain the following main columns:
i. LFQ Intensity CD7_DMSO_ctrl
ii. LFQ Intensity CD9_DMSO_ctrl
iii. LFQ Intensity CD11_DMSO_ctrl
iv. LFQ Intensity CD12_DMSO_ctrl
v. LFQ Intensity CD13_DMSO_ctrl
vi. LFQ Intensity CD14_DMSO_ctrl
vii. LFQ Intensity CD7_DMSO_Cep63
viii. LFQ Intensity CD11_DMSO_Cep63
ix. LFQ Intensity CD12_DMSO_Cep63
x. LFQ Intensity CD14_DMSO_Cep63
Optional: Remove any outlier samples, if necessary.
i. Following plots can be used for determining the outliers:
1) PCA plot.
2) Profile plot (i.e., box plot).
3) Multi-scatter plot.
4) tSNE/UMAP (R software).
ii. Matrix > Processing > Rearrange > Reorder/remove columns > transfer respective outlier column(s) to the left, if any.
Filter out the proteins detected in less than two replicates [ExampleAnalysis_Part2_CEP63: Filtered: 2+ valid].
Matrix > Processing > Filter rows > Filter rows based on valid values.
i. Min. valids > Number: 2.
ii. Mode: In at least one group.
iii. Grouping: IP (one of your annotations).
iv. Values should be > Greater than: 0.
v. Filter mode: Reduce matrix.
Apply t-test (unpaired one-tailed Student’s t-test): Matrix > Analysis > Volcano plot.
i. Grouping: IP.
1) First group (right): CEP63.
2) Second group (left): ctrl.
ii. Test: t-test.
iii. Side: Left.
iv. Number of randomizations: 1,000.
v. FDR: 0.1, S0: 10 (initial settings).
Rank the proteins into statistical significance categories using π-value (Xiao et al., 2014):
Transfer the data to an Excel file.
In the table part of volcano plot: Points > Right-click on the data > Select all > Right-click on the data.
i. > Copy selected rows > paste to an Excel file.
ii. Alternatively: > Plain matrix export… > .txt which can then be imported into Excel as a tab delimited file.
In Excel, calculate the π-values [Perseus Analysis Files: 10_π-values.xlsx].
Create a new column and calculate -log(p-value) × Difference.
i. Should be 2nd and 3rd columns. -log(p-value) column might be converted to #NAME?
ii. “Difference” corresponds to fold-change.
Sort the proteins based on the π-values:
i. Using the natural separation of the π-values on the plot (Figure 18), set thresholds.
In this case, we used around 100, 30, 12, and 8:
1) Score 4: 100 > π-value
2) Score 3: 100 > π-value > 30
3) Score 2: 30 > π-value > 12
4) Score 1: 12 > π-value > 8
ii. Regardless of its π-value, DO NOT accept anything with:
1) Difference (fold-change) < 5
2) -log(p-val) < 0.9
iii. The FDR and S0 values in the volcano plot corresponding to these cut-offs in this case were:
1) CEP63: FDR: 0.03, S0: 10
2) CEP170: FDR: 0.06, S0: 10
iv. Go back to Perseus, and color the proteins on the volcano plot according to the scores:
1) CRITICAL: These colors will subsequently be used in the R script to assign scores. Therefore, make sure to use the exact same colors across analyses.
2) TIP: It can be helpful to search the highest and lowest few proteins of a given score in the volcano plot, color them first, and then color the ones that fall in between.
Figure 18. Volcano plot of the proteins ranked and colored based on π-value. From highest to lowest rank: ● Score 4; ●Score 3; ● Score 2; ● Score 1
Transfer the significant ranked values to Excel:
First, select all the non-colored (gray) points; then, invert the selection to keep the colored (significant) ones, as follows:
i. In the table part of volcano plot: Points >.
ii. Click on a gray (non-colored) dot in the “Symbol color” column and copy the value shown below the table [i.e., default value: Color2 (A = 255, R = 169, G = 169, B = 169)].
iii. Right-click on the data:
1) > Find…: Color2 [A = 255, R = 169, G = 169, B = 169].
2) > Look in: Symbol color.
3) > Find all > Right-click on the rows > Select all and close this window.
iv. Points > Right-click on the data > Invert selection. Now, all the significant, colored (non-gray) points should be selected.
v. Right-click on the data > Copy selected rows.
vi. Paste the copied data to the Excel file, corresponding tab [Perseus Analysis Files: 11_Selection_from_Volcano.xlsx].
1) Create a separate tab for each bait analyzed.
2) Rename the “-Log(p-Value)” column (which might be changed to “#NAME?” by Excel) as “minus_log10_p”.
3) Correct the Septins and other problematic gene names (mentioned in section B6 and Figure 13) if converted to date.
Part III. Further analysis
Combine and summarize the data (TIMING: 0.5 day)
Combine and clean-up the data saved in Selection_from_Volcano.xlsx using R [Perseus Analysis Files: 12_Tidy_Data.R]. This script generates two Excel files:
Significant_Final.xlsx. Each tab of this file lists the proteins enriched by one bait, including Gene names, Protein IDs, Protein names, -log(p-value), log(fold-change) (i.e., Difference), and significance score (4: highest, 1: lowest).
Significant_Final_Frequencies.xlsx. This table summarizes by how many baits each protein was pulled down and with which score.
Further analysis suggestions (TIMING: variable)
Networks can be generated using bait-protein pairwise interactions using the Cytoscape software: https://cytoscape.org/.
Protein-protein interactions among the interactors of the baits can be analyzed for Gene Ontology (GO) terms using STRING database: https://string-db.org/.
Use the Multiple proteins search option and use the Gene.names2 column of the Significant_Final.xlsx file for the search, as it lists single protein names instead of protein groups.
Recipes
Note: For the solutions that require pH adjustment, use initially ~60%–80% of the volume of the water to solve the reagents, and complete the volume only after adjusting the pH.
2 M NaCl
Reagent MW Quantity
NaCl 58.44 g/mol 23.376 g
Distilled water
Final 200 mL
*Can be kept at room temperature.
0.5 M EDTA, pH 7.5
Reagent MW Quantity
EDTA 292.24 g/mol 14.612 g
Distilled water
Final 100 mL
*Dissolves after adding NaOH pellets and warming, pH 7.5. Can be kept at room temperature.
0.5 M EGTA, pH 7.6
Reagent MW Quantity
EGTA 380.35 g/mol 19.018 g
Distilled water
Final 100 mL
*Dissolves after adding NaOH crystals and warming, pH 7.6. Can be kept at room temperature.
1 M Tris-HCl, pH 7.5
Reagent MW Quantity
Tris base 121.14 g/mol 24.228 g
Distilled water
Final 200 mL
*Adjust the pH to 7.5 with concentrated HCl. Can be kept at room temperature.
0.5 M Tris-HCl, pH 6.8
Reagent MW Quantity
Tris base 121.14 g/mol 12.11 g
Distilled water
Final 200 mL
*Adjust the pH to 6.8 with concentrated HCl. Can be kept at room temperature.
10% Triton X-100 (v/v)
Reagent Stock Final concentration Quantity
Triton X-100 100% 10% 20 mL
Distilled water 180 mL
Final 200 mL
*Can be kept at room temperature.
10% SDS (w/v)
Reagent MW Quantity
SDS (powder) 288.372 g/mol 10 g
Distilled water, up to 100 mL
Final 100 mL
*Can be kept at room temperature.
ROCK inhibitor, 10 mM
Reagent MW Quantity
Y-27632 (dihydrochloride), powder 320.3 g/mol 5 mg
Water, sterile 1.56 mL
Final 1.56 mL
*Aliquot and keep at -20 °C. Once thawed, the vial can be kept at 4 °C.
Dorsomorphin, 5 mM
Reagent MW Quantity
Dorsomorphin, powder 399.5 g/mol 5 mg
DMSO, sterile 2.5 mL
Final 2.5 mL
*Observe the solution closely when dissolving dorsomorphin and look for floating tiny particles. If it is not dissolved completely, try warming up to 37 or 50 °C for 15–20 min. Aliquot and keep at -20 °C. Once thawed, the vial can be kept at 4 °C.
SB431542, 10 mM
Reagent MW Quantity
SB431542 hydrate, powder 384.39 g/mol 5 mg
DMSO, sterile 1.3 mL
Final 1.3 mL
*Aliquot and keep at -20 °C. Once thawed, the vial can be kept at 4 °C.
Nocodazole, 3.3 mM
Reagent MW Quantity
Nocodazole, powder 301.32 g/mol 2 mg
DMSO, sterile 2.0114 mL
Final 2.0114 mL
*Aliquot and keep at -20 °C. Thawed vial can be re-frozen and used a few times.
Poly-L-Ornithine, 10 mg/mL
Reagent MW Quantity
Poly-L-Ornithine hydrobromide, powder 30–70 kDa 500 mg
Water, sterile 50 mL
Final 50 mL
*Aliquot and keep at -20 °C. Once thawed, the vial can be kept at 4 °C.
Neural maintenance medium (N3)
Reagent Stock Dilution Final concentration Volume
DMEM/F-12, GlutaMAXTM supplement ~1:2 ~0.5× 241 mL
Neurobasal medium (1×) [-] L-Glutamine ~1:2 ~0.5× 241 mL
B27 with vitamin A 50× 1:100 0.5× 5 mL
Penicillin-Streptomycin 100× 1:100 1× 5 mL
N2 supplement 100× 1:200 0.5× 2.5 mL
Non-essential amino acids 100× 1:200 0.5× 2.5 mL
GlutaMAXTM supplement 100× 1:200 0.5× 2.5 mL
2-mercaptoethanol 50 mM 1:1,000 50 μM 500 μL
Insulin 10 mg/mL 1:4,000 2.5 μg/mL 125 μL
Final 500 mL
*Filter media using a 0.22 μm vacuum filter. N3 medium can be stored for up to three weeks at 4 °C.
Neural induction medium (N3+SMADi)
Reagent Stock Dilution Final concentration Volume
Neural maintenance medium (N3) - - - 50 mL
SB431542 10 mM 1:1,000 10 μM 50 μL
Dorsomorphin 5 mM 1:5,000 1 μM 10 μL
Final 50 mL
* After adding SMAD inhibitors (i.e., as N3+SMADi), keep the medium at 4 °C and use within five days.
Protein standard (BSA) (64 mg/mL)
Reagent Stock
BSA 320 mg
IP/wash buffer 5 mL
Final 5 mL
*Aliquot and keep at -20 °C. Thawed vial can be re-frozen and used a few times.
1× Laemmli buffer
Reagent Stock Final concentration Volume
Tris-HCl, pH 6.8 0.5 M 32.9 mM 658 μL
SDS 10% 1% 1 mL
Distilled water 8.092 mL
Final 9.75 mL
*Stable at room temperature for six months. Mix with 2-mercaptoethanol at 39:1 ratio as a small aliquot just before using (e.g., 975 μL of 1× Laemmli + 25 μL of 2-mercaptoethanol).
IP/wash buffer A
Reagent Stock Final concentration Volume
Distilled water 38.05 mL
Tris-HCl, pH 7.5 1 M 50 mM 2.5 mL
NaCl 2 M 150 mM 3.75 mL
EDTA 0.5 M 1 mM 100 μL
EGTA, pH 7.6 0.5 M 1 mM 100 μL
Triton X-100 10% 1% 5 mL
SDS 10% 0.1% 500 μL
Protease inhibitor 1 tablet
Final 50 mL
*Make fresh on the day of use.
Wash buffer B
Reagent Stock Final concentration Volume
Distilled water 43.55 mL
Tris-HCl, pH 7.5 1 M 50 mM 2.5 mL
NaCl 2 M 150 mM 3.75 mL
EDTA, pH 7.5 0.5 M 1 mM 100 μL
EGTA, pH 7.6 0.5 M 1 mM 100 μL
Protease inhibitor 1 tablet
Final 50 mL
*Same as IP/wash buffer A, but without Triton X-100 and SDS. Make fresh on the day of use.
Acknowledgments
We thank Magdalena Götz and Stefanie Hauck for excellent technical help and mentorship during the progress of this project. Juliane Merl-Pham and Julia Schessner for their feedback on proteomic analyses. Florian Kammerstetter for the comprehensive feedback on the Perseus pipeline and for providing the cell count ranges for various cell splitting ratios. Adam O’Neill was supported by a Philip Wrightson Postdoctoral Fellowship from the Neurological Foundation of New Zealand. Funding for this work and Fatma Uzbas was provided from the ERC (advanced grant NeuroCentro, 885382) and the EU (NSC-Reconstruct, 874758). This protocol was derived and validated from original research described in O’Neill et al. (2022).
Competing interests
The authors declare no competing interests.
References
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Supplementary information
The following supporting information can be downloaded here:
MaxQuant Analysis Files
MaxQuant version 1.6.17.0
Perseus Analysis Files
Perseus version 1.6.14.0
Article Information
Copyright
© 2023 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
Molecular Biology > Protein > Protein-protein interaction
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Improve Research Reproducibility
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Peer-reviewed
An in vitro Assay to Probe the Formation of Biomolecular Condensates
YZ Yu Zhang
SL Shen Lisha
Published: Vol 13, Iss 17, Sep 5, 2023
DOI: 10.21769/BioProtoc.4813 Views: 1443
Reviewed by: Emilia KrypotouLu Liu Anonymous reviewer(s)
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Bio-protocol journal peer-reviewed
Sep 05, 2023 | This version
Preprint
Aug 14, 2022
Original Research Article:
The authors used this protocol in Science Advances Jun 2022
Abstract
Biomolecular condensates are membrane-less assemblies of proteins and nucleic acids formed through liquid–liquid phase separation (LLPS). These assemblies are known to temporally and spatially regulate numerous biological activities and cellular processes in plants and animals. In vitro phase separation assay using recombinant proteins represents one of the standard ways to examine the properties of proteins undergoing LLPS. Here, we present a detailed protocol to investigate in vitro LLPS using in vitro expressed and purified recombinant proteins.
Keywords: Biomolecular condensates Liquid–liquid phase separation In vitro phase separation Recombinant protein
Background
Liquid–liquid phase separation (LLPS) is mostly triggered by multivalent interactions between macromolecules including proteins and nucleic acids (Boeynaems et al., 2018). Several factors, such as protein concentration, pH, post-translational modifications, or the presence of other molecules, affect the process of phase separation by affecting the multivalent interactions (Saha et al., 2016; Dao et al., 2018; Dignon et al., 2020; Li et al., 2022). Both in vitro and in vivo strategies have been developed to test whether a protein undergoes LLPS, including the in vitro phase separation using recombinant proteins and fluorescence recovery after photobleaching (FRAP). In vitro phase separation assay offers a simple and fast detection method and is necessary to determine whether a protein undergoes phase separation. This approach could be used to evaluate any protein of interest. In this protocol, we take GFP and GFP-HRLP (Zhang et al., 2022) as examples to present a step-by-step guide for this approach.
Materials and reagents
pGEX-6p-2 vector (Pharmacia, 27-4598-01)
Primers used to construct GST-GFP and GST-GFP-HRLP:
GST-GFP-F-BamHI: 5′-CGCGGATCCATGAGTAAAGGAGAAGAACT-3′
GST-GFP-R-EcoRI: 5′-CCGGAATTCTTTGTATAGTTCATCCATGC-3′
HRLP-F-SalI: 5′-ACGCGTCGACTCATGCCACCGAAGGTTGTGAAG-3′
HRLP-R-NotI: 5′-AAGGAAAAAAGCGGCCGCTCAGTAGTATGATCCTGGAC-3′
Rosetta (DE3) competent cells (Novagen, catalog number: 70954)
Ampicillin (Sigma-Aldrich, CAS number: 7177-48-2)
Isopropyl β-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, CAS number: 367-93-1)
Glutathione Sepharose beads (GE Healthcare, catalog number: GE17-0756-05)
PreScission protease (GE Healthcare, catalog number: 27-0843-01)
Bio-Rad protein assay dye reagent concentrate (Bio-Rad, catalog number: 5000006)
Bovine serum albumin (BSA) (Sigma-Aldrich, CAS number: 9048-46-8)
Precision Plus protein standard all blue marker (Bio-Rad, catalog number: 1610373)
PEG-8000 (Sigma-Aldrich, catalog number: 25322-68-3)
Hole microscope slide (BoliOptics, catalog number: SL39101004)
Cover glass (TRUSCO, catalog number: 122-9777)
Tryptone (Thermo Fisher Scientific, catalog number: 211705)
NaCl (Sigma-Aldrich, CAS number: 7647-14-5)
Yeast extract (Thermo Fisher Scientific, catalog number: 211929)
Agar (for LB plates) (BD, catalog number: 214010)
Methanol (Sigma-Aldrich, CAS number: 67-56-1)
Acetic acid (Sigma-Aldrich, CAS number: 64-19-7)
Coomassie Brilliant Blue R-250 (Bio-Rad, catalog number: 1610400)
Tris (Vivantis, catalog number: PR0612)
EDTA (Sigma-Aldrich, CAS number: 60-00-4)
Triton X-100 (Bio-Rad, catalog number: 1610407)
DTT (Roche, CAS number: 3483-12-3)
Complete EDTA-free protease inhibitor cocktail (Roche, catalog number: 5056489001)
Glycine (Bio-Rad, catalog number: 1610718)
SDS (Vivantis, catalog number: 151-21-3)
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)
LB medium (see Recipes)
Coomassie blue staining buffer (see Recipes)
Destaining solution (see Recipes)
Lysis buffer (see Recipes)
Cleavage buffer (see Recipes)
SDS running buffer (see Recipes)
10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel (see Recipes)
Recipes
LB medium
10 g/L tryptone
10 g/L NaCl
5 g/L yeast extract
15 g/L agar (for LB plates)
Coomassie blue staining buffer
20% methanol
10% acetic acid
0.1% Coomassie Brilliant Blue R-250
Destaining solution
20% methanol
10% acetic acid
Lysis buffer
10 mM Tris-HCl pH 8.0
15 mM NaCl
1 mM EDTA
1% Triton X-100
5 mM DTT
1× Complete EDTA-free protease inhibitor cocktail
Cleavage buffer
50 mM Tris-HCl pH 7.0
150 mM NaCl
1 mM EDTA
1 mM DTT
0.01% Triton X-100
SDS running buffer
3.03 g/L Tris
14.4 g/L glycine
1 g/L SDS
10% SDS-PAGE gel (for two 1.0 mm gels)
Separation gel:
4.1 mL of H2O
3.3 mL of 30% acrylamide/bis 37.5:1
2.5 mL of 1.5 M Tris-HCl pH 8.8
100 μL of 10% SDS
100 μL of APS
10 μL of TEMED
Stacking gel:
3.05 mL of H2O
0.65 mL of 30% Acrylamide/bis 37.5:1
0.625 mL of 1 M Tris-HCl pH 6.8
50 μL of 10% SDS
50 μL of APS
5 μL of TEMED
Equipment
Incubator (Panasonic, catalog number: 24127-88)
Shaker (INFORS, model: SKU BLE2000100)
Sonicator (Sonics, model: VCX-130)
Microcentrifuge (Thermo Fisher Scientific, catalog number: 75003424)
Centrifuge (Beckman Coulter, model: Allegra X-22)
Nutating mixer (Labnet, model: S0500-230V-EU)
Protein vertical electrophoresis cell (Bio-Rad, catalog number: 658004)
PowerPac power supply (Bio-Rad, catalog number: 1645070)
Confocal microscope (Olympus, model: FV3000)
Software
FV31-SW (Olympus)
Procedure
Recombinant protein expression
Clone the coding sequences of GFP and GFP fused with HRLP into the pGEX-6p-2 vector to generate GST-GFP and GST-GFP-HRLP constructs, respectively.
Transform the constructs of GST-GFP and GST-GFP-HRLP into DE3 competent cells.
Spread the transformed competent cells onto LB plates containing 100 mg/L ampicillin and incubate overnight at 37 °C to form colonies.
Inoculate one single colony into 2 mL of LB liquid medium supplemented with ampicillin at the final concentration of 100 mg/L and incubate at 37 °C overnight with vigorous shaking at 220 rpm.
Transfer the overnight cell culture to 200 mL of LB liquid medium supplemented with 100 mg/L ampicillin and grow at 37 °C with shaking at 220 rpm until the OD600 value reaches 0.6–1.0.
Add IPTG at the final concentration of 0.2 mM to the cultured cells to induce proteins of interest at 16 °C overnight (Note 1).
Test the solubility of GST-GFP and GST-GFP-HRLP fusion proteins by running a protein gel followed by Coomassie blue staining through the following steps:
Transfer 2 mL of induced and non-induced cell cultures to Eppendorf tubes, followed by centrifugation at maximum speed for 1 min to pellet the cells. Meanwhile, keep the rest of the cell cultures at 4 °C.
Add 1 mL of lysis buffer (Note 2) to resuspend the cells and further break cells by sonication (output watt at 6, 30 s on/30 s off) until the solution becomes clear.
Centrifuge the lysed cells at maximum speed at 4 °C for 10 min and transfer the supernatant into a new 1.5 mL tube as the soluble proteins.
Add 6× SDS loading dye to the supernatant and denature the proteins at 100 °C for 10 min.
Run an SDS-PAGE gel. After running is completed, stain the gel with Coomassie blue staining buffer for at least 1 h at room temperature.
Destain the gel by soaking the gel in destaining solution. Induced recombinant protein bands should appear after IPTG induction (Figure 1A).
Figure 1. Protein staining with Coomassie blue. (A) GST-GFP and GST-GFP-HRLP expression after IPTG induction. Red asterisks indicate the protein bands of GST-GFP and GST-GFP-HRLP. (B) GFP and GFP-HRLP proteins after cleavage. The red asterisks indicate the protein bands of GFP and GFP-HRLP.
Recombinant protein purification
After confirming the solubility of proteins, harvest the induced cells containing GST-GFP and GST-GFP-HRLP by centrifugation at 2,400× g for 5 min at 4 °C.
Resuspend the cells in 10 mL of ice-cold lysis buffer, followed by sonication (output watt at 6, 30 s on/ 30 s off) until the cell suspension becomes clear.
Centrifuge the suspension at 10,000× g for 20 min at 4 °C.
Transfer the supernatant containing induced proteins into new 15 mL Falcon tubes.
Equilibrate the Glutathione Sepharose beads by washing the beads with 1 mL of lysis buffer.
Transfer 150 μL of equilibrated beads to each supernatant, followed by an incubation for 2 h at 4 °C with gentle rotation.
Centrifuge at 200× g for 1 min at 4 °C to pellet the beads.
Discard the supernatants and wash beads with 5 mL of ice-cold lysis buffer for 5 min.
Repeat the washing step twice.
To test the protein quality and size, boil 2 μL of washed beads in 10 μL of lysis buffer supplemented with 6× SDS loading dye at 100 °C for 10 min and run in an SDS-PAGE gel.
After running is completed, stain the gel with Coomassie blue buffer for 1 h at room temperature.
Destain the gel with destaining solution; the target protein bands should appear after IPTG induction. For the rest of the beads, keep at 4 °C until the cleavage step.
Cleavage of recombinant protein
Add 5 μL of PreScission Protease and 200 μL of cleavage buffer to the washed beads.
Incubate the beads on a nutating mixer at 4 °C for 4 h (Note 3).
Centrifuge the beads at 600× g for 1 min at 4 °C.
Transfer the green supernatants to new 1.5 mL tubes to store GFP and GFP-HRLP proteins (Figure 2). The proteins can be stored in cleavage buffer in small aliquots at -80 °C for several months (Note 4).
Figure 2. GFP (left) and GFP-HRLP (right) proteins after cleavage
Measure the protein concentration with Bio-Rad protein assay dye reagent concentrate (BSA protein could be used to generate the standard curve).
Run 2 μL of GFP and GFP-HRLP proteins on a protein gel followed by Coomassie blue staining to check the protein quality and size (Figure 1B) (Note 5).
In vitro phase separation
Dilute the generated GFP and GFP-HRLP proteins at the final concentration of 10 μM (Note 6) with the cleavage buffer on ice.
Add 2 μL of PEG-8000 (Note 7) at the final concentration of 10% to the 18 μL of diluted proteins in a PCR tube on ice. Mix the samples by pipetting up and down.
Transfer the 20 μL mixture to a Hole microscope slide immediately to avoid protein degradation or fluorescence decay.
Carefully cover the slide with a cover glass.
Observe the fluorescence with 40× microscope objective under an FV3000 Olympus confocal microscope. GFP protein exhibits uniform signals under the microscope, whereas GFP-HRLP forms droplets (Figure 3) (Notes 8–10).
Figure 3. In vitro phase separation of GFP and GFP-HRLP proteins with the addition of PEG-8000. Scale bars, 10 μm.
Data analysis
The in vitro phase separation assay should be repeated through the above procedures using independently expressed and purified recombinant proteins. The negative control protein GFP should always be included. If a protein of interest but not the GFP control consistently form droplets in several independent experiments, this protein of interest undergoes LLPS in vitro. Other methods including timelapse imaging and FRAP are needed to further confirm the LLPS property of the protein of interest.
Notes
After IPTG induction, the color of the cell culture usually turns green, which is an indicator of successful induction of GFP fusion proteins.
The concentration of NaCl in the lysis buffer could be adjusted based on the specific properties of the protein of interest, as certain phase-separating proteins may require higher concentrations of NaCl to enhance their solubility. This adjustment is aimed at optimizing the conditions for maintaining the protein of interest in a soluble state.
The concentration of PreScission protease and the duration of incubation time could be adjusted on a case-by-case basis to avoid protein degradation.
The protein aliquots should be stored in small volumes, and repeated freeze-thaw cycles should be avoided to maintain the integrity and activity of proteins.
A size-exclusion chromatography assay is suggested to effectively eliminate nonspecific proteins and contaminants.
Optimal protein concentrations should be determined on a case-by-case basis. It is recommended to evaluate whether liquid droplet formation is concentration dependent.
The concentrations of PEG-8000 could be adjusted. Other crowding reagents, such as dextran and Ficoll, could also be used.
Some proteins may need longer time to form droplets.
The formation of protein droplets is affected by multiple factors such as pH, salt concentration, protein concentration, temperature, and PEG concentration. Modifying these parameters may promote droplet formation.
The presence of a GFP tag may influence the phase separation properties of certain proteins. In such cases, it is recommended to remove the GFP tag and conduct LLPS observations under a Differential Interference Contrast (DIC) microscopy. DIC microscopy allows for the visualization of phase-separated structures without the potential interference from the GFP tag, providing a clearer assessment of a protein's inherent phase separation behavior.
Acknowledgments
This work was supported by the National Research Foundation Competitive Research Programme (NRF-CRP22-2019-0001) and the intramural research support from Temasek Life Sciences Laboratory.
This protocol was derived from the original work of Zhang et al. (2022).
Competing interests
The authors declare no competing interests.
References
Boeynaems, S., Alberti, S., Fawzi, N. L., Mittag, T., Polymenidou, M., Rousseau, F., Schymkowitz, J., Shorter, J., Wolozin, B., Van Den Bosch, L., et al. (2018). Protein Phase Separation: A New Phase in Cell Biology. Trends Cell Biol. 28(6): 420–435.
Dao, T. P., Kolaitis, R. M., Kim, H. J., O’Donovan, K., Martyniak, B., Colicino, E., Hehnly, H., Taylor, J. P. and Castañeda, C. A. (2018). Ubiquitin Modulates Liquid-Liquid Phase Separation of UBQLN2 via Disruption of Multivalent Interactions. Mol. Cell 69(6): 965–978.e6.
Dignon, G. L., Best, R. B. and Mittal, J. (2020). Biomolecular Phase Separation: From Molecular Driving Forces to Macroscopic Properties. Annu. Rev. Phys. Chem. 71(1): 53–75.
Li, J., Zhang, M., Ma, W., Yang, B., Lu, H., Zhou, F. and Zhang, L. (2022). Post-translational modifications in liquid-liquid phase separation: a comprehensive review. Mol. Biomed. 3(1): e1186/s43556-022-00075-2.
Saha, S., Weber, C. A., Nousch, M., Adame-Arana, O., Hoege, C., Hein, M. Y., Osborne-Nishimura, E., Mahamid, J., Jahnel, M., Jawerth, L., et al. (2016). Polar Positioning of Phase-Separated Liquid Compartments in Cells Regulated by an mRNA Competition Mechanism. Cell 166(6): 1572–1584.e16.
Zhang, Y., Fan, S., Hua, C., Teo, Z. W. N., Kiang, J. X., Shen, L. and Yu, H. (2022). Phase separation of HRLP regulates flowering time in Arabidopsis. Sci. Adv. 8(25): eabn5488.
Article Information
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© 2023 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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4,814 | https://bio-protocol.org/en/bpdetail?id=4814&type=0 | # Bio-Protocol Content
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Peer-reviewed
Quantitative Analysis of Clot Deposition on Extracorporeal Life Support Membrane Oxygenators Using Digital and Scanning Electron Microscopy Imaging Techniques
YZ Yanyi Zang
TR Teryn R. Roberts
GH George T. Harea
BB Brendan M. Beely
LP Leonardo J. Olivera Perez
SA Sreedevi Ande
MB Maria Batchinsky
JL Ji H. Lee
MT Marianne A. Thrailkill
MR Melissa M. Reynolds
AB Andriy I. Batchinsky
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4814 Views: 497
Reviewed by: Komuraiah MyakalaBhanu JagilinkiAndrew Doyle
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Abstract
Device-induced thrombosis remains a major complication of extracorporeal life support (ECLS). To more thoroughly understand how blood components interact with the artificial surfaces of ECLS circuit components, assessment of clot deposition on these surfaces following clinical use is urgently needed. Scanning electron microscopy (SEM), which produces high-resolution images at nanoscale level, allows visualization and characterization of thrombotic deposits on ECLS circuitry. However, methodologies to increase the quantifiability of SEM analysis of ECLS circuit components have yet to be applied clinically. To address these issues, we developed a protocol to quantify clot deposition on ECLS membrane oxygenator gas transfer fiber sheets through digital and SEM imaging techniques. In this study, ECLS membrane oxygenator fiber sheets were obtained, fixed, and imaged after use. Following a standardized process, the percentage of clot deposition on both digital images and SEM images was quantified using ImageJ through blind reviews. The interrater reliability of quantitative analysis among reviewers was evaluated. Although this protocol focused on the analysis of ECLS membrane oxygenators, it is also adaptable to other components of the ECLS circuits such as catheters and tubing.
Key features
• Quantitative analysis of clot deposition using digital and scanning electron microscopy (SEM) techniques
• High-resolution images at nanoscale level
• Extracorporeal life support (ECLS) devices
• Membrane oxygenators
• Blood-contacting surfaces
Graphical overview
Keywords: Quantitative analysis Scanning electron microscopy ImageJ Membrane oxygenator Extracorporeal life support (ECLS) Clot deposition Thrombosis
Background
Device-induced thrombosis remains one of the major complications of extracorporeal life support (ECLS) (Jaffer et al., 2015; Doyle and Hunt, 2018). The interaction of blood and artificial surfaces of ECLS components results in clot deposition on these surfaces and in severe instances can lead to occlusion of the circuit and systemic complications (Roberts et al., 2020a). Clinically available antithrombotic treatments can introduce hemorrhagic complications, especially in patients with preexisting bleeding disorders (MacLaren et al., 2022). Non-thrombogenic circuits that reduce thrombotic complications and enable improved patient outcomes are needed. An important prerequisite to developing such circuits is to more thoroughly understand how blood components interact with the artificial surfaces of ECLS devices (Beely et al., 2016). However, there is currently no methodology for routine examination of these devices after use. Surfaces of ECLS components could be specifically engineered to address the issue of thrombosis with a better understanding of the interactions between blood and ECLS surfaces. Examination and analysis of extracorporeal circuitry following clinical use in a standardized and validated method could inform both clinical decision-making regarding anticoagulation as well as device design.
Digital images have been used to record and visualize the extent of clot formation within ECLS devices in small pilot research studies, both clinically and in translational research laboratories (Lehle et al., 2008; Beely et al., 2016; Diehl and Gantner, 2018; Chlebowski et al., 2020; Naito et al., 2021). Blood clots are easily observed through digital images, while some microscopic thrombotic structures are not. Scanning electron microscopy (SEM), an advanced technology, produces high-resolution images at nanoscale level. Although most of the clinical use of SEM is qualitative, new methodologies that increase the quantifiability of SEM have been developed (Kundu et al., 1988; Di Iorio et al., 2005). We have previously used SEM to visualize and characterize clots and cell deposits in ECLS circuits following use in translational research studies in swine up to 72 h in duration (Beely et al., 2016; Roberts et al., 2020b). In this protocol, we developed and standardized a method to quantitatively analyze clot deposition using both digital and SEM images, allowing to routinely evaluate ECLS components following clinical use. Although this protocol focused on the analysis of ECLS membrane oxygenators only, this method could be applied to other components of the ECLS with necessary adaptations.
Materials and reagents
Reagents
Glutaraldehyde, 50% biological grade (Electron Microscopy Sciences, catalog number: 16520)
Sucrose, ≥ 99.5% (Sigma-Aldrich, catalog number: S9378)
Sodium cacodylate trihydrate, ≥ 98% (Sigma-Aldrich, catalog number: C0250)
Phosphate buffered saline (PBS) powder, 10×, pH 7.4 (Fisher BioReagents, catalog number: BP665-1)
Dehydrant alcohol, 100% (Epredia, catalog number: 6215)
Milli-Q water, 18.2 MΩ·cm, obtained through BarnsteadTM Smart2PureTM Pro Water Purification System
Solutions
Scanning electron microscopy fixative (SEM fixative) (see Recipes)
Scanning electron microscopy buffer (SEM buffer) (see Recipes)
Phosphate buffered saline, pH 7.4 (PBS) (see Recipes)
Recipes
SEM fixative
Reagent Final concentration Quantity
Sucrose 0.1 M 8.5575 g
Sodium cacodylate trihydrate 0.1 M 5.3508 g
Glutaraldehyde, 50% 3% 15 mL
Milli-Q water n/a 235 mL
SEM buffer
Reagent Final concentration Quantity
Sucrose 0.1 M 8.5575 g
Sodium cacodylate trihydrate 0.1 M 5.3508 g
Milli-Q water n/a 250 mL
PBS (based on vendor instruction)
Reagent Final concentration Quantity
PBS powder, 10× 1× 98.9 g
Milli-Q water n/a 10 L
Laboratory supplies
Slotted tissue cassette (Epredia, catalog number: B851729WH)
Container, multi-purpose, PP, with separate snap lid, 120 mL (Globe Scientific, catalog number: 271004)
Copper conductive tape, single adhesive surface (Ted Pella, catalog number: 16072-1)
Carbon conductive tape, double coated (Ted Pella, catalog number: 16073-5)
T10 Torx Screwdriver (Tekton, model number: DST31100)
Sterilization wrap (Halyard, catalog number: 34162)
Premium blue indicating silica gel desiccant beads 3–5 mm (Dry & Dry, catalog number: X000TS1IZP)
Straight connector, female luer lock port (Qosina, catalog number: 73317)
Needleless injection site, swappable, male luer lock (Qosina, catalog number: 80147)
50 mL syringe, luer-lok tip (BD, catalog number: 309653)
Scissors
Surgical grade stainless steel forceps, nonlocking thumb handle straight delicate, serrated tips, 4 3/4" in. (McKesson, catalog number: 43-1-772)
Pathology metric ruler
Weighing paper or weighing boat
Spatula
Cylinder
Desiccator, 6"
SEM fixative/buffer waste bottle
Dehydration/ethanol waste bottle
Labels with study ID, protocol number, and date
Personal protective equipment (gloves, goggles, hearing protection, and scrubs/lab coats)
Equipment
Miter saw coupled with diamond blade (Miter saw DeWALT, model: DWS779; blade RIDGID, model: 12 in. dual-purpose)
Digital camera with lens (Nikon, camera model: D3300, lens model: AF-P DX NIKKOR 18-55 f/3.5-5.6G VR)
BarnsteadTM Smart2PureTM Pro Water Purification System (Thermo Fisher Scientific, catalog number: 50157873)
Gold sputter coater (Denton Vacuum, model: Desk II)
InTouchScopeTM scanning electron microscope (JEOL, model: JSM-IT100)
Impact suction unit (Envi health solutions, model: 326/326M)
Software and datasets
ImageJ (version: 1.53k, release date: 07/06/2021, free)
Analytics Software & Solutions (SAS) (version 9.4, release date: 07/2013, license needed)
Procedure
Membrane oxygenator preservation post-use
Disconnect membrane oxygenator from ECLS system pump.
Flush membrane oxygenator from the inlet port with 1× PBS and drain blood/PBS flush from the outlet port until flush exiting outlet is clear (standardize flush flow rate to the flow rate in which the ECLS device is operated).
Connect inlet and outlet tubing (10–15 cm length) into a closed loop with 3/8" × 3/8" connector (Figure 1).
Attach needleless injection port to 3/8" × 3/8" straight connector.
Inject SEM fixative into the oxygenator via a priming port using a 50 mL syringe while simultaneously withdrawing air from the oxygenator via a separate port. This process is continued until the membrane is saturated with fixative.
Store the membrane oxygenator in the refrigerator (4 °C) until ready for section B.
Figure 1. Membrane oxygenator preservation with SEM fixative
Membrane oxygenator disassembly (General note 1)
Drain SEM fixative slowly into SEM fixative/buffer waste bottle.
Rinse the membrane oxygenator with 500 mL of 1× PBS.
Unscrew and disassemble membrane oxygenator pump using a T10 Torx screwdriver.
Take digital images of membrane oxygenator pump, inlet face, and outlet face of the membrane oxygenator on sterilization wrap (use as a blue background) with labels and pathology metric ruler. Examples are in Figure 2.
Figure 2. Example images of (a) membrane oxygenator pump, (b) inlet/top/pre-membrane/venous face of the membrane oxygenator, and (c) outlet/bottom/post-membrane/arterial face of the membrane oxygenator
Cut off water connections to get a flat edge, then cut off post-membrane sample port on outlet side so that the membrane can sit flat on the miter saw table (Figure 3).
Figure 3. Examples of water connection and post-membrane sample port of a membrane oxygenator
Cut the four sides of the membrane case with inlet face facing up using a miter saw coupled with vacuum system to remove debris (schematic shown in Figure 4).
Figure 4. Schematic of membrane oxygenator dissection
Gently remove inlet face plastic cover using forceps.
The first gas transfer fiber sheet should be exposed and ready for imaging.
Membrane oxygenator gas transfer fiber sheet digital imaging (Troubleshooting 1)
Gently remove gas transfer fiber sheets individually in order from inlet face to outlet face.
Place every four gas fiber sheets next to each other horizontally with total three rows.
Take digital images of every 12 gas fiber sheets with Study ID and number labels as shown in Figure 5.
Figure 5. Layout example of membrane gas transfer fiber sheets for digital imaging
Take individual images of top inlet membrane gas fiber sheet, middle membrane gas fiber sheet, and bottom outlet membrane gas fiber sheet with study ID, date, and pathology metric ruler as shown in Figure 6.
Figure 6. Example of individual membrane gas fiber sheet for digital imaging
Cut top inlet, middle, and bottom outlet membrane gas transfer fiber sheets each into nine squares (total 27 squares). Number squares from top inlet membrane gas fiber sheet from 1 through 9, middle membrane gas fiber sheet from 10 through 18, and bottom outlet membrane gas fiber sheet from 19 through 27, as shown in Figure 7.
Figure 7. Schematic diagram of cutting individual membrane gas fiber sheets and numbering. Numbers 1–9 are assigned to inlet/top membrane gas fiber sheet (left), numbers 10–18 are assigned to middle membrane gas fiber sheet (middle), and numbers 19–27 are assigned to outlet/bottom membrane gas fiber sheet (right).
Place randomly selected squares into labeled cassettes with the surface that was closest to the membrane inlet facing the lid of the cassette facing upward.
Place cassettes with sample into 120 mL container, add SEM fixative, and store in refrigerator (4 °C) until ready for section D.
SEM sample dehydration (General Note 2)
Drain SEM fixative from each container into SEM fixative/buffer waste bottle.
Add SEM buffer into each container and incubate for 10 min; drain SEM buffer into SEM fixative/buffer waste bottle.
Add 30% dehydrant into each container and incubate for 10 min; drain 30% dehydrant into dehydrant/ethanol waste bottle.
Add 50% dehydrant into each container and incubate for 10 min; drain 50% dehydrant into dehydrant/ethanol waste bottle.
Add 70% dehydrant into each container and incubate for 10 min; drain 70% dehydrant into dehydrant/ethanol waste bottle.
Add 100% dehydrant into each container and incubate for 10 min; drain 100% dehydrant into dehydrant/ethanol waste bottle.
Add 100% dehydrant into each container and incubate for 10 min; drain 100% dehydrant into dehydrant/ethanol waste bottle.
Place container with samples into desiccator overnight under vacuum.
After overnight vacuum drying, fill 1/5 of the container with silica gel desiccant beads and close the lid tightly.
Keep the container with samples at room temperature until ready to do SEM imaging.
SEM imaging
SEM sample preparation (Figure 8)
Attach sample onto an SEM sample holder using double-sided conductive carbon tape.
Sputter coat with 20 nm of gold nanoparticles using Denton Vacuum Desk II Gold sputter coater.
Tape the opposite borders of the sample with conductive copper tape perpendicular to the sample fibers.
Figure 8. Schematic of SEM sample preparation process
SEM imaging for percentage of clot deposition quantitative analysis
Set up 14 kV accelerating voltage with 14 mm working distance using InTouchScopeTM scanning electron microscope.
Take images at 25× magnification horizontally from the upper left corner of sample to the lower right corner to cover the entire surface of each sample.
Quantitative analysis of digital images using ImageJ
Open ImageJ software and open the image for analysis. Click File and select Open to open an image.
Set up scale of the image (Figure 9).
Click Analyze and select Set Scale…
Enter 0 next to Known distance.
Enter pixels next to Unit of length.
Figure 9. Screenshot of Set Scale window
Select total area of interest (Figure 10).
Click Image, Adjust, and then Color Threshold…
Set Thresholding method as Default.
Set Color space as HSB to start and change to RGB if needed.
Check Dark background box.
Set threshold to cover the entire surface of the membrane gas fibers and exclude blue background.
Figure 10. Screenshot after selecting total area of interest
Click Select to highlight the total area of interest (Figure 11).
Figure 11. Screenshot of highlighted total area of interest after selection
Click Analyze and select Measure to show the results in Results window (Figure 12).
Figure 12. Screenshot of Results window
Enter the Area value into the Excel spreadsheet as total area.
Select clot deposition area of interest (Figure 13).
Click Original on the Threshold Color window to revert to the original image.
Adjust threshold to cover clot deposition area only.
Figure 13. Screenshot after selecting clot deposition area of interest
Click Select to highlight the clot deposition area.
Click Analyze and select Measure to show the results in Results window.
Enter the Area value into the spreadsheet as clot deposition area.
Calculate percentage of coverage using clot deposition area and total area on the spreadsheet (an example is shown below, Figure 14).
Figure 14. Screenshot of a calculation spreadsheet example
Quantitative analysis of SEM images using ImageJ (General Note 3)
Open ImageJ software and open the image for analysis. Click File and select Open to open an image.
Set up scale of the image.
Click Straight line icon and draw a line on the scale bar (Figure 15).
Figure 15. Screenshot of setting up scale for SEM images
Click Analyze and select Set Scale… to change unit from pixel to micrometer (Figure 16).
Enter scale bar value next to Known distance (Figure 16).
Enter scale bar unit next to Unit of length (Figure 16).
Click OK.
Figure 16. Screenshot of Set Scale window for SEM images
Click Analyze and then Set Measurements. Check boxes Area and Display Label. Click OK.
Click Analyze, Tools, and then ROI Manager… Check boxes Show All and Labels.
Use Polygon selections or Freehand selections to draw areas of interest (Figure 17).
Figure 17. Screenshot of Polygon and Freehand selections
After drawing area of interest, click Add [t] to add it to the list (same for all other areas of interest) (Figure 18).
Figure 18. Screenshot of drawing area of interest example for SEM images
Click one of the areas of interest. Click Rename and then enter the number of areas of interest next to Rename As. Click OK (same for all other areas of interest).
Select all areas of interest, and then click Measure.
Save Results into .csv format. Click File, and then Save As…
Open saved file.
Use the value under Area column divided by total area (Figure 19), which is shown above each image in ImageJ to get percentage of coverage.
Figure 19. Screenshot of the location of total area
Screenshot and save image with areas of interest as for recorder cord.
Data analysis
Perform statistical comparison on digital and SEM images percentage of total clot coverage of each set of samples (two-sided test with α = 0.05 for significance via SAS 9.4). Perform interrater reliability of quantitative analysis to evaluate the agreement among reviewers (General note 4).
Validation of protocol
Currently, there is no gold standard method for determining total clot deposition on medical device surfaces, therefore limiting the ability to compare our methods tested to an established methodology. This protocol has been validated through a blind review process in which individuals that are not familiar with this method were required to follow the protocol and perform the analysis without help. By the end of the process, we collected feedback and improved the protocol. So far, at least six individuals from three institutions have tried this protocol and there are no outstanding concerns or issues. In addition, this protocol results in a podium presentation at 37th Annual Children’s National Symposium: ECMO & the Advanced Therapies for Respiratory Failure (Virtual).
Zang et al. (2021). Quantitative Analysis of Digital vs. Scanning Electron Microscopy Images as An Assessment Tool for Post-Extracorporeal Life Support Clot Formation Evaluation. Oral Presentation at Bradley Hill Best Abstract Session.
General notes and troubleshooting
General notes
These steps only apply to Maquet HLS7.0 membrane and may require modifications for other membrane oxygenators.
It is recommended to perform SEM imaging the day after SEM dehydration to avoid sample moisturization during storage. If it is not practicable, we recommend checking and changing silica gel desiccant beads on a regular basis.
It took approximately 28 h to image and process a set of SEM images for one sample compared to approximately 3 min for one digital image of the same sample.
To avoid bias, we suggest choosing individuals who are not familiar with the studies involved in membrane oxygenator or de-identified samples before performing the analysis. For routine post-study analysis, interrater reliability score is used to see the reliability and validity to ensure the results are accurate and replicable.
Hazardous exposure (e.g., blood, hazardous chemicals in fixative, etc.) that could occur when assessing materials after clinical use may raise safety and health concerns.
SEM is expensive and time consuming, especially at institutions where SEM is not available or there are not enough trained technicians to perform SEM on a routine basis. Digital imaging could be an alternative approach to access clot deposition; however, some nanoscale clots will be missing from digital images. Therefore, we recommend the researchers and clinicians to balance the advantages and disadvantages of these methods and make a conscious decision based on their needs.
The most challenging part during the development of this protocol is section B. We highly recommend trying at least one test membrane oxygenator before performing on the membrane oxygenator to be analyzed.
Troubleshooting
Some of the thrombi were noticed to be stripped from the sample surface during sample processing and handling as the thrombus did not adhere to the surface tightly. This may lead to variability in quantitative analysis vs. the clinical state of the device during use. Please be as gentle as possible in this step.
Acknowledgments
The work was supported by the Assistant Secretary of Defense for the Health Affairs endorsed by the Department of Defense through the Peer Reviewed Medical Research Program, Technology/Therapeutic Development Award, under Grant no. W81XWH-18-2-0048, PI Andriy Batchinsky MD; Co-PI Teryn Roberts PhD.
The authors acknowledge Dr. Patrick McCurdy and Dr. Roy Geiss from the Central Instrument Facility at Colorado State University for their help on SEM. The authors also acknowledge the SEM facility at the School of Mathematics, Science and Engineering, University of the Incarnate Word and Dr. Carlos Garcia, the dean, for his support on SEM.
Competing interests
There are no conflicts of interest.
Ethical considerations
Membrane oxygenators were obtained from animal studies that were carried out in compliance with the Animal Welfare Act, implementing Animal Welfare Regulations, and the principles of the Guide for the Care and Use of Laboratory Animals, National Research Council at the Autonomous Reanimation and Evacuation Research Program (AREVA) (San Antonio, TX). The University of Texas at San Antonio Institutional Animal Care and Use (UTSA IACUC) approved all research conducted (Protocol SU001-03-23). The AREVA laboratory facility is fully accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care. The protocol and facility were also approved by the Department of Defense, Medical Research and Development Command Animal Care and Use Review Office.
References
Beely, B. M., Campbell, J. E., Meyer, A., Langer, T., Negaard, K., Chung, K. K., Cap, A. P., Cancio, L. C. and Batchinsky, A. I. (2016). Electron Microscopy as a Tool for Assessment of Anticoagulation Strategies During Extracorporeal Life Support: The Proof Is on the Membrane. ASAIO J. 62(5): 525–532.
Chlebowski, M. M., Baltagi, S., Carlson, M., Levy, J. H. and Spinella, P. C. (2020). Clinical controversies in anticoagulation monitoring and antithrombin supplementation for ECMO. Crit. Care 24(1): e1186/s13054-020-2726-9.
Di Iorio, D., Traini, T., Degidi, M., Caputi, S., Neugebauer, J. and Piattelli, A. (2005). Quantitative evaluation of the fibrin clot extension on different implant surfaces: Anin vitro study. J. Biomed. Mater. Res. Part B Appl. Biomater. 74B(1): 636–642.
Diehl, A. and Gantner, D. (2018). Pump head thrombosis in extracorporeal membrane oxygenation (ECMO). Intensive Care Med. 44(3): 376–377.
Doyle, A. J. and Hunt, B. J. (2018). Current Understanding of How Extracorporeal Membrane Oxygenators Activate Haemostasis and Other Blood Components. Front. Med. 5: e00352.
Jaffer, I. H., Fredenburgh, J. C., Hirsh, J. and Weitz, J. I. (2015). Medical device-induced thrombosis: what causes it and how can we prevent it?. J. Thromb. Haemost. 13: S72–S81.
Kundu, S. K., Klein, M. D., Whittlesey, G. C., Barmatoski, S. P. and Salley, S. O. (1988). Quantitative scanning electron microscopy for the evaluation of thrombosis in extracorporeal circuits. ASAIO Trans. 34(3): 568–572.
Lehle, K., Philipp, A., Gleich, O., Holzamer, A., Müller, T., Bein, T. and Schmid, C. (2008). Efficiency in Extracorporeal Membrane Oxygenation—Cellular Deposits on Polymethypentene Membranes Increase Resistance to Blood Flow and Reduce Gas Exchange Capacity. ASAIO J. 54(6): 612–617.
MacLaren, G., Peek, G., Lorusso, R., Brodie, D., Thiagarajan, R. and Vercaemst, L. (2022). Extracorporeal Life Support: The ELSO Red Book 6th Edition. Extracorporeal Life Support Organization.
Naito, N., Ukita, R., Wilbs, J., Wu, K., Lin, X., Carleton, N. M., Roberts, K., Jiang, S., Heinis, C., Cook, K. E., et al. (2021). Combination of polycarboxybetaine coating and factor XII inhibitor reduces clot formation while preserving normal tissue coagulation during extracorporeal life support. Biomaterials 272: 120778.
Roberts, T. R., Garren, M. R., Handa, H. and Batchinsky, A. I. (2020a). Toward an artificial endothelium: Development of blood-compatible surfaces for extracorporeal life support. J. Trauma Acute Care Surg. 89: S59–S68.
Roberts, T. R., Harea, G. T., Singha, P., Sieck, K. N., Beely, B. M., Wendorff, D. S., Choi, J. H., Ande, S., Handa, H., Batchinsky, A. I., et al. (2020b). Heparin-Free Extracorporeal Life Support Using Tethered Liquid Perfluorocarbon: A Feasibility and Efficacy Study. ASAIO J. 66(7): 809–817.
Zang, Y., Olivera Perez, L., Roberts, T., Ande, S., Lee, J., Reynolds, M. M. and Batchinsky, A. I. (2021). Quantitative Analysis of Digital vs. Scanning Electron Microscopy Images as An Assessment Tool for Post-Extracorporeal Life Support Clot Formation Evaluation. Virtual, February 23rd, 2021 (Oral Presentation at Bradley Hill Best Abstract Session).
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4,815 | https://bio-protocol.org/en/bpdetail?id=4815&type=0 | # Bio-Protocol Content
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Functional Phenotyping of Lung Mouse CD4+ T Cells Using Multiparametric Flow Cytometry Analysis
CM Céline M. Maquet
LG Laurent Gillet
BM Bénédicte D. Machiels
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4815 Views: 1346
Reviewed by: Alka MehraMartin V Kolev Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Science Immunology Jul 2022
Abstract
Gammaherpesviruses such as Epstein-Barr virus (EBV) are major modulators of the immune responses of their hosts. In the related study (PMID: 35857578), we investigated the role for Ly6Chi monocytes in shaping the function of effector CD4+ T cells in the context of a murine gammaherpesvirus infection (Murid gammaherpesvirus 4) as a model of human EBV. In order to unravel the polyfunctional properties of CD4+ T-cell subsets, we used multiparametric flow cytometry to perform intracellular staining on lung cells. As such, we have developed herein an intracellular staining workflow to identify on the same samples the cytotoxic and/or regulatory properties of CD4+ lymphocytes at the single-cell level. Briefly, following perfusion, collection, digestion, and filtration of the lung to obtain a single-cell suspension, lung cells were cultured for 4 h with protein transport inhibitors and specific stimulation media to accumulate cytokines of interest and/or cytotoxic granules. After multicolor surface labeling, fixation, and mild permeabilization, lung cells were stained for intracytoplasmic antigens and analyzed with a Fortessa 4-laser cytometer. This method of quantifying cytotoxic mediators as well as pro- or anti-inflammatory cytokines by flow cytometry has allowed us to decipher at high resolution the functional heterogeneity of lung CD4+ T cells recruited after a viral infection. Therefore, this analysis provided a better understanding of the importance of CD4+ T-cell regulation to prevent the development of virus-induced immunopathologies in the lung.
Key features
• High-resolution profiling of the functional properties of lung-infiltrating CD4+ T cells after viral infection using conventional multiparametric flow cytometry.
• Detailed protocol for mouse lung dissection, preparation of single-cell suspension, and setup of multicolor surface/intracellular staining.
• Summary of optimal ex vivo restimulation conditions for investigating the functional polarization and cytokine production of lung-infiltrating CD4+ T cells.
• Comprehensive compilation of necessary biological and technical controls to ensure reliable data analysis and interpretation.
Graphical overview
Graphical abstract depicting the interactions between immune cells infiltrating the alveolar niche and the lung during respiratory infection with a gammaherpesvirus (Murid herpesvirus 4, MuHV-4). Two distinct situations are represented: the inflammatory response developed during viral replication in the lung, either in the presence (WT mice) or absence of regulatory monocytes (CCR2KO mice). Sequential process of the experiment is represented, starting from intratracheal instillation of MuHV-4 virions to tissue dissociation and multicolor staining for flow cytometry analysis.
Keywords: Immunology Cytotoxicity Virology Multiparametric flow cytometry T-cell response Pro-inflammatory/regulatory cytokines CD4+ T-cell heterogeneity Lung immunity
Background
The complexity and heterogeneity of CD4+ T cells are increasingly being investigated. Indeed, in addition to their multiple roles as helper cells orchestrating the orientation of immune responses, CD4+ T cells also play major direct roles as regulatory or cytotoxic cells. In particular, it is becoming increasingly clear that CD4+ T cells, alongside with CD8+ T cells, are key determinants in anti-viral and anti-tumor immunity. Investigating the heterogeneity of CD4+ T-cell response, as well as their possible plasticity in different contexts of inflammation, can be achieved by high-resolution transcriptomic approaches such as sc-RNA sequencing. However, this expensive technique, relying on strong bioinformatics expertise, cannot always be used as a first line of investigation and/or requires additional validation of protein expression. In this context, the development of multicolor flow cytometry staining is a reliable and accurate tool as it allows the precise definition of the expression profile of surface molecules, while evaluating the functional properties of these populations in the presence of relevant stimuli. Multiparametric flow cytometry is therefore a widely used method for immunoprofiling circulating or tissue-resident cells. The originality and interest of the present protocol is to combine the staining of surface molecules suggestive of functional cell polarization (e.g., the checkpoint inhibitor PD-1 or the degranulation marker CD107a) with the quantification of the immune mediators expressed and/or released by these target cells. In that regard, the design of the restimulation cocktail is extremely important and requires a precise knowledge of the biological context of expression of each particular mediator. Indeed, the use of certain drugs such as phorbol 12-myristate 13-acetate and ionomycin triggers a massive and non-specific release of certain cytokines that does not necessarily reflect the behavior and level of activation of the cells of interest in vivo and makes it difficult to interpret the results. In addition, these aggressive treatments may also introduce a bias by affecting cell viability and leading to the selective loss of activated cells in vivo and therefore more sensitive to this type of treatment. Furthermore, while staining of some chemokines such as CXCL9 or cytotoxic mediators such as granzyme B can be achieved directly without any accumulation, others require simple accumulation (e.g., iNOS) or accumulation in presence of conditioned medium (e.g., IFNγ) for reliable detection. More specifically, the protocol presented here details the optimal conditions for assessing the regulatory vs. cytotoxic properties of CD4+ T cells recruited to the lung after respiratory infection with a gammaherpesvirus.
Materials and reagents
GentleMACS C tube (Miltenyi, catalog number: 130-093-237)
Hank’s buffered saline solution (HBSS) (Lonza, catalog number: BE10-543F)
DNase I (Roche, catalog number: 11284932001)
Collagenase D (Roche, catalog number: 11088866001)
Falcon, 15 and 50 mL conical centrifuge tubes (Corning, catalog numbers: 352096 and 352070)
Bovine serum albumin (BSA) (Sigma, catalog number: A8412)
Fetal calf serum (FCS) (Gibco, catalog number: 10082147)
EDTA (CorningTM, catalog number: 46-034-CI)
Falcon® 70 μm cell strainer (Corning, catalog number: 352350)
cOmpleteTM protease inhibitor cocktail (Roche, catalog number: 11697498001)
RBC lysis buffer (Thermo Fisher Scientific, catalog number: 00433357)
Trypan Blue solution, 0.4% (Sigma-Aldrich, catalog number: 93595)
96-well round (U) bottom plate, TC surface, pack of 1 (Thermo ScientificTM, catalog number: 163320)
RPMI 1640 medium (GibcoTM, catalog number: 11875093)
Monensin solution (BioLegend, catalog number: 420701)
Brefeldin A solution (BioLegend, catalog number: 420601)
β-mercaptoethanol (Sigma-Aldrich, catalog number: 3148)
L-Glutamine (CorningTM, catalog number: 25005CI)
Sodium pyruvate (CorningTM, catalog number: 25-000-CIR)
Sodium azide (Sigma, catalog number: 8223350100)
Penicillin-streptomycin (GibcoTM, catalog number: 15070063)
Purified anti-mouse CD28 antibody (BioLegend, catalog number: 102102)
Ultra-LEAFTM purified anti-mouse CD3ε antibody (BioLegend, catalog number: 100340)
Ultra-LEAFTM purified anti-mouse CD49d antibody (BioLegend, catalog number: 103709)
Anti-mouse CD107a allophycocyanin (APC) conjugated antibody (BioLegend, catalog number: 121613)
Anti-mouse CD16/32 Fc block (BioLegend, catalog number: 101301)
Zombie Aqua Fixable Viability kit (BioLegend, catalog number: 423101)
Paraformaldehyde (PFA) (Santa Cruz Biotechnology, catalog number: sc-281692)
Saponin (Sigma-Aldrich, catalog number: 47036)
BD FACSuiteTM CS&T research beads (BD Biosciences, catalog number: 650621)
Lung digestion medium (see Recipes)
Collagenase D working stock (see Recipes)
DNase I working stock (see Recipes)
Culture medium (see Recipes)
Wash buffer (see Recipes)
FACS buffer (see Recipes)
Permeabilization buffer (see Recipes)
Intracellular staining buffer (see Recipes)
Fc receptor blockade (see Recipes)
Intracellular staining mix (see Recipes)
Surface staining mix (see Recipes)
Zombie VioletTM Fixable Viability Kit (see Recipes)
Restimulation medium 1 (see Recipes and Table 1)
Restimulation medium 2 (see Recipes and Table 1)
Table 1. Summary of the specific cocktails of stimulation medium
Specific medium Milieu a (Il-10 and IFNγ producing T cells) Milieu b (Granzyme B and Granzyme A producing T cells) Milieu c (Perforin-1 producing T cells)
Media RPMI complete medium RPMI complete medium RPMI complete medium
Protein transport inhibitors Monensin (2 μM) and brefeldin (2 μM) Monensin (2 μM) and brefeldin (2 μM) Monensin (2 μM) and brefeldin (2 μM)
Additives β-mercaptoethanol (50 μM) β-mercaptoethanol (50 μM) β-mercaptoethanol (50 μM)
purified CD28 antibody (1 μg/mL) purified CD28 antibody (1 μg/mL)
purified CD3ε antibody (2 μg/mL) purified CD3ε antibody (2 μg/mL)
purified CD49d antibody (1 μg/mL)
APC anti-mouse CD107a (0.5 μg/mL; LAMP-1) antibody
Recipes
Lung digestion medium
HBSS medium supplemented with 5% heat-inactivated fetal calf serum
Collagenase D working stock
Resuspend lyophilized collagenase D with HBSS at a concentration of 20 mg/mL (stock solution). The reconstituted solution can be stored at -15 to -25 °C.
DNase I working stock
Resuspend lyophilized DNase I with nuclease-free water at a concentration of 1 mg/mL (stock solution). The reconstituted solution can be stored at -15 to -25 °C.
Culture medium
RPMI medium supplemented with 10% heat-inactivated FCS, 100 U/mL penicillin, 100 U/mL streptomycin, 2 mM L-glutamine, and 50 μM of β-mercaptoethanol.
Wash buffer
2 mM PBS-EDTA with 10% FCS
FACS buffer
PBS, 0.5% BSA, EDTA 2 mM, 2 mM NaN3 sodium azide
Note: FACS Buffer is a buffered saline solution that can be used for immunofluorescence staining protocols, antibody and cell dilution steps, wash steps required for surface staining, and flow cytometry analysis. The buffer contains sodium azide as preservative and animal serum proteins (BSA) to help minimize non-specific binding of antibodies. The addition of EDTA prevents cell-to-cell adhesion and clumping.
Permeabilization buffer
PBS, 0.1% saponin
Intracellular staining buffer
FACS buffer, 0.1% saponin
Fc receptor blockade
Add purified anti-mouse CD16/32 antibody diluted 1:500 in FACS buffer. Put 100 μL of total volume per well.
Surface staining mix
Add antibodies with appropriate dilutions predetermined by titration experiments in FACS buffer. Put 100 μL of total volume per well.
Zombie VioletTM Fixable working stock
Place the DMSO provided with the kit in a 37 °C water bath until it is completely thawed. Add 100 μL of DMSO to one vial of Zombie VioletTM dye and mix until fully dissolved. Store at -20 °C in 10 μL aliquots. Add appropriate volumes of Fixable Viability solution, with dilutions predetermined by titration experiments (1:1,000) in PBS. Put 100 μL of total volume per well.
Intracellular staining mix
Add antibodies with appropriate dilutions predetermined by titration experiments in intracellular staining buffer. Put 100 μL of total volume per well.
Equipment
BD LSR Fortessa X-20 flow cytometer (BD Biosciences)
Beckman Coulter Allegra X-15R centrifuge (Beckman Coulter); μ-plate carrier for SX4750A (392806)
GentleMACS dissociator (Miltenyi)
Software
FACS Diva software (Version 6.0)
FlowJo software v10
Procedure
Cell suspension preparation from the lungs
Open the thoracic cavity (Figure 1).
Figure 1. Pictures illustrating the different steps of dissection, perfusion, and isolation of the lungs for flow cytometry analysis of immune cells in the context of a respiratory viral infection. (A) Fixation of the mouse on a platform to be stuck in this position. (B) Dissection of the skin and subcutaneous tissue to visualize the upper part of the respiratory system and the thoracic cavity. (C) Opening of the thoracic cavity by cutting the thoracic ribs at the basis to visualize the lungs in place. In the axillary cavity, cut the blood vessels. Insertion of the needle in the right part of the heart (darker side of the heart) and instillation of ice-cold PBS (5 mL) carefully with the syringe. The lungs turn white as the instillation progresses. (D) Isolation of the perfused lung lobes on a dish.
Perfuse the lungs with ice-cold PBS through the right ventricle of the heart.
Collect the lobes of the lung into a pre-warmed GentleMACS C-tube containing 3 mL of complete HBSS, collagenase D (final concentration: 1 mg/mL), and DNase I (final concentration 50 μg/mL).
Process the lungs with a gentleMACS dissociator.
Run the m_lung_01.01 protocol (pre-programmed by manufacturer: 8 s at 168 rounds per run no heat).
Critical: During the dissociation process, some tissue may get stuck to the plastic blades in the C tube. If this occurs, use forceps to transfer the tissue to the liquid prior to centrifugation.
Incubate in water bath under agitation (200 rpm) during 30 min at 37 °C.
Use again the gentleMACS dissociator and run the m_lung_02.01 protocol (pre-programmed by manufacturer: 38 s 2,083 rounds per run no heat) to finely grind the lungs.
Strain the cell suspensions through a 70 μm Falcon filter with wash buffer into a 50 mL centrifuge tube.
Spin down the tubes at 350× g for 5 min at 4 °C.
Discard the supernatant, resuspend with 1 mL of 1× RBC lysis buffer, mix well, and incubate during 5 min.
Fill the tubes with 20 mL of wash buffer.
Spin down the tubes at 350× g for 5 min at 4 °C.
Discard the supernatant and resuspend the pellet into 1 mL of FACS buffer.
Count cells with hemocytometer using trypan blue staining.
Plate 2 million cells into a 96-well U-bottom plate after counting.
Ex vivo stimulation of lung cells
Prepare culture media according to the particular mediators/cytokines to be quantified.
IL-10- and IFNγ-producing T cells: mouse lung cells are stimulated for 4 h at 37 °C in RPMI complete medium containing protein transport inhibitors (2 μM monensin and 2 μM brefeldin), 50 μM β-mercaptoethanol, purified CD28 antibody (1 μg/mL), and purified CD3ε antibody (2 μg/mL).
Granzyme B and Granzyme A producing T cells: mouse lung cells accumulate granzymes for 4 h at 37 °C in RPMI medium containing protein transport inhibitors (2 μM monensin and 2 μM brefeldin) and 50 μM β-mercaptoethanol.
Perforin-1-producing T cells: mouse lung cells are stimulated for 4 h at 37 °C in RPMI medium containing protein transport inhibitors (2 μM monensin and 2 μM brefeldin), 50 μM β-mercaptoethanol, purified CD28 antibody (1 μg/mL), purified CD3ε antibody (2 μg/mL), purified CD49d antibody (1 μg/mL), and APC anti-mouse CD107a (0.5 μg/mL; LAMP-1) antibody.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Resuspend the cell in 200 μL of specific culture media as described above.
Incubate at 37 °C during 4 h and mix regularly by pipetting the solution once an hour.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Wash the cells with 200 μL of ice-cold FACS buffer.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Flow cytometry staining
Block Fc receptors by adding the purified CD16/32 antibody diluted 1:500 in ice-cold FACS buffer. Put 100 μL of total volume per well.
Incubate for 15 min at 4 °C.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Stain for cell surface antigens with antibodies diluted in ice-cold FACS buffer. Put 100 μL of total volume per well. The surface antibodies used (panel, dilution, clone, provider) are provided in Figures 2 and 3.
Incubate for 30 min at 4 °C.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Wash the cells with 200 μL of FACS buffer per well.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Resuspend the cells in 100 μL of fixable viability solution.
Incubate for 30 min at 4 °C.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Discard the supernatant.
Wash the cells with 200 μL of PBS per well.
Centrifuge the cells at 350× g for 5 min at 4 °C.
Resuspend the cells in PFA 4% (diluted in PBS) for 15 min or, alternatively, in PFA 1% overnight at 4 °C.
After the incubation time, centrifuge the cells at 400× g for 5 min at 4 °C.
Discard the supernatant.
Resuspend the cells in 200 μL of permeabilization buffer.
Incubate for 20 min at 37 °C.
Centrifuge the cells at 400× g for 5 min at 4 °C.
Discard the supernatant.
Stain for intracellular cytokines with antibodies diluted in the intracellular staining buffer. The intracellular antibodies used in this study (panel, dilution, clone, provider) are provided in Figures 2 and 3.
Incubate for 20 min at 4 °C.
Centrifuge the cells at 400× g for 5 min at 4 °C.
Discard the supernatant.
Resuspend the cells in 200 μL of intracellular staining buffer.
Centrifuge the cells at 400× g for 5 min at 4 °C.
Discard the supernatant.
Resuspend the cells in 200 μL of FACS buffer.
Analyze by flow cytometry.
Figure 2. Panel 1 antibodies, useful controls, and plate design for Granzyme A, Interleukin-10, and IFN-γ staining. (A). Details of antibodies against surface and intracellular antigens used in the panel 1 (fluorochromes, clones, dilutions, references). (B). Various controls useful for setting up and validating the proposed multicolor panel 1 and example of a 96-well plate design corresponding to the controls used for setting up the panel (C).
Figure 3. Panel 2 antibodies, useful controls, and plate design for Granzyme B and Perforin-1 staining. (A). Antibodies against surface and intracellular antigens used in the panel 2 (fluorochromes, clones, dilutions, references). (B). Various controls useful for setting up and validating the proposed multicolor panel 1 and example of a 96-well plate design corresponding to the controls used for setting up the panel (C).
Staining setup for flow cytometry
In general, there are important considerations that need to be taken into account when performing multiparameter flow cytometry. The Figure 4 provides a schematic representation of the steps in the protocol to be followed.
Figure 4. Protocol flowchart. Schematic representation of the steps in the protocol to be followed and the importance of the controls to be included at key times.
As such, to avoid possible technical variances that may affect data interpretation, it is critical to properly set up the instrument with these following steps by adding technical and biological controls. To circumvent any issues, biological control samples and experimental samples must be processed and analyzed concurrently.
Before each use, calibration of the flow cytometer has to be done by running CST beads provided by BD Biosciences.
We strongly recommend performing titrations of each antibody, single staining controls, and fluorescence minus one (FMO) controls to obtain maximum resolution of cell populations and consistent results across experiments in these multicolor panels. Technical control samples such as FMO controls are essential to help define the boundaries between negative and positive populations. All of these controls allow adjusting the settings of flow cytometer voltages and automatically calculate a compensation matrix, which depends on spillover spreading. We recommend including (in addition of compensation beads) an unstained control by using the cells of interest in order to identify and correct for any intrinsic potential autofluorescence.
In addition to these technical controls, we recommend adding biological controls for each experiment to control the intracellular staining and put the gates properly. As biological controls, we always include control mock mice, ex vivo unstimulated sample controls with and without Golgi Stop (Brefeldin, Monensin). These controls are crucial to validate the signals and to interpret correctly the data obtained by eliminating bias due to technical problems.
Specific features
The protocol we describe in detail here was used to define the functional profile of lung-infiltrating CD4+ T cells during acute respiratory infection induced by a gammaherpesvirus and to understand the intercellular interactions controlling CD4+ T-cell activation. In order to characterize the phenotype of CD4+ cells and to study the mechanisms that control their level of activation and degranulation that take place in vivo, we established an ex vivo stimulation protocol, following which we used multicolor staining and multiparametric flow cytometry. Specifically, we isolated cells from the whole lung (without prior enrichment or sorting) and plated 2 million of cells per well in order to respect the existing in vivo stoichiometry and to preserve the intercellular interactions that participate in the local control of cytotoxic CD4+ T cells.
Data analysis
Regarding the gating strategy, the cells were first selected based on their size and granulosity and the doublets were excluded. We then focused on live cells (not stained with fixable aqua zombie viability dye) and excluded alveolar macrophages based on the CD11c marker expression and autofluorescence in the FITC channel. We subsequently excluded B cells and neutrophils according to lineage markers (Ly6G, CD19) and focused on lin-CD11b- cells. Within these lin-CD11b- cells, T cells were selected based on CD3 expression and the CD4+ and CD8+ T-cell subsets were subsequently selected according to these respective markers.
Detailed information on data analyses appears in the original research article (Maquet et al., 2022) and in the Figure 5 provided in this Bio-protocol. The technical and biological controls included during the process are provided in Figure 4 as a flowchart.
Figure 5. Immunoprofiling of CD4+ T cells isolated from mouse lung after viral infection. (A) Gating strategy used to identify by flow cytometry lung CD4+ T cells isolated from Mock and MuHV-4 infected mice at day 8 after infection. The first step is to distinguish cell populations based on the forward (FSC) and side scatter properties related to cell size and granularity, respectively. After exclusion of doublet cells based on FSC-W vs. FSC-A distribution, dead cells (stained by the fixable viability dye) are identified and eliminated from the gating. Among the living cells, alveolar macrophages (AMs) are gated based on an autofluorescent signal falling in the FITC channel and positive CD11c marker. By excluding these AMs, other immunes cells can be identified as follows: neutrophils gated as live, non-autofluorescent/CD11c-CD19-CD3-Ly6G+CD11b+; monocytes (MOs) gated as live, non-autofluorescent/CD11c-CD19-CD3-Ly6G-CD11b+Ly6C+; CD4+ or CD8+ T cells gated as live, non-autofluorescent/CD11c-CD19-CD3+CD4+ or CD8+, respectively. After selection of the cell subset of interest, the specific gates for the cytotoxic markers and Interleukin-10 are adjusted according to the FMO, biological controls, and positive and negative controls. Representative flow cytometry plots of lung CD4+ T cells, stained for the indicated cytotoxic mediators (B), and for Interleukin-10 (C) after 4 h of anti-CD3ε and anti-CD28 ex vivo stimulation. (D) Expected numbers of live total lung cells, CD4+ T cells, and MOs recovered after lung digestion and multicolor staining for flow cytometry, as described in the methods. (E) Percentage of lung CD4+ T cells expressing the indicated marker of cytotoxicity or interleukin-10 after cell isolation, restimulation, and staining, as described in the methods.
Validation of protocol
This protocol allows for successful isolation of lung cells. Successful dissociation/digestion should yield a viability of 70% or greater. The cell numbers are dependent upon the status of the mice (control or virally infected mice). In Figure 5, we have detailed the gating strategy used to identify the lung cells of interest in the context of the study published in Maquet et al. (2022) on day 8 post infection with MuHV-4, as well as the expected percentages and total numbers of these target cells.
General notes and troubleshooting
General consideration
It is important to highlight that we have described the staining for the intracellular cytokines IL-10 and IFNγ as an example and that other cytokines (in other inflammatory contexts) could be targeted in a similar manner. Indeed, the methods we detail here are directly related to the study we published in Science Immunology, which demonstrated that the absence of recruited monocytes in the lung transforms an asymptomatic viral infection into a severe immunopathology. In particular, in infected WT mice, monocytes provide a balance between cytotoxic and regulatory responses developed by the CD4+ T cells infiltrating the lung. In the specific context of this gammaherpesvirus respiratory infection, it is relevant to quantify the production of IL-10 and IFNγ to determine the functional heterogeneity of CD4+ T cells. This intracellular staining is proposed as an example: the quantification of other cytokines such as IL-6 (Th17), IL-12 (Th1), or the cytokines IL-5 and IL-13 in the context of a Th2 response could be addressed using a similar protocol.
Besides, it is also important to note that treatment of CD4+ T cells with monoclonal anti-CD3 and anti-CD28 antibodies provides a co-stimulatory signal that engages the T-cell receptor (TCR), which can be used for mimicking antigen-induced activation. However, to investigate the presence, activation, and/or proliferation of T cells specific to a given antigen, other protocols should be considered, such as the addition of specific peptides to the stimulation medium or the use of transgenic mice whose CD4+ T cells display a TCR specific to a given antigen.
Critical steps/Troubleshooting
Throughout this protocol, problems may be encountered, and we have compiled a list of critical steps to consider in order to avoid troubleshooting.
Reagents storage
Antibody conjugates and reagents are light and temperature sensitive and must be always protected from light and maintained at room temperature to preserve their stability and shelf life.
Zombie Fixable Viability kit should be stored for up to one year at -20 °C. Unused content in the aliquots should be discarded.
Protocol adjustments
We have tested saving the samples in the fridge overnight after PFA (1% instead of 4% with 15 min fixation) in case samples cannot be immediately acquired by the flow cytometer. Cells can be stored overnight at 4 °C in 1% PFA protected from light, and permeabilized and intracellular-stained the day after, without alteration of the results.
Due to cross-laser excitation, some dyes spill into other channels inducing spillover-spreading errors. Alternatively, and depending on the required panel, antibodies-conjugated fluorochromes could be swapped with another dye. For more information on spreading, use BD Biosciences Spectrum Viewer.
Depending on the biological questions, intracellular staining of Granzymes A and B can be performed directly without any pre-incubation with protein transport inhibitors.
Cautionary points
Lung samples should be kept as much as possible on ice until stimulation at 37 °C to preserve viability. After digestion, the only step that has to be performed at room temperature is the RBC lysis during 5 min. Use ice-cold solutions.
Saponin permeabilization is a reversible step, and intracellular staining must be carried out in the presence of permeabilization buffer.
Respect the washing steps correctly during staining and permeabilization.
Acknowledgments
The protocol was adapted from the previously published paper: Maquet et al. (2022). This work was supported in part by the F.R.S./FNRS (research fellow for C.M.; research associate support for B.M.; F.R.S./FNRS “incentive grant for scientific research to B.M., 40003542”), in part by the ERC Starting Grant (to B.M.) (ERC-StG-2019 VIROME, ID:853608), in part by the grant from Leon Fredericq Foundation (to C.M.), and by the EOS joint program of F.R.S./FNRS-FWO (EOS ID: 30981113) (to L.G.).
Competing interests
The authors declare no competing interests.
Ethical considerations
All work in the development of this protocol was approved by the Committee on the Ethics of Animal Experiments of ULiège (permit numbers 2015 and 2215).
References
Maquet, C., Baiwir, J., Loos, P., Rodriguez-Rodriguez, L., Javaux, J., Sandor, R., Perin, F., Fallon, P. G., Mack, M., Cataldo, D., et al. (2022). Ly6Chimonocytes balance regulatory and cytotoxic CD4 T cell responses to control virus-induced immunopathology. Sci. Immunol. 7(73): eabn3240.
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© 2023 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-based analysis > Flow cytometry
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4,816 | https://bio-protocol.org/en/bpdetail?id=4816&type=0 | # Bio-Protocol Content
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Peer-reviewed
A Post-translational Modification–enhanced Pull-down Method to Study Degron Domains and the Associated Protein Degradation Complexes
PS Pierluigi Scalia
SW Stephen J. Williams
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4816 Views: 816
Reviewed by: Ralph Thomas BoettcherQingliang Shen Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in International Journal of Molecular Sciences Apr 2023
Abstract
The identification and characterization of the ubiquitin E-ligase complexes involved in specific proteins’ degradation via the ubiquitin-proteasome system (UPS) can be challenging and require biochemical purification processes and in vitro reconstitution assays. Likewise, evaluating the effect of parallel phosphorylation and ubiquitination events occurring in vivo at dual phospho/ubiquitin-regulated motifs (called Phospho-Degrons or pDegrons) driving UPS degradation of the targeted protein has remained elusive. Indeed, the functional study of such E1-E2-E3 complexes acting on a protein-specific level requires previously or otherwise acquired knowledge of the nature of such degradation complex components. Furthermore, the molecular basis of the interaction between an E3 ligase and its pDegron binding motif on a target protein would require individually optimized in vitro kinase and ubiquitination assays. Here, we describe a novel enzymatically enhanced pull-down method to functionally streamline the discovery and functional validation of the ubiquitin E-ligase components interacting with a phospho-degron containing protein domain and/or sub-domain. The protocol combines key features of a protein kinase and ubiquitination in vitro assay by including them in a pull-down step exerted by a known or putative pDegron-tagged peptide using the cell extracts as a source of enzymatically active post-translational modification (PTM) modifying/binding native proteins. The same method allows studying specific stimuli or treatments towards the recruitment of the molecular degradation complex at the target protein’s phospho-degron site, reflecting in vivo–initiated events further enhanced through the assay design. In order to take full advantage of the method over traditional protein–protein interaction methods, we propose to use this PTM-enhanced (PTMe) pull down both towards the degradation complex discovery/ID phase as well as for the functional pDegron recruitment validation phase, which is the one described in the present protocol both graphically and in a stepwise fashion for reproduceable results.
Key features
• Suitable to study UPS-regulated (a) cytosolic and/or nuclear proteins, (b) intracellular region of transmembrane proteins, and (c) protein sub-domains bearing a known/putative pDegron motif.
• Requires a biotin-tagged recombinant version of the target protein and/or sub-domain.
• Allows the qualitative and quantitative analysis of endogenous ubiquitin (Ub) E-ligases recruitment to a known or putative pDegron bearing protein/sub-domain.
• Allows simultaneous testing of various treatments and/or conditions affecting the phosphorylative and/or ubiquitylation status of the studied pDegron bearing protein/sub-domain and the recruited factors.
Graphical overview
Keywords: UPS Ubiquitin-proteasome system Ubiquitin E-ligase pDegron motif Phospho-degron motif PTM Post-translationally modified PTMe PTM-enhanced
Background
Regulation of protein degradation is particularly relevant for the cell to control the expression levels of regulatory proteins along with the underlying biological processes. The major known cellular proteostasis mechanism is mediated by the ubiquitin-proteasome system (UPS) and uses a dedicated set of widespread enzymes called ubiquitin E-ligases. Their role is to flag specific proteins by adding a ubiquitin moiety to specific lysine residues that will then be further identified, targeted, and complexed by E-ligases (typically initiated by E3, followed by E2 and E1 types) and ultimately degraded by the UPS (reviewed in [1]). The essential amino-acid sequence allowing ubiquitin-targeted protein degradation has been called Degrons [2–4]. Degrons in which phosphorylation residue(s) play either a permissive or inhibitory role towards degradation are best defined as phospho-(regulated) Degrons and are ubiquitously found in regulatory proteins involved in growth and developmental processes [4,5]. Phospho-Degrons in which the alternative phosphorylation vs. ubiquitylation requirement establishes a biological switch are also referred as phospho-inhibited degrons [6]. Phospho-Degron motifs can be predicted by the parallel presence of in vivo PTM-modified Ser/Thr/Tyr amino-acid residues within a relative short distance from an in vivo modified lysine [7–9]. At present, the post-translational modification (PTM) status of putative Phospho-Degrons can be exerted by independent assays looking either at the ubiquitylation (in vitro ubiquitination assay) or phosphorylation status (through in vitro kinase assays). We have designed an integrated strategy and method to enhance specificity towards both (a) identifying a putative phospho-degron and simultaneously (b) functionally confirm the ID of the ubiquitin E1,2,3-ligases recruited by the phospho-degron-bearing protein/domain. Central to the described method is the synthesis of a biotin-tagged protein domain/subdomain or peptide bearing the native ubiquitin-binding motif amino acidic sequence for the putative phospho-Degron. This recombinant biotin-tagged protein domain/peptide is used as the bait component of the PTM-enhanced (PTMe) pull-down method. The method relies on the ex vivo–enhanced phosphorylation and ubiquitylation (PTMs) events induced in vivo. This bears increased advantages over performing separate co-immunoprecipitations (co-IP)/far westerns and individual kinase or ubiquitylation assays in immunoprecipitated proteins, as shown in Figure 1. The method can indeed be used prior to further refining individual protein interactions within a protein degradation complex, requiring deletion and single-point recombinant mutants, therefore sparing lengthy preparatory work for the expression and purification of the individual recombinant proteins before having clarified their functional involvement in the degradation complex. PTMe pull-down conditions can also be used for proteomics profiling in the degradation complex discovery phase, but the LC-Ms/Ms approach may not be available to all laboratories. We have successfully used this method towards the characterization of a growth factor–regulated phospho-Degron that we initially identified in an angiogenic/metastatic membrane receptor kinase of the Ephrin family, over-expressed in malignant mesothelioma cells [6]. We have adopted the described protocol in combination with immune-enriched proteomics towards further identifying the protein complex for the degradation of the angiogenic kinase EphB4 [6], whose inhibition by an IGF-II autocrine signal determines its over-expression in a malignant mesothelioma cell line [5].
Figure 1. Immuno-blotting detection branch of the post-translational modification–enhanced (PTMe) pull-down method and comparison with traditional co-immune precipitation (co-IP) on the same cellular extract batch from treated vs. untreated cells. An aliquot of the material used in each condition was run on a gel and used for Coomassie stain. The asterisk-marked arrows refer to the complex components for which the method can provide results not observed with traditional co-IP as specified under background. The right-side image is part of the study by Scalia et al. (2023) [6], in which the method was first used. *Please note that two different areas of the same Coomassie gel are shown for comparing the post-normalization protein amount of the same cell extract batches used for both the displayed co-IP and PTMe pull-down experiments.
Figure 2. Membrane partition suggested for parallel analysis of ubiquitin (Ub)-E-ligases detection/pDegron recruitment study using post-translational modification–enhanced (PTMe) pull-down assay. Membrane partition is suggested given the relevance of studying same-cell extract batches deriving from individual in vivo treatment conditions and underlying PTMe pulled-down complexes from individual experiment. Proper control suggested between conditions is a Coomassie stain of equal amounts of (protein-normalized) cell extract input resolved by SDS-PAGE.
Materials and reagents
Materials include disposable sterile items like tubing and cell culture supplies obtained from Thermo Inc. (Waltham, MA, USA). Reagents, along with the working concentrations used, have been conveyed in Table 1.
Table 1. Reagents
Name Reagent Specs Working conc. Provider Product#
Putative PD-synthetic peptide (N/C-biotin tagged) Assay substrate/bait
HPLC-grade
50 μg/ pull-down reaction (rxn) New England Peptides (MA)
custom
synthesis
Ubiquitin, human synthetic Ub-moiety donor 1 μg/rxn Boston Biochemical U-100H
Tris-HCl pH buffer 10 mM Sigma-Aldrich 77-96-1
PBS pH buffer 1× to final rxn vol. Sigma-Aldrich D1408
ATP phosphate moiety donor 4 mM Sigma-Aldrich A1852
DTT reducing agent 4 mM Sigma-Aldrich D9779
MnCl2 prot kinase cofactor (Mn++) 2 mM Sigma-Aldrich 1375137
MgCl2 prot kinase cofactor (Mg++) 2 mM Sigma-Aldrich 1374248
NP40 Rxn/pull-down facilitator 0.05% Sigma-Aldrich 9036-19-5
D5/W5 inhibitors cocktail Protease inhibitors See Recipe 2
Kaleidescope prestained protein markers
broad range visible
protein markers
6 µL/lane Bio-Rad
1610324
WB-Master Protein Standard
Chemiluminescence
Sensitive Markers
10 µL/lane Genscript
M00521
SuperSignalTM West Femto Chemiluminescent Substrate
Chemiluminescence
Ultrasensitive
Reagents system
0.5 mL/reagent/
Membrane strip
Fisher
34095
Antigen (/Ab Clone) Animal source/ clonality Working concentration Provider Clone /product#
Ubiquitin (P4D1) mouse monoclonal 1:1,000 Santa Cruz P4D1/sc8017
Anti-mouse IgG-HRP chicken monoclonal 1:2,000 Santa Cruz sc2954
Anti-IgGk-LC-HRP Mouse monoclonal 1:1,000 Santa Cruz sc156102
Streptavidin mag beads n/a 2 μL/mL Genscript L00936
Recipes
The key component of the PTMe pull-down method stands in the homonymous reaction/incubation mixture described below.
PTMe pull-down reaction mixture
PTMe Reaction (rxn) Stock conc. Volume per rxn Final conc.
N-Biotin pDegron-peptide coupled w/STP-magnetic beads (see text) n/a 50 μg-STP bound/rxn
Tris-HCl, pH 7.5 80 μL 10 mM
DTT, 100 mM 8 μL 4 mM
ATP, 100 mM 8 μL 4 mM
MnCl2,100 mM 4 μL 2 mM
MgCl2, 100 mM 4 μL 2 mM
Ubiquitin, human 1 µg/µL 1 μL 1 μg
Cell extracts (2.5–3.5 µg/µL) Up to 95 μL 0.2–0.3 mg
PBS, 0.05% NP40 +D5/W5 protease inhibitors mix
To 200 μL (as needed) 1×
Protease and related activities inhibitors stocks
Water soluble inhibitors (W5)
Inhibitor [Stock]
µL stock to produce [working]
for 5 mL pull-down buffer
[Working]
NaVO3 0.2 M 25 µL 1 mM
Benzamide 0.25 M 40 µL 2 mM
β-glycerophosphate 1 M 50 µL 10 mM
NaF 1 M 50 µL 10 mM
DTT 1 M 5 µL 1 mM *
Sub-total volume (165 µL) -
Mol Grade H2O to 10× 483.5 µL n/a
PBS, 0.05% NP40 n/a 4,500 µL 1×
DMSO soluble inhibitors stock (D5)
Inhibitor [Stock]
µL stock to produce [working]
5 mL pull-down buffer
[Working]
Pepstatin 10 mg/mL 5 µL 10 µg/mL
Leupeptin 10 mg/mL 5 µL 10 µg/mL
Aprotonin 10 mg/mL 5 µL 10 µg/mL
Okadaic acid 100 µM 5 µL 100 nM
PMSF 0.5 M 5 µL 0.5 mM
Sub-total volume (25 µL) -
DMSO for 10× 487.5 µL n/a
PBS, 0.05% NP40 n/a 4,500 µL 1×
It is advisable to prepare a larger volume of 10× W5 and D5 stocks, make 0.5 mL aliquots, and save them for single fast thawing (water bath, room temperature). Keep at -20 °C for three months or at -80 °C for six months.
*DTT is added separately to the final buffer containing water-soluble and DMSO-soluble protease inhibitors, to produce a buffer containing 1 mM DTT.
PBST is the antibody probing buffer and is prepared starting by PBS at 1× dilution in sterile deionized H2O with Tween 20 detergent at 0.001% (w/w).
Note: All reagents were purchased from Sigma-Aldrich, St. Louis, MO, USA
Equipment
Cell culture facility (SterilGard III tissue culture hood; CO2 Auto Zero tissue culture incubator, Thermo Scientific, Waltham, MA or comparable models)
4 °C refrigerator or same temperature cold room
-20 °C and -80 °C freezers
Benchtop microcentrifuge supporting 14,000× g spins (e.g., Eppendorf 5418 model or comparable)
Gentle orbital rotation instrument (Boekel Orbital rotator, Feasterville, PA or comparable)
Thermo block with heating capability of 95 °C (Thermo Digital Block Dry Heater, Thermo Scientific, Waltham MA or comparable)
Protein electrophoresis/western blot equipment (NOVEX XCell II and NOVEX TransBlot system, Thermo Scientific, Waltham MA or comparable)
Gel image multi-signal detection system (Odissey XF imaging system, LICOR, Lincoln, NE or comparable). The suggested equipment brands are only indicative and not considered as limiting the outcome of the described protocol
Procedure
The protocol has been conveyed in three sequential workflows that can take place on sequential days, with the exception of Workflow B that must be continued until proteins transfer to PVDF membrane. Day three can start from the post-overnight membrane blocking step in workflow C. The author acknowledges that the optimization of the present protocol may be favored by previous investigator’s hands-on experience with protein immunoprecipitation and protein kinase or other PTM-based in vitro assay. Therefore, in order to speed up the optimization of the present method, it is advised getting familiar with the mentioned procedures prior to dive straight into the integrated approach described herein.
Benchwork considerations (Figure 3)
Figure 3. Stepwise procedure, workflow A
An N-/C-terminally tagged protein domain bearing a pDegron or a corresponding Biotin-conjugated synthetic peptide is used as a substrate/bait (New England Peptides, Gardner, MA) in a modified pull-down/ubiquitination/phosphorylation in vitro assay designed to enhance the specific post-translational modifications (ubiquitination and phosphorylation) induced by the ubiquitin-ligase/phosphorylation native kinase enzymatic activities in the cell extracts from either treated cells or those obtained from parallel untreated cells.
The pDegron protein/peptide bait can be either synthetized by a commercial provider or generated through recombinant DNA cloning and protein purification methods, depending on the length of the domain and on the laboratory cost/time benefit evaluations. The pDegron synthetic peptide was reconstituted in sterile molecular biology-grade water as per manufacturers’ recommendation at a 1 mg/mL concentration, aliquoted in sterile microfuge tubes, and stored at -20 °C until use. We did not find signs of degradation or loss within six months but have not tested longer storage times. The purified phospho-Degron-tagged peptide is therefore used as a bait for the in vitro protein complex capture assay in the presence of pre-treated cell lysates as a source of the native enzymatic factors targeting the degron peptide (namely, protein kinases or PKs and Ub-ligases), along with defined reconstituted inorganic components aimed to enhancing both protein kinase and ubiquitylation activities triggered by the in vivo treatment during the in vitro pull-down incubation step. In the case of the pDegron characterized by the authors through the described method, its putative presence in the studied protein kinase (PK) was first demonstrated by looking at the differential effect of a cancer-secreted growth factor signal on the (Cyclohexymide-sensitive) protein levels on the over-expressed PK [5]. The same GF signal was then specifically correlated to the phosphorylation and ubiquitination status of the C-terminal region of the PK, allowing the definition of the pDegron as a phospho-inhibited type. This means that the pDegron is active and leads to the protein UPS-mediated degradation upon de-phosphorylation. Therefore, the recruitment of degradation factors (namely the E3-E2-E1-binding ubiquitin ligases) to the pDegron (peptide mimicking the C-tail region of the PK) was enhanced through a comparative experimental design (as shown in Figure 1), which included untreated cell extracts (with intact GF signal, inhibited pDegron, and over-expressed target protein degradation rescue status) vs. GF-signal-deprived cell extracts (with dephosphorylated pDegron and recruited degradation factors) [6].
Benchwork considerations (Figure 4)
Figure 4. Stepwise procedure, workflow B
For protocol validation and during the discovery phase, two cell pellets (for untreated and treated conditions, respectively) were pooled from 4× 150 mm plates at semi-confluency and used to generate the two cell extract batches (with several frozen aliquots generated to avoid multiple freezing/thawing cycles) displayed in Figure 1. Specifically, a single preparation (of treated and untreated cells) was used towards performing, at different times, (a) co-IP, (b) PTMe pull down, and (c) Ms/Ms profiling from a single biological source. The traditional co-IP method demanded the highest starting protein amount (we used up to 0.5–0.7 mg of cell extract per IP condition for the results shown in Figure 1), followed by the PTMe pull down (0.2–0.3 mg/condition); the most sensitive was the immuno-enriched LC-Ms/Ms (0.05–0.150 mg/condition).
Although we obtained good results on the PTMe pull down with less than 100 μg whole cell extracts for the detection of binding proteins through mass spectrometry, we suggest increasing the protein amount to the maximum usable volume of normalized protein extracts for the pull-down reaction step towards immuno-blot detection (0.2–0.3 mg of enzymatically active protein extracts). Enzymatically active cell extracts were obtained by (1) adding the described protease inhibitors (Recipe 2) to the lysis buffer, (2) performing all cell-free steps on ice, (3) adopting fast freezing (liquid nitrogen immersion) and slow thawing (on ice) of the tested cell extracts, and (4) avoiding multiple freezing/thawing cycles.
First, combine 50 μg of Nt-Biotin-tagged synthetic pDegron-bearing peptide with 6.25 μL of Streptavidin (STP) magnetic beads (Genscript, Piscataway NJ) for 2 h at 4 °C with rotation, followed by a PBS 0.05% NP40 wash. Then, use the STP beads–coupled N-Biotin peptide bait to assemble a 200 μL mixture containing 0.75 mg whole-cancer cell extracts (enzymatically active) along with 10 mM Tris pH 7.5, 4 mM ATP, 2 mM MnCl2, 2 mM MgCl2, 1 μg human recombinant Ubiquitin, and protease/phosphatase inhibitors at the final concentrations summarized in Recipe 2.
In order to determine the optimal amount of starting cell extracts needed for a successful result, an aliquot of the pulled down product was used in parallel with 2–5 μL of total cell extracts for SDS-PAGE followed by gel staining with either Coomassie blue or Silver Stain. Protein–protein interactions along with cell extracts–induced phosphorylation/ubiquitination post-transcriptional modifications were allowed to occur by incubating the mixture for 24–48 h at 4 °C with gentle rotation.
At the end of the pull-down incubation, use the microfuge tubes containing pDegron beads–bound proteins either (a) without wash for LC-Ms/Ms analysis or (b) transferred to a thermo block pre-equilibrated at 30 °C for further incubation, enhancing the cell extracts endogenous protein kinase and ubiquitination events. Alternatively, transfer an aliquot (e.g., 1/3 volume) of the PTMe-complexed beads into a microfuge tube and wash twice with 1 mL of ice-cold PBS to stop the reaction; use the beads for LC-Ms/Ms (as displayed in the graphic overview). The need for PTMe to detect native proteins bound to our pDegron through mass spectrometry was mitigated by the ex vivo treatment of our cells to increase the specific proteins interaction to our pDegron [6]. Nonetheless, in those cases where no biologic background is available about the regulation of the putative pDegron domain, PTMe for both the proteomic step and the immuno-blot detection may become the preferred approach.
After incubation in step B3, add Stop/SDS Gel loading (Laemmli) buffer and DTT (1 mM final concentration) to the beads and perform denaturation at 95 °C for 3 min. Following brief vortexing and full-speed centrifugation, load the supernatant sample on a polyacrylamide gel (fixed 10%, 12%, or 8%–16% gradient) for SDS-PAGE.
Although the workflow ends with protein separation on SDS-PAGE for conceptual simplicity, for practical reasons it must continue to protein transfer to the suggested solid membrane (PVDF), in order for the protocol to be suspended as suggested in workflow C (Figure 5).
Benchwork considerations (Figure 5)
Figure 5. Stepwise procedure, workflow C
Transfer resolved proteins onto PVDF (which requires pre-activation with soaking for 5 s in organic solvent, as suggested by the manufacturer, followed by equilibration in PBST buffer for 2–5 min).
Block non-specific binding sites through incubation in PBST 5% BSA overnight at 4 °C.
To obtain the most from each cell extract preparation and single experiment, cut membrane strips from the post gel-to-membrane transfer solid support (PDVF) as shown in Figure 2 and perform MW-specific immunoblotting on each strip immediately after step C3. Membrane partition and differential p-Degron binding components immune detection of the same extracts (rate limiting or PTMe) require former establishment of the identity and MW of the protein complex components, through either the suggested enriched proteomic approach or other proteomic workflows.
The use of enhanced luminescence signal (we used SuperSignal West Femto kit from Pierce) for detection of femto-molar amounts of PVDF-transferred protein turned out to be a key factor for the results of the described method (see under troubleshooting tips, Table IV).
The PTMe pull-down method described herein was optimized using a tagged peptide of the studied protein bearing a previously identified phospho-Degron motif [5].
Upon denaturation of the pull-down complex bound to the studied (tagged) phospho-Degron peptide, proceed to either a proteomic study or a co-IP assay, by resolving the pull-down endogenous proteins by SDS-PAGE and by detecting the resolved factors by western blotting.
In case of using the PTMe pull down towards protein complex components ID by LC-Ms/Ms profiling (not part of the present protocol), we suggest adding an ex vivo or in vitro reversible cross-linking step, as described in Scalia et al. (2023) [6].
Validation of protocol
Given the biological complexity driving the formation of degradation complexes and the context-specific variables involved, the direct comparison of traditional co-IP with pull down using the same cell extracts is a mandatory strategy we suggest for protocol validation. The case-specific advantages observed will drive the choice. As shown in Figure 1, PTMe pull down provided greater specificity for the treatment-induced recruitment of all the degradation complex components, while co-IP was accurate only for two of the pDegron binding components tested (identified by previous proteomic screening), which included the complete set of Ub-E-ligases along with a key chaperone/unfoldase co-factor involved in the underlying process. The advantage of validating all components biologically responsible for the degradation of the studied protein in a single experiment using a single cellular extract/preparation/treatment per se justifies the implementation of the proposed PTMe pull-down method over other protocols.
General notes and troubleshooting
The described protocol is best suited for proteins and sub-domains with predicted pDegron function. This can be accomplished through preliminary sub-domain comparisons with existing protein domains known to be ubiquitylated in vivo (the best is pre-tested by western blot for K48 lysine ubiquitylation using available commercial antibodies). Troubleshooting has been conveyed in Table 2 below.
Table 2. PTMe pull-down method troubleshooting
Problem Potential technical issues Solution
Immunoblot signal of the target proteins pulled down by the phospho-Degron bait
in PTMe assay not optimal
Spurious protein degradation
Include protease inhibitors in your cell extract/lysis buffer (see Recipe 2).
Low amount or expression of target protein(s) of interest in lysates Confirm presence of expected or proband protein bands at specific MW range by either silver stain or Coomassie blue method.
Insufficient gel visualization of expected band
Increase the amount of pull-down proteins by:
1) increasing the starting amount of protein cell extracts and in parallel;
2) increasing the amount of STP-coupled biotin-tagged synthetic peptide bait.
Potential consumption of ECL reagent in the PVDF membrane from other proteins than the one studied Cut and perform western blot (WB) of single strip of transfer membrane corresponding to the protein of interest as in Figure 2.
Amount of pull-down protein factors lower than detection limit of chemiluminescent detection system used Use femto-molar grade WB luminescence detection kit (when using HRP-conjugated secondary Ab) or use IR-labeled secondary Ab.
Acknowledgments
No public funding was used for this study and the study relative to the described protocol.
Method first reported in Scalia et al. (2023) [6]; discovery and design of the pDegron domain used for the present protocol development: Scalia et al. (2019) [5].
Competing interests
The authors declare no competing interests.
References
Hershko, A. and Ciechanover, A. (1992). The ubiquitin system for protein degradation. Annu. Rev. Biochem. 61(1): 761–807. doi: 10.1146/annurev.bi.61.070192.003553
Van Roey, K., Uyar, B., Weatheritt, R. J., Dinkel, H., Seiler, M., Budd, A., Gibson, T. J. and Davey, N. E. (2014). Short Linear Motifs: Ubiquitous and Functionally Diverse Protein Interaction Modules Directing Cell Regulation. Chem. Rev. 114(13): 6733–6778. doi: 10.1021/cr400585q
Davey, N. E. and Morgan, D. O. (2016). Building a Regulatory Network with Short Linear Sequence Motifs: Lessons from the Degrons of the Anaphase-Promoting Complex. Mol. Cell 64(1): 12–23. doi: 10.1016/j.molcel.2016.09.006
Hunter, T. (2007). The Age of Crosstalk: Phosphorylation, Ubiquitination, and Beyond. Mol. Cell 28(5): 730–738. doi: 10.1016/j.molcel.2007.11.019
Scalia, P., Pandini, G., Carnevale, V., Giordano, A. and Williams, S. J. (2019). Identification of a novel EphB4 phosphodegron regulated by the autocrine IGFII/IRA axis in malignant mesothelioma. Oncogene 38(31): 5987–6001. doi: 10.1038/s41388-019-0854-y
Scalia, P., Merali, C., Barrero, C., Suma, A., Carnevale, V., Merali, S. and Williams, S. J. (2023). Novel Isoform DTX3c Associates with UBE2N-UBA1 and Cdc48/p97 as Part of the EphB4 Degradation Complex Regulated by the Autocrine IGF-II/IRA Signal in Malignant Mesothelioma. Int. J. Mol. Sci. 24(8): 7380. doi: 10.3390/ijms24087380
Tang, X., Orlicky, S., Liu, Q., Willems, A., Sicheri, F. and Tyers, M. (2005). Genome‐Wide Surveys for Phosphorylation‐Dependent Substrates of SCF Ubiquitin Ligases. Meth. Enzymol.: 433–458. doi: 10.1016/s0076-6879(05)99030-7
Hao, B., Oehlmann, S., Sowa, M. E., Harper, J. W. and Pavletich, N. P. (2007). Structure of a Fbw7-Skp1-Cyclin E Complex: Multisite-Phosphorylated Substrate Recognition by SCF Ubiquitin Ligases. Mol. Cell 26(1): 131–143. doi: 10.1016/j.molcel.2007.02.022
Nash, P., Tang, X., Orlicky, S., Chen, Q., Gertler, F. B., Mendenhall, M. D., Sicheri, F., Pawson, T. and Tyers, M. (2001). Multisite phosphorylation of a CDK inhibitor sets a threshold for the onset of DNA replication. Nature 414(6863): 514–521. doi: 10.1038/35107009
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© 2023 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 > Ubiquitinylation
Molecular Biology > Protein > Protein-protein interaction
Molecular Biology > Protein > Targeted degradation
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Double Staining with Fluorescent Tracers to Determine Myeloid Cell Migration of Leishmania-infected Cells from Mouse Skin to Lymphatic Tissues by Flow Cytometry
AU Ashanti C. Uscanga-Palomeque
EO E. Yaneth Osorio
PM Peter C. Melby
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4817 Views: 697
Reviewed by: Alexandros AlexandratosMaria Agallou Anonymous reviewer(s)
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Cited by
Original Research Article:
The authors used this protocol in PLOS Neglected Tropical Diseases Jan 2023
Abstract
Immune cell trafficking in steady-state conditions and inflammatory cell recruitment into injured tissues is crucial for the surveillance of the immune system and the maintenance of body homeostasis. Tracking the cell journey from the infection site in the skin to lymphoid tissues has been challenging, and is typically determined using fluorescent cell tracers, antibodies, or photoconvertible models. Here, we describe the detailed method to track Leishmania-infected myeloid cells migrating from the skin to lymphatic tissues by multiparametric flow cytometry. These methods involve labeling of infective Leishmania donovani parasites with fluorescent cell tracers and phenotyping of myeloid cells with fluorescent antibodies, to determine the infection status of migratory myeloid cells. We also describe the detailed protocol to trace donor monocytes transferred intradermally into recipient mice in Leishmania donovani infection. These protocols can be adapted to study skin-lymphoid tissue migration of dendritic cells, inflammatory monocytes, neutrophils, and other phagocytic myeloid cells in response to vaccine antigens and infection.
Key features
• Cell-tracking of cell-trace-labeled parasites and monocytes from the skin to lymphatic tissues after transference into donor mice.
• Identification of migratory cells labeled with fluorescent cell tracers and antibodies by flow cytometry.
• Isolation, labeling, and transference of bone marrow monocytes from donor mice into the skin of recipient mice.
• Description of a double-staining technique with fluorescent cell tracers to determine cell and parasite dissemination from the skin to lymphoid tissues.
Graphical overview
Overview of the methods to trace the migration of Leishmania and monocytes from the skin to lymphatic tissues by flow cytometry. Infective metacyclic promastigotes (from axenic culture) and monocytes (isolated from the bone marrow of donor mice) are labeled with fluorescent cell tracers. After intradermal injection into the test mouse (1, 2), migratory cells and infected cells are isolated from the skin and lymphoid tissues of the test mouse. These cells are then labeled with fluorescent antibodies against myeloid cells and recognized according to the differential excitation/emission wavelengths of the fluorochromes by flow cytometry.
Keywords: Leishmania donovani Pathogen labeling Myeloid cells Fluorescent staining Cell tracking In vivo Flow cytometry Adoptive transfer of monocytes
Background
The migration of immune cells is a fundamental function of the immune system in health and disease. Immune cell movement through the lymphatic network plays a crucial role in immune surveillance, the initiation of the immune response, and the development of tolerance. Innate immune cells, such as neutrophils, macrophages, and dendritic cells capture antigens and capture and kill pathogens, activating the adaptative immune response at the draining lymph node (Hampton and Chtanova, 2019). However, intracellular parasites have also developed strategies to circumvent the immune response, survive, and disseminate through the lymphatics to other tissues (Hampton and Chtanova, 2019).
Cell migration can be studied by diverse approaches with different drawbacks. Time-lapse microscopy tracks short fragments of the individual cell migration and generates large amounts of digital data, making the analysis complicated, even with access to software tools for automated cell tracking (Al-Zaben et al., 2019). Intravital microscopy (IVM) is useful for the visualization of cell trafficking, proliferation, and cell death at single-cell resolution level (Maiorino et al., 2022). However, IVM is an invasive method that can generate local inflammation and tissue damage (Maiorino et al., 2022). Most IVM techniques are based on fluorescence microscopy, but in the last two decades, different biological processes have been explored using IVM together with two/three-photon microscopy to avoid dye use. However, the long duration of image acquisition can cause photodamage and an inadequate spatial co-registration between sequentially detected signals (You et al., 2018). Currently, photoconvertible transgenic mice are being used for cell migration studies; however, the establishment of photoactivation conditions is tedious, requiring the coupling of the photoactivation technique and instrumentation for signal detection. The most popular photoactivation/conversion model, Kaede mice, helps to understand cell migration at single-cell level or by invasive intracranial imaging. However, this technique is not suitable for models involving photoactivation of cells in the skin or for studies requiring deep tissue photoactivation (Tomura et al., 2021). Similarly, in our hands, the photoactivable model R26-PAGFP-LysMcre [B6.Cg-Gt(ROSA)26Sortm1(CAG-PA-GFP)Rmpl/J mice mated with B6.129P2-Lyz2tm1(cre)Ifo/J] was not suitable for identifying migratory cells photoactivated in the skin (Figure S1, Video S1 and Table S1). Therefore, fluorescent cell tracers are the most inexpensive, straightforward technique to study cell migration from skin to lymphatic organs.
Fluorescent cell trackers are powerful tools that enable the direct visualization of biological processes, such as cell proliferation, cell migration, and cell–cell interactions (Halabi et al., 2020). Thus, it is possible to stain cells or pathogens with fluorescent tracers to follow their migration through the lymphoid tissues of the host. The use of cell trackers coupled with flow cytometry makes it possible to identify the infected cells and determine the cell populations carrying parasites from the skin (Ibrahim et al., 2013). The higher specificity, sensitivity, and robustness of this method are reached by the combination with flow cytometry. The protocol described here was applied to the study of infection status and migration of innate immune cells (monocytes, dendritic cells, and neutrophils) to lymph node and spleen of after-skin-infection with Leishmania spp. (Osorio et al., 2023).
Materials and reagents
Biological materials
Balb/c mice (4–7 weeks old) (Envigo, Inotiv)
Wildtype or mCherry Leishmania donovani metacyclic promastigotes (IS strain; MHOM/SD/00/S-2D) (Osorio et al., 2023)
Reagents
RPMI 1640 medium (Thermo Fisher Science, GibcoTM, catalog number: 11875093)
M199 medium (Thermo Fisher Science, GibcoTM, catalog number: 11150067)
Penicillin-streptomycin solution (Thermo Scientific, GibcoTM, catalog number: 15140163)
Fetal bovine serum (FBS), inactivated by heating at 56 °C for 20 min (Thermo Scientific, GibcoTM, catalog number: 16000044)
0.4% Trypan Blue solution (Sigma-Aldrich, catalog number: T8154)
Adenine (Sigma-Aldrich catalog number: A8626)
Hemin (Millipore, Sigma, catalog number: H9039)
HEPES buffer solution 1 M (Millipore, Sigma, catalog number: 83264)
Sodium chloride (NaCl) (Millipore, Sigma, catalog number: S9888)
Potassium chloride (KCl) (Millipore, Sigma, catalog number: P3911)
Magnesium chloride hexahydrate (MgCl2·6H2O) suitable for cell culture (Millipore, Sigma, catalog number: M2393)
Calcium chloride (CaCl2) suitable for cell culture (Millipore, Sigma, catalog number: C7902)
Collagenase D (Millipore, Sigma, Roche, catalog number: 11088858001)
DNase I (Millipore, Sigma, Roche, catalog number: 10104159001)
LiberaseTM TL (Millipore, Sigma, Roche, catalog number: 5401020001)
Accutase cell detachment solution (STEMCELL Technologies, catalog number: 07920)
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7030)
10× phosphate-buffered saline (PBS) (Corning®, catalog number: 46-013-CM)
Dimethyl sulfoxide (DMSO) (hybridoma grade) (Sigma-Aldrich, catalog number: D2650)
MojoSortTM mouse Ly-6G Selection kit (BioLegend, catalog number: 480123)
MojoSortTM buffer (5×) (BioLegend, catalog number: 480017)
MojoSortTM magnet (BioLegend, catalog number: 480020)
Anti-CD11b magnetic particles DM Clone M1/70 (RUO) (BD IMagTM, catalog number: 558013)
Lectin from Arachis hypogaea (peanut) PNA agglutinin (Sigma-Aldrich, catalog number: L-08881)
Dibutyl phthalate (Sigma-Aldrich, catalog number: 524980)
1-step Fix/Lyse solution (10×) (eBioscienceTM, catalog number: 00533354)
BD IMagTM buffer (10×) (BD, catalog number: 552362)
RBC lysis buffer (10×) (BioLegend, catalog number: 420301)
TruStain FcXTM (anti-mouse CD16/32) antibody (BioLegend, catalog number: 101320)
Fixable Viability Dye eFluor 455UV (eBioscienceTM, catalog number: 65-0868)
Antibodies for flow cytometry (see Table 1)
Cell tracer dyes (see Table 2)
Table 1. Antibodies and probes for ex vivo staining of myeloid cells by flow cytometry
Antibody name (Concentration of stock) Ex/Em (nm)
Working concentration
μg/tube (μL)
Clone Brand and catalog number
Anti-mouse CD3- FITC (0.5 mg/mL) 495/519 1.0 (0.25) 17A2 BioLegend, 100204
Anti-mouse CD19-FITC (0.5 mg/mL) 495/519 0.125 (0.25) 1D3/CD19 BioLegend, 152404
Anti-mouse NK-1.1-FITC (0.5 mg/mL) 495/519 0.25 (0.25) PK136 BioLegend, 108706
Anti-mouse CD45-PE-Cy7 (0.2 mg/mL) 488, 532, 561/767 0.125 (0.5) 30-F11 BioLegend, 103114
Anti-mouse/human CD11b-eFluor 506
(0.2 mg/mL)
419/508 0.125 (0.5) M1/70 eBioscience, 69-0112-82
Anti-mouse Ly6C-BV 785 (0.2 mg/mL) 405/785 0.25 (0.5) HK1.4 BioLegend, 128041
Anti-mouse Ly6G–APC-Cy7 (0.2 mg/mL) 650, 755/767 0.25 (0.5) 1A8 BioLegend, 127624
Anti-mouse CD11c-BV 605 (0.2 mg/mL) 405/605 0.25 (1) N418 BioLegend, 117334
Viability (Fixable Viability Dye-eFluor 455) (100 test) 355/450 0.1 (0.01) NA eBioscience, 65-0868
Note: Antibody titration is recommended for optimal performance. Avoid direct exposure of labeled samples to light. Acquire samples in a flow cytometer as soon as possible to one week at 4 °C (better within 24 h). Use FMOs, unstained control (no-cell trace control tissues) for gating. For myeloid cell phenotyping, use fluorochromes compatible with your flow cytometer. Use FluoroFinder or other software for panel design (to avoid spectral overlap of fluorescence based on the configuration of your flow cytometer).
Table 2. Cell tracer dyes used in the study; the listed dyes can be used to track cells or parasites*
Cell tracer name
(Concentration of stock)
Storage Excitation/Emission (nm) Final concentration Brand and catalog number
CellTrackerTM Orange CMTMR (10 mM) -20 °C 541/565 nm 2.5 μM Invitrogen C2927
CellTrackerTM Green CMFDA (10 mM) -20 °C 492/517 nm 5 μM Invitrogen C2925
CellTraceTM Far-Red (1 mM) -20 °C 630/661 nm 1 μM Invitrogen C34572
PKH26 Red Fluorescent Cell Linker 4 °C 551/567 nm 4 μM Sigma-Aldrich MINI26-1KT
PKH 67 Green Fluorescent Cell Linker 4 °C 490/502 nm 4 μM Sigma-Aldrich MINI67-1KT
*mCherry (5871/610 nm)
Solutions
1× phosphate-buffered saline (1× PBS) (see Recipes)
1× PBS with 0.1% BSA: FACS buffer (see Recipes)
1× PBS + 2% FBS (see Recipes)
1× 1-Step Fix/Lyse solution (see Recipes)
1× RBC lysis buffer (see Recipes)
1× BD IMagTM buffer (see Recipes)
Complete M199 medium (see Recipes)
Complete RPMI 1640 medium (see Recipes)
Collagenase D + Liberase TL Mix (see Recipes)
Collagenase D solution + DNase I (see Recipes)
Recipes
1× phosphate-buffered saline (1× PBS) from 10× PBS (4 °C storage)
Reagent Final concentration Quantity
10× PBS 1× 100 mL
Distilled H2O n/a 900 mL
Total n/a 1,000 mL
1× PBS with 0.1% BSA. FACS buffer (4 °C storage)
Reagent Final concentration Quantity
1× PBS 1× 1,000 mL
BSA 0.1% 1 g
Total n/a 1,000 mL
1× PBS + 2% FBS (4 °C storage)
Reagent Final concentration Quantity
1× PBS 1× 980 mL
100% FBS 2% 20 mL
Total n/a 1,000 mL
1× 1-Step Fix/Lyse solution (4 °C storage)
Reagent Final concentration Quantity
10× 1-step Fix/Lyse 1× 100 mL
Distilled H2O n/a 900 mL
Total n/a 1,000 mL
1× RBC lysis buffer (4 °C storage)
Reagent Final concentration Quantity
10× RBC lysis buffer 1× 10 mL
Distilled H2O n/a 90 mL
Total n/a 100 mL
1× BD IMagTM buffer (4 °C storage)
Reagent Final concentration Quantity
10× BD IMagTM buffer 1× 10 mL
Distilled H2O n/a 90 mL
Total n/a 100 mL
Complete M199 medium (4 °C storage)
Reagent Final concentration Quantity
M199 medium n/a 500 mL
FBS 10% 50 mL
Adenine 1% 6 mL
Hemin 0.1% 1.2 mL
Penicillin-streptomycin solution 1× 6 mL
Total n/a 563.2 mL
Complete RPMI 1640 medium (4 °C storage)
Reagent Final concentration Quantity
RPMI medium n/a 445 mL
FBS 10% 50 mL
Penicillin-streptomycin solution 1× 5 mL
Total n/a 500 mL
Collagenase D + Liberase TL Mix (storage at -20 °C)
Reagent Final concentration Quantity
20 mg/mL Collagenase D 2 mg/mL 2 mL
1 mg/mL Liberase TL 160 μg/mL 640 μL
RPMI complete medium n/a 17.36 mL
Total n/a 20 mL
Collagenase D Solution + DNase I (storage at -20 °C)
Reagent Final concentration Quantity
20 mg/mL Collagenase D 2 mg/mL 5 mL
2 mg/mL DNase I 20 μg/mL 500 μL
1 M HEPES pH 7.4 10 mM 500 μL
5 M NaCl 150 mM 1.5 mL
1 M KCl 5 mM 250 μL
0.5 M MgCl2 1 mM 100 μL
0.5 M CaCl2 1.8 mM 180 μL
Distilled H2O n/a 42.5 mL
Total n/a 50 mL
Laboratory supplies
Ultra-fine insulin syringe with needle (BD, catalog number: 324920)
Sterile 5 mL Polystyrene round bottom tube (Falcon, catalog number: 352054)
Sterile 70 μm cell strainer (any vendor)
Sterile 15 mL polypropylene centrifuge tubes (any vendor)
Sterile 50 mL polypropylene centrifuge tubes (any vendor)
Sterile nuclease-free filter tips (10, 200, and 1,000 μL) (any vendor)
Cell culture plates, different sizes, 6-, 12-, and 24-wells flat bottom, TC treated, sterile (any vendor)
Vented flask surface area 25 cm2 and 175 cm2 (any vendor)
Micro-dissecting scissors (Millipore, Sigma, catalog number: S3271)
Scissor-handle forceps (Millipore, Sigma, catalog number: Z168831)
Disposable plastic serological pipettes, sterile, individually wrapped, 10 mL, 25 mL capacity (any vendor)
Equipment
Flow cytometer with at least two lasers and seven parameter analysis capacity (BD LSRFortessa IITM Cell Analyzer, BD Biosciences; SE500, Stratedigm)
Eclipse TE300 Microscope (Nikon)
Software and datasets
Panel Builder, Fluorofinder (3, 2014, free)
FlowJo V10.6 software (V10.6, 2019, license required)
BD FACSDiva 9.0 (flow cytometer software)
GraphPad Prism 9.0 software (9, 2020, license required)
Procedure
Labeling of Leishmania donovani and bone marrow cells
Selection of L. donovani metacyclic promastigotes with PNA agglutinin (adapted from Sacks and Melby, 2015). This selection is adequate for L. donovani expressing mCherry fluorescent protein (Figure 1).
Figure 1. Representative fluorescent signal of Leishmania donovani infective promastigotes. A. L. donovani expressing mCherry fluorescent protein (red). B. L. donovani stained with PKH67 (green) (step A2). Parasites were selected as mentioned in step A1. Images were taken with a Nikon Eclipse TE300 Microscope at 20×.
Grow L. donovani (MHOM/SD/00/S-2D): add approximately 40 mL of parasites in logarithmic phase of culture (3–4 days) at 1 × 106 cells/mL to 400 mL of complete M199 medium. Incubate for 5–6 days (until stationary phase) at 27 °C.
Harvest the parasite by centrifugation in sterile 50 mL tubes at 700× g for 10 min.
Note: All the centrifugation steps are performed at room temperature (RT), unless the text indicates otherwise.
Discard supernatant. Caution! Be careful to not disturb the pellet.
Resuspend the pellet in 10 mL of RPMI medium and take 10 μL to count parasites. Use dilution if the number of parasites is too high and difficult to count in a Neubauer camera.
Centrifuge at 700× g for 10 min.
Discard the supernatant and resuspend in 1× PBS to a final concentration of 1 × 108 parasites/mL.
Add 50 μg/mL of PNA or 10 μL/mL.
Vortex for 30 s and incubate at RT for 30 min without agitation.
Centrifuge at 200× g for 2 min (low speed).
Take the supernatant in a fresh tube. The supernatant contains metacyclic promastigotes.
Wash parasites with 10 mL of RPMI medium. Proceed with counting viable parasites. Parasite viability is assessed by the motility of parasites.
To count viable parasites, take 10 μL of parasites from step A1k into a Neubauer chamber covered with a cover slide and count motile parasites. Parasites with any movement of the body or flagellum are counted. Then, calculate the number of parasites using the following equation:
Number of parasites per milliliter = Number of motile parasites per quadrant × dilution factor × 10,000.
Centrifuge at 700× g for 10 min.
Resuspend parasites at desired concentration (normally 1 × 106 parasites/10 μL of PBS) for intradermic (ID) inoculation or 2 × 107 parasites/mL. If the parasite is not fluorescent, proceed with staining with PKH26/67 (step A2).
Note: The parasite staining is not necessary when the L. donovani strain is expressing a fluorescent protein (mCherry). In this case, skip step A2. The mCherry L. donovani parasites are ready for intradermal infection (step A7). To perform intradermic inoculation, check Machholz et al. (2012).
Parasite staining with PKH26 (red) or PKH67 (green) for intradermal infection (modified from manufacturer’s protocol):
Place a suspension containing 2 × 107 single parasites in a 15 mL polypropylene tube and wash once using medium without serum or 1× PBS.
Centrifuge the parasites at 400× g for 5 min into a loose pellet at RT.
After centrifuging, carefully aspirate the supernatant, being careful not to remove any cells but leaving no more than 25 μL of supernatant.
Prepare a 2× parasite suspension by adding 1 mL of Diluent C to the pellet and resuspend with gentle pipetting to ensure complete dispersion. Caution! Do not vortex and do not let cells stand in Diluent C for long periods of time.
Immediately prior to staining, prepare a 2× dye solution (4 μM) in Diluent C by adding 4 μL of the PKH26 to 1 mL of Diluent C in a polypropylene centrifuge tube and mix well to disperse.
Rapidly add the 1 mL of 2× parasite suspension to 1 mL of 2× dye solution and immediately mix the sample by pipetting. Final concentrations after mixing the indicated volumes will be 1 × 107 cells/mL and 2 μM PKH26/67.
Incubate the parasite/dye suspension for 5 min with periodic mixing at 27 °C (or as best suitable for the cell you are using) in the dark. Because staining is so quick, longer times provide no advantage.
Stop the staining by adding an equal volume (2 mL) of serum or 10 mL of complete medium and incubate for 1 min to allow binding of excess dye.
Centrifuge the parasites at 400× g for 10 min at RT and carefully remove the supernatant. Resuspend cell pellet in 10 mL of complete medium, transfer to a fresh sterile conical polypropylene tube, centrifuge at 400× g for 5 min at RT, and wash the parasite pellet two more times with 10 mL of complete medium to ensure removal of unbound dye.
After the final wash, resuspend the parasite pellet in 10 mL of complete RPMI medium for assessment of cell recovery and cell viability. Optional: check fluorescence intensity in a fluorescence microscope (Figure 1). Proceed with counting viable parasites (step A1l).
Centrifuge and resuspend to a desired final concentration of viable parasites in a volume of 10 μL (1 × 106/10 μL 1× PBS).
Note: The procedure above uses 2 mL as the final staining volume, containing final concentrations of 2 μM of PKH26/67 and 1 × 107 cells/mL. Volume is adjusted to the concentration of parasites obtained.
Optional: L. donovani expressing mCherry fluorescent protein can be used (Figure 1A) instead of metacyclic promastigotes labeled with a fluorescent cell tracer. The selection of infective metacyclic promastigotes should be performed as mentioned before.
Isolation of bone marrow monocytes:
Euthanize the mice according to the approved IACUC protocol (exposure to carbon dioxide followed by cervical dislocation) and proceed with bone marrow collection. Place the mouse in a container with 70% alcohol for disinfection for 30 s. Work in a laminar flow hood.
Remove femurs and tibias from the mice using scissors and forceps. Place the tissues on the plate containing 1 mL of RPMI complete media. Keep the tissues on ice until all mice have been prepared.
Using sterile utensils, remove the muscles from the bones and place the clean bones in a Petri dish. (Estimate that one mouse femur yields approximately 1 million monocytes.)
Obtention of bone marrow cells: using sterile utensils, cut both ends of the bone with scissors as close to the joints as possible. Fill a 1 mL insulin syringe with ice-cold RPMI complete medium, insert the syringe needle into the end of the bone marrow cavity, and flush out the bone marrow into a 15 mL centrifuge tube on ice. Flush five times (5 mL) until the bones are completely white. Break up any cell clumps by pipetting up and down. Alternatively, place the bone in one sterile mortar and carefully break the bone with the pestle. Transfer the bone fragments to a sterile Petri dish containing 1 mL of Accutase solution. Incubate for 5 min at 37 °C. Add 1 mL of culture medium and flush the material out of the bones (by pipetting up and down until the cells detach from the bone). Repeat, if necessary, until the reddish bones turn white.
Obtain a single-cell suspension by filtering the solution through a 70 μm cell strainer, place it in a fresh 15 mL centrifuge tube, and centrifuge at 500× g for 5 min at RT. Discard the supernatant.
Resuspend the bone marrow cell pellet in 1–3 mL of 1× RBC lysis buffer and incubate for 5 min at RT.
Stop the lysis reaction by adding 10 mL of 1× PBS + 2% FBS. Centrifuge immediately at 500× g for 5 min at RT. Decant the supernatant and resuspend the cells in RPMI complete media (5 mL).
Proceeded with Neutrophil depletion using MojoSortTM Mouse Ly-6G Selection Kit (step A4).
Note: The cell purification kit varies in accordance with the lineage of cells to be traced.
Neutrophil depletion using MojoSortTM Mouse Ly-6G Selection kit (modified from manufacturer’s protocol). This step excludes neutrophils and enriches the sample for monocytes.
Centrifuge the bone marrow cells obtained above at 300× g for 5 min and resuspend in 4 mL of 1× MojoSort buffer. Count the cells with 0.4% trypan blue and adjust to 1 × 108 cell/mL in 1× MojoSort buffer.
Take 100 μL of cell suspension (107 cells) into a new 5 mL polypropylene tube. Add 10 μL of the Biotin anti-mouse Ly-6G antibody (clone 1A8) per 100 μL of cell suspension (107 cells/100 μL). Mix well and incubate on ice for 15 min.
Resuspend the Streptavidin nanobeads by vortex at maximum speed for 10 s and add 10 μL of Streptavidin nanobeads to the cell suspension. Mix well and incubate on ice for 15 min.
After incubation time, add 2.5 mL of MojoSort buffer.
Place the tube in the magnet for 5 min.
Optional: take a small aliquot before placing the tube in the magnet to monitor purity and yield.
Transfer the unlabeled cell fraction still in suspension (not held by the magnet) by pouring the liquid into a fresh 5 mL tube. The unlabeled fraction is the cells of interest.
Resuspend the labeled cells in MojoSort buffer and repeat steps A4e–A4g on the labeled cells at least two more times.
Pool the unlabeled fractions.
Continue with monocytes positive selection with CD11b magnetic particles DM (step A5).
Note: When working with less than 107 cells, use the indicated volumes for 107 cells. To scale up, adjust the volumes of cells and beads proportionately. For example, add 100 μL of nanobeads to separate 1 × 108 cells in 1 mL of MojoSort buffer. To maximize yield, you can disrupt the aggregates by pipetting the solution up and down. Filter the suspension using a 70 μm cell strainer.
Positive selection of monocytes with CD11b magnetic particles DM (modified from manufacturer’s protocol).
Centrifuge the unlabeled negative fraction from the previous procedure at 400× g for 5 min.
Resuspend the cell in 3 mL of 1× BD IMag buffer. Count the cells with 0.4% trypan blue and adjust to 1 × 107 cell/mL in 1× BD IMag buffer.
Centrifuge at 400× g for 5 min and carefully aspirate all the supernatant.
Vortex to maximum speed the BD IMag anti-CD11b magnetic particles DM. Add 50 μL of particles for every 1 × 107 total cells. Mix thoroughly. Refrigerate for 30 min at 4 °C.
Bring the BD IMag–particle labeling volume up to 1 × 107–8 × 107 cells/mL with 1× BD IMag buffer and immediately place the tube on the cell separation magnet. Incubate at RT for 8 min.
Keep the tube on the cell separation magnet and carefully aspirate off the supernatant with a Pasteur pipette. This supernatant contains the negative fraction. (This time, we want the CD11b positive fraction, which is attached to the magnet.)
Remove the tube from the cell separation magnet and add 1× BD IMag buffer to the same volume as in step A5e. Gently resuspend cells by pipetting briefly and return the tube to the cell separation magnet for another 4 min.
With the tube on the cell separation magnet, carefully aspirate off the supernatant and discard.
Repeat steps A5g and A5h one more time.
After the final wash, resuspend the positive fraction in 1× PBS and adjust the purified monocytes to 1 × 106 cells/500 μL for staining with the cell tracer (step A6).
Monocyte staining using cell tracers:
Label the monocytes with any of the cell tracers in Recipe 9. However, make sure that the excitation/emission wavelength is different from the parasites. For this, dissolve the lyophilized CMTMR (orange) or CMFDA (green) CellTrackerTM in high-quality DMSO to get a 10 mM stock or dilute the Far-Red CellTraceTM to obtain a 1 mM stock with high-quality DMSO.
Dilute the stock solution to a 2× working concentration of 2.5 μM CMTMR (orange), 5 μM CMFDA (green), or 1 μM Far-Red CellTraceTM (Figure 2) in 500 μL of serum-free medium or 1× PBS. For this, add either 0.5 μL of CMTMR, 1 μL of CMFDA, or 2 μL of Far-Red to 500 μL of serum-free medium or 1× PBS. Vortex.
Note: Cell tracer dyes and PKH linkers are equally useful to trace lymphoid cells and parasites.
Figure 2. Representative fluorescent signal of monocytes stained with cell tracers. A. Mouse bone marrow monocytes stained with 5 μM CMFDA (green). B. Monocytes stained with 1 μM Far-Red cell trace (magenta). Monocytes were isolated and labeled as in steps A3–A6. Images were taken with a Nikon Eclipse TE300 Microscope at 40×.
Warm the 2× cell tracker working solution to 37 °C and add 500 μL to the cells obtained in step A5 (1 × 106 cells in 500 μL of 1× PBS) for a final volume of 1 mL. Mix the tube by inversion.
Incubate for 20 min at 37 °C with 5% CO2 protected from light.
Add five times the original staining volume of complete culture medium to the cells and incubate for 5 min. This step removes any free dye remaining in the solution. Count the cells.
Centrifuge the cells at 300× g for 5 min. Carefully remove all the supernatant with a pipette.
Add RPMI complete culture medium to adjust the cells to 2.5 × 106–4 × 106 per 10–20 μL.
Optional: Check the fluorescent signal at the microscope (Figure 2).
Intradermal transfer of cell tracer–labeled monocytes and intradermal infection using fluorescent L. donovani (in vivo).
Cell tracer–labeled monocytes (step A6) and fluorescent parasites (mCherry) or pre-labeled with a cell tracer (step A2) are inoculated by ID injection.
Place the parasite (1 × 106 parasite/10 μL) into an ultra-fine insulin syringe (protected from light).
In an ABSL2 room, put the mice under anesthesia and transfer the cell tracer–labeled monocytes (Figure 2) by ID injection (2.5 × 106–4 × 106 per 10 μL).
Immediately inoculate intradermally the fluorescent parasite (1 × 106/10 μL) in each foot (dorsal hind paw) or as needed.
Twenty-four to forty-eight hours after transfer of cell tracer labeled cells and infection, proceed with popliteal lymph node and spleen isolation for flow cytometry acquisition and analysis (follow sections B and C).
Collection of in vivo samples and tissue processing
Isolation of cells from skin
Euthanize the mice as mentioned before and proceed with the skin tissue collection. For this, hold the skin with forceps and cut the skin of the infected hind paw with a scalpel.
Note: Collect the draining lymph nodes and spleen and treat them as in section C.
Place the inoculation site tissue (skin of hind paw) into a 24-well plate in 1 mL of complete RPMI medium on ice (4 °C).
Transfer skin tissue to 500 μL of 2 mg/mL collagenase D + 160 μg/mL of Liberase (see Recipes). Reserve the complete medium from the step above (B1b) at 4 °C to use in step B1e.
Cut the tissue in small parts with a blade or scissors (< 2 mm), incubate at 37 °C for 30 min, and mix at 200 rpm if an incubator with shaker is available. Alternatively, mix the tube by inversion every 5 min during the incubation time.
Add the 1 mL complete medium saved from above (B1c) to stop the enzymatic digestion (B1d).
Collect the cell suspension together with any tissue in the suspension and press the tissue through a 70 μm cell strainer with a 1 mL syringe plunger. Use another well or a small Petri dish to collect the cell suspension.
Add 1 mL of culture medium to the cell strainer to help pass through the cells.
Transfer the cell homogenate to a 5 mL Falcon tube.
Pellet the cells by centrifuging at 300× g for 7 min at 4 °C.
Remove the supernatant and add 1 mL of FACS buffer (see Recipes).
Count cells with 0.4% trypan blue (use 1:10 dilution).
Centrifuge cells at 300× g for 7 min at 4 °C, remove the supernatant, and adjust cell count to 1 × 106/50 μL per tube by adding FACS buffer.
Proceed with cell surface staining for flow cytometry (see below).
Note: Use uninfected (stained samples without parasites) as control to establish flow cytometry thresholds for infected cells [dot-plot of the forward scatter (FSC) of cell population of interest vs. fluorescence channel of parasites].
Isolation of cells from spleen and draining lymph node (dLN)
Disinfect the euthanized mice with 70% alcohol.
Proceed to make an incision on the mouse’s left side, high under the rib cage, where the spleen is located. Collect the spleen and place it into a Petri dish with RPMI complete medium at 4 °C.
Make an incision on the right and left hind leg using scissors. Pushing the knee up to expose the popliteal fossa and using forceps, carefully remove the popliteal lymph node draining the infection site (Figure 3) and place it into a Petri dish with RPMI complete medium at 4 °C. Use a stereoscope as an alternative to facilitate visualization of lymph node.
Figure 3. Mouse popliteal lymph node (pLN) localization. A. Popliteal lymph node drawing. B. Photo of the pLN (dot circle) using a stereoscope for localization and isolation. C. Comparison between an uninfected pLN (left) and Leishmania donovani–infected pLN (right) view under a stereoscope.
Transfer the tissues into a Petri dish with Collagenase D solution + DNase I (see Recipes).
Inject the mouse spleen with 1 mL of Collagenase D solution + DNase I per spleen using a 1 mL syringe, then cut the tissue into smaller (1 mm) pieces using sharp scissors. Skip this step for the popliteal lymph node.
Incubate for 30 min at 37 °C with 5% CO2.
Inactivate the reaction by adding 1 mL of RPMI complete media.
Transfer the suspension of tissue pieces into a 70 μm cell strainer placed on a Petri dish. Add 1 mL of 1× PBS to the strainer and gently press the tissue through the strainer with a 1 mL syringe plunger.
Transfer the cell suspension to a 5 mL polypropylene tube.
Centrifuge at 500× g for 5 min. Discard the supernatant. Skip step B2k–m for lymph nodes.
To eliminate red blood cells from spleen tissues, resuspend the pellet in 1 mL of 1× RBC lysis buffer for 5 min.
Stop the lysis reaction by adding 3 mL of 1× PBS + 2% FBS.
Centrifuge immediately at 300× g for 5 min at RT. Decant the supernatant.
Resuspend the cells in 300 μL to 1 mL of PBS to adjust the cell concentration to 1× 106 cells/50 μL.
Proceed with cell surface staining for myeloid cell phenotyping flow cytometry.
Finally, proceed with flow cytometry acquisition and analysis.
Note: If you are working with groups of mice, 24-well plates can be used to preserve the spleen or other organs at 4 °C. However, it is important to work fast to avoid cell death.
Phenotyping of harvested cells
Cell surface staining for myeloid cell phenotyping flow cytometry
Samples are adjusted at 1 × 106/50 μL of 1× PBS (section B). Distribute 50 μL of cell suspension to a 5 mL polypropylene tube.
Add 50 μL of 1× PBS with FC block antibody (dilution 1:100) and Fixable Viability Dye eFluor 455UV (dilution 1:1,000) per tube. Add only PBS to one non-stained control tube.
Note: The staining for viability is done in 1× PBS as per manufacturer’s instructions; use FACS buffer otherwise.
Incubate for 20 min at 4 °C, protected from light.
Wash cells using 2 mL of FACS buffer and centrifuge at 300× g for 5 min at 4 °C. Decant supernatant.
Resuspend samples in 50 μL of FACS buffer with antibody staining cocktail per tube as below. Reserve tubes: unstained and FMO controls as needed for gating.
Prepare the antibody staining cocktail to identify the cell population of interest. In this case, we are using a panel to identify the main myeloid cell populations: monocytes, neutrophils, dendritic cells, and monocyte-derived dendritic cells (see suggested flow cytometry panel in Table 1). The number of markers depends on the flow cytometer configuration and the antibodies available. Incubate for 30 min on ice in the dark.
After incubation, add 3 mL of FACS buffer and centrifuge at 400× g for 5 min at 4 °C.
Discard the supernatant and resuspend cell pellet in 300 μL of 1× Fix/Lyse.
Incubate for 15 min at RT in the dark.
Add 3 mL of FACS buffer and centrifuge at 400× g for 5 min at 4 °C.
Wash the pellet with 3 mL of FACS buffer.
Resuspend the cell pellet in 300 μL of FACS buffer.
Store fixed cells in the dark at 4 °C for flow cytometry acquisition and analysis.
Note: Make sure to include fluorescence minus one (FMO) controls for the flow cytometry analysis.
Flow cytometry acquisition and analysis
For BD LSRFortessa IITM Cell Analyzer, turn on the machine and check fluid levels. Set the instrument settings (assistance for sample acquisition and analysis may be necessary if the user is not familiar with flow cytometry). Proceed with acquisition of the single-color compensation controls, FMO controls, and negative controls.
Collect between 100,000 and 1,000,000 events to capture enough infected cells for analysis.
Note: To generate a compensation matrix, it is preferable to use compensation beads (UltraComp eBeads, Thermo Fisher Scientific). For the compensation of cell tracers, is necessary to use cells reserved from the labeling with the fluorescent cell tracer or parasites expressing a fluorescent protein, if that is the case.
Data analysis
Flow cytometry analysis
For the flow cytometry analysis, we recommend using FlowJo. Basic analysis features may be available in each instrument.
To analyze data in FlowJo, follow the gating schemes in Figure 4 (adapted from Rose et al., 2012; Osorio et al., 2023). We performed different gating strategies depending on the cell population to be analyzed.
Figure 4. Representative gating strategy to detect fluorescent Leishmania in infected myeloid cells on lymphoid tissues. A. Representative gating strategy to evaluate infected cells in draining lymph node (dLN) and spleen (evaluated at 6 h and 12 h post-infection, respectively). B. Gating strategy to determine cells stained with cell tracer infected with Leishmania. Representative dot-plot skin (left) or lymph node (right) of monocytes labeled with Far-Red cell tracer positive for fluorescent Leishmania parasites (mCherry + or PKH26 cell tracer +) after 48 h of transference. Dot-plot from Ly6G-CD11b+ cells (gates A6). The threshold is established above the fluorescent signal of FMO controls and uninfected tissue controls. See detailed gating strategy in Data analysis.
Gating strategy to identify cell tracer–labeled Leishmania in neutrophils, monocytes, conventional dendritic cells (cDCs), and monocyte-derived dendritic cells (MoDCs) (dLN and spleen): first, gate events on FSC-A and SSC-A (A1), and then gate on FSC-A vs. FSC-H to exclude doublets (A2). Cells are sub-gated to exclude dead cells in 455 UV (A3) and then T cells (CD3+), B cells (CD19+), and NK cells (NK-1.1+) in the FITC dump channel for T, B, and NK cells (A4). Leukocytes CD45+ (PE-Cy7) are sub-gated (A5). Then, CD11b+ (eFluor 506) population are sub-gated using the expression of Ly6G+ (APC-Cy7) to determine monocytes (CD11b+ Ly6G-) (A6) or neutrophils (CD11b+Ly6G+). cDCs and MoDCs are sub-gated using the expression of CD11c+ and CD11b+ to determine cDCs (CD11c+ CD11b-) (A7) or MoDCs (CD11c+ CD11b+). Monocytes are determined as CD11c- CD11b+ (A9). Monocytes could further be classified according to Ly6C expression as resident monocytes (Ly6Clow/-) or inflammatory monocytes (Ly6Chi+/int) (A10).
All populations are sub-gated in a dot-plot with FSC-A and the fluorescence of Leishmania (mCherry or labeled with a cell tracer such as PKH26) to identify infected cells (Leishmania) (A8 and 11).
Similarly, isolated, labeled, and transferred monocytes can be identified using the gating strategy explained above (Figure 4B). Here, after sub-gating monocytes as CD11b+ Ly6G- (A6 or B6), transferred monocytes labeled with Far-Red cell tracer are gated with the fluorescent Leishmania. The double staining indicates the transferred monocytes are infected with Leishmania parasites.
Validation of protocol
Parts of these procedures were standardized and validated previously (Ibrahim et al., 2013 and Osorio et al., 2023). For flow cytometry analysis, the threshold was established as the fluorescent signal above the FMO controls and uninfected tissue controls. Data were analyzed with GraphPad Prism 9.0 software (GraphPad, LLC).
General notes and troubleshooting
Troubleshooting (Table 3)
Table 3. Troubleshooting
Problem Possible reason Solution
Cell clumping after using collagenase D or other proteases Extended time using enzymatic degradation
• Reduce the time of cell exposure to collagenase D solution
• Be sure to add the DNase I into collagenase D solution to remove free DNA from the destroyed tissue
Incomplete disaggregation of the cells/cell suspension is not achieved property Short time using enzymatic degradation or ineffective filtration When cell agglomeration occurs, passing the cell suspension through a 70 or 50 μm strainer can help. This will also avoid clogs when using the Flow cytometer
Compensation problem
Inadequate fluorochrome panel design
Lack of controls
• Design your fluorochrome panel using specialized software (Fluorofinder). Ask a flow cytometrist for help
• Reduce the number of fluorochromes to adapt the protocol to your instrument configuration. Reconfigure the panel if necessary
• Use no-cell trace negative tissues and FMO controls for threshold and analysis.
Acknowledgments
This work was supported by the U.S. National Institutes of Health (NIH/NISID) grant numbers AI107419 and AI13012 to P.C.M. We thank the staff at the Animal Resources Center at the University of Texas Medical Branch for excellent care provided to the experimental animals. This protocol was adapted from the less-detailed original (Ibrahim et al., 2013 and Osorio et al., 2023).
Competing interests
The authors have declared that no competing interests exist.
Ethical considerations
The protocol was approved by the Institutional Animal Care and Use Committee of the University of Texas Medical Branch, Galveston, Texas (protocol number 1306027).
References
Al-Zaben, N., Medyukhina, A., Dietrich, S., Marolda, A., Hünniger, K., Kurzai, O. and Figge, M. T. (2019). Automated tracking of label-free cells with enhanced recognition of whole tracks. Sci. Rep. 9(1): e1038/s41598-019-39725-x.
Halabi, E. A., Arasa, J., Püntener, S., Collado-Diaz, V., Halin, C. and Rivera-Fuentes, P. (2020). Dual-activatable cell tracker for controlled and prolonged single-cell labeling. ACS Chem. Biol. 15(6): 1613–1620.
Hampton, H. R. and Chtanova, T. (2019). Lymphatic Migration of Immune Cells. Front. Immunol. 10: e01168.
Ibrahim, M. K., Barnes, J. L., Anstead, G. M., Jimenez, F., Travi, B. L., Peniche, A. G., Osorio, E. Y., Ahuja, S. S. and Melby, P. C. (2013). The Malnutrition-Related Increase in Early Visceralization of Leishmania donovani Is Associated with a Reduced Number of Lymph Node Phagocytes and Altered Conduit System Flow. PLoS Negl.Trop. Dis. 7(8): e2329.
Machholz, E., Mulder, G., Ruiz, C., Corning, B. F. and Pritchett-Corning, K. R. (2012). Manual Restraint and Common Compound Administration Routes in Mice and Rats. J. Vis. Exp.: e3791/2771.
Maiorino, L., Shevik, M., Adrover, J. M., Han, X., Georgas, E., Wilkinson, J. E., Seidner, H., Foerschner, L., Tuveson, D. A., Qin, Y. X., et al. (2022). Longitudinal Intravital Imaging Through Clear Silicone Windows. J. Vis. Exp.: e3791/62757–v.
Osorio, E. Y., Uscanga-Palomeque, A., Patterson, G. T., Cordova, E., Travi, B. L., Soong, L. and Melby, P. C. (2023). Malnutrition-related parasite dissemination from the skin in visceral leishmaniasis is driven by PGE2-mediated amplification of CCR7-related trafficking of infected inflammatory monocytes. PLoS Negl.Trop. Dis. 17(1): e0011040.
Rose, S., Misharin, A. and Perlman, H. (2012). A novel Ly6C/Ly6G-based strategy to analyze the mouse splenic myeloid compartment. Cytometry Part A 81(4): 343–350.
Sacks, D. L. and Melby, P. C. (2015). Animal Models for the Analysis of Immune Responses to Leishmaniasis. Curr. Protoc. Immunol. 108(1): eim1902s108.
Tomura, M., Ikebuchi, R., Moriya, T. and Kusumoto, Y. (2021). Tracking the fate and migration of cells in live animals with cell-cycle indicators and photoconvertible proteins. J. Neurosci. Methods 355: 109127.
You, S., Tu, H., Chaney, E. J., Sun, Y., Zhao, Y., Bower, A. J., Liu, Y. Z., Marjanovic, M., Sinha, S., Pu, Y., et al. (2018). Intravital imaging by simultaneous label-free autofluorescence-multiharmonic microscopy. Nat. Commun. 9(1): 2125.
Supplementary information
The following supporting information can be downloaded here:
Figure S1. Presence of the Green Fluorescent Protein(GFP) in the photoactivable mice(PA-GFP)
Video S1. Single-cell photoactivation
Table S1. Conditions tested in the photoconvertible model PA-GFP for identifying migratory cells photoactivated in skin. Most of the conditions showed no significant photoconversion
Article Information
Copyright
© 2023 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
Immunology > Immune cell staining > Flow cytometry
Cell Biology > Cell movement > Cell migration
Immunology > Immune cell isolation > Myeloid cell
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4,818 | https://bio-protocol.org/en/bpdetail?id=4818&type=0 | # Bio-Protocol Content
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Differentiation of Bone Marrow Monocytes into Alveolar Macrophages-like Cells through Co-culture with Lung Epithelial Cells and Group 2 Innate Lymphoid Cells
PL Pauline Loos
TM Thomas Marichal
BM Bénédicte Machiels *
LG Laurent Gillet *
(*contributed equally to this work)
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4818 Views: 834
Reviewed by: Rajesh RanjanDebashis DuttaThirupugal Govindarajan
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Original Research Article:
The authors used this protocol in Science Immunology Feb 2023
Abstract
During life, the embryonic alveolar macrophage (AM) population undergoes successive waves of depletion and replenishment in response to infectious and inflammatory episodes. While resident AMs are traditionally described as from embryonic origin, their ontogeny following inflammation or infection is much more complex. Indeed, it appears that the contribution of monocytes (MOs) to the AM pool is variable and depends on the type of inflammation, its severity, and the signals released in the microenvironment of the pulmonary niche (peripheral imprinting) and/or in the bone marrow (central imprinting). Deciphering the cellular and molecular mechanisms regulating the differentiation of MOs into AMs remains an area of intense investigation, as this could potentially explain part of the inter-individual susceptibility to respiratory immunopathologies. Here, we detail a relevant ex vivo co-culture model to investigate how lung epithelial cells (ECs) and group 2 lung innate lymphoid cells (ILC2s) contribute to the differentiation of recruited MOs into AMs. Interestingly, the presence of lung ILC2s and ECs provides the necessary niche signals to ensure the differentiation of bone marrow MOs into AMs, thus establishing an accessible model to study the underlying mechanisms following different infection or inflammation processes.
Key features
• Ex vivo co-culture model of the alveolar niche.
• Deciphering the particular niche signals underlying the differentiation of MO into AMs and their functional polarization.
Graphical overview
This protocol described the isolation of bone marrow monocytes (MOs), lung epithelial cells (ECs), and lung group 2 lung innate lymphoid cells (ILC2s) and the ex vivo co-culture of these cells to drive the differentiation of bone marrow MOs into alveolar macrophages (AMs).
This co-culture experiment is composed of three steps (Graphical overview):
1. Identification and FACS-sorting of ECs and MOs isolated from the lung and the bone marrow of naive mice, respectively.
2. Culture of these ECs and bone marrow MOs for three days.
3. Addition of ILC2s isolated from the lung of naïve mice or mice subjected to a treatment/infection of interest.
Keywords: Group 2 innate lymphoid cells Monocytes Alveolar macrophages Alveolar Niche Co-culture Monocytes differentiation and polarization
Background
Innate lymphoid cells (ILCs) were firstly described in 2010 and have since been described as important regulators of both inflammation and homeostasis throughout the body (Moro et al., 2010; Ebbo et al., 2017). They affect the function of their neighboring cells through the cytokines they produce and also via direct cell–cell interactions (Spits and Di Santo, 2011; Oliphant et al., 2014). Their role in modulating alveolar macrophage (AMs) polarization has been described in early life through the IL-33/ILC2/IL-13 activation cascade, which confers an M2 phenotype to AMs (Saluzzo et al., 2017). In the context of a persistent viral infection, we recently highlighted the essential role of ILC2s as lung niche cells that imprint the tissue-specific identity of monocytes (MOs)-derived AMs and shape their function (Loos et al., 2023).
The protocol presented here details the optimal settings to recapitulate an alveolar niche ex vivo allowing the differentiation of MOs into AMs. In particular, this protocol describes the experimental conditions for co-culture of epithelial cells (ECs) and bone marrow MOs, with group 2 lung innate lymphoid cells (ILC2s) isolated from mouse lung. In the related publication, we showed that bone marrow MOs differentiate in vitro into AM-like cells in the presence of ILC2s and ECs, as evidenced by the expression of genes related to macrophage differentiation and activation (Csf1, PPARg, and Il4ra) (Loos et al., 2023). In addition, we also showed the long-term effect of lung gammaherpesvirus infection on ILC2s and the subsequent consequences on ILC2-mediated differentiation of MOs into AMs. In particular, we highlighted the key role of granulocyte-macrophage colony-stimulating factor (GM-CSF) produced by ILC2s in the differentiation of MO-AMs. The main advantage of this protocol is that it reduces the cell types involved in the differentiation of MO-AMs to a number of selected candidates that we can study ex vivo. This approach therefore allows to test the influence of either environmental conditions (such as viruses or pollutants) or defined mediators in a very controlled environment with every other condition being equal. Subsequently, the differentiation of MOs into AM-like cells can be analyzed classically by multiparametric flow cytometry or by transcriptomic analyses such as bulk or single-cell RNA sequencing. Besides, modifications in intracellular metabolism driving epigenetic reprogramming could be investigated via metabolic (Cedex BioAnalyzer) and epigenetic analysis (ATAC-seq, CHIP-seq). Subsequently, those differences could be confirmed using specific neutralizing antibodies or metabolic (OXPHOS inhibitor, such as oligomycin, or glycolysis manipulator, such as PKM2-pyruvate) (Surace et al., 2021) or epigenetic (BET bromodomain inhibitor IBET151) inhibitors (Kerscher et al., 2019). Altogether, this protocol presents a method to study the different aspects of the biology of MO-derived AMs under controlled and reproducible experimental conditions.
Materials and reagents
Biological materials
Female C57BL/6 mice (8 weeks old) (Charles River, C57BL/6NCrl, catalog number: 000664)
Reagents
Dulbecco’s modified Eagle medium (DMEM, high glucose, GlutaMAX Supplement, pyruvate) (Thermo Fisher Scientific, Gibco, catalog number: 31966021)
RPMI 1640 medium (Thermo Fisher, catalog number: 11875093)
Minimum essential medium (MEM) (Sigma, catalog number: M0446-500ML)
Bovine serum albumin (BSA) (Sigma, catalog number: A8412)
Fetal bovine serum (FBS) (Gibco, catalog number: 10082147)
L-Glutamine (Corning, catalog number: 25005CI)
Sodium pyruvate (Corning, catalog number: 25-000-CIR)
Penicillin-streptomycin (Gibco, catalog number: 15070063)
β-mercaptoethanol (Sigma-Aldrich, catalog number: M3148)
Phosphate-buffered saline, pH 7.4 (Gibco, catalog number: 10010-023)
EDTA (Corning, catalog number: 46-034-CI)
Dispase (Sigma-Aldrich, catalog number: 42613-33-2)
Liberase research grade (Sigma-Aldrich, catalog number: 5401119001)
DNase (Sigma-Aldrich, catalog number: 11284932001)
GentleMACS C tube (Miltenyi, catalog number: 130-093-237)
Red blood cells (RBC) lysis buffer (Thermo Fisher Scientific, catalog number: 00433357)
MojoSort mouse anti-CD45 nanobeads (BioLegend, catalog number: 480027)
MojoSort mouse anti-APC nanobeads (BioLegend, catalog number: 480072)
Recombinant murine GM-CSF (Peprotech, catalog number: 315-03)
Recombinant murine IL-2 (Peprotech, catalog number: 212-12)
Zombie Aqua Fixable Viability kit (BioLegend, catalog number: 423101)
Sytox Blue Dead Cell stain (Thermo Fisher, catalog number: s34857)
Anti-mouse CD16/32 Fc block clone 93 (BioLegend, catalog number: 101301)
Paraformaldehyde (PFA) (Santa Cruz Biotechnology, catalog number: sc-281692)
Saponin (Sigma-Aldrich, catalog number: 47036)
Sodium azide (NaN3) (Sigma-Aldrich, catalog number: S2002)
TRIzol reagent (Thermo Fisher, catalog number: 15596026)
Flow cytometry antibodies
Antibodies for bone marrow MO enrichment (bead-based negative selection) and sorting:
Negative selection
i. Anti-mouse B220/CD45R APC (clone RA3-6B2) (BioLegend, catalog number: 103212)
ii. Anti-mouse Ly6G APC (clone 1A8) (BioLegend, catalog number: 127613)
Flow cytometry mastermix staining
i. Anti-mouse CD11b BV711 (clone M1/70) (BD Biosciences, catalog number: 563168)
ii. Anti-mouse Ly6G APC-Cy7 (clone 1A8) (BD Biosciences catalog number: 560600)
iii. Anti-mouse Ly6C BV785 (clone HK1.4) (BioLegend, catalog number: 128041)
iv. Anti-mouse I-A/I-E PE/Cyanine7 (clone M5/114.15.2) (BioLegend, catalog number: 107630)
v. Anti-mouse CD3e FITC (clone 145-2C11) (BioLegend, catalog number: 100306)
Antibodies for lung EC enrichment (CD45 bead-based negative selection) and sorting:
Flow cytometry mastermix staining
i. Anti-mouse CD45 PE/Cyanine7 (clone 30-F11) (BioLegend, catalog number: 103114)
ii. Anti-mouse EpCAM FITC (clone G8.8) (BioLegend, catalog number: 118207)
iii. Anti-mouse CD31 PE (clone 390) (BioLegend, catalog number: 102407)
Antibodies for lung ILC2 enrichment (bead-based negative selection) and sorting:
Negative selection
i. Anti-mouse B220/CD45R APC (clone RA3-6B2) (BioLegend, catalog number: 103212)
ii. Anti-mouse CD11c APC (clone N418) (BioLegend, catalog number: 117310)
iii. Anti-mouse CD3e APC (clone 145-2C11) (BioLegend, catalog number: 100312)
iv. Anti-mouse CD4 APC (clone RM 4-5) (BioLegend, catalog number: 100515)
v. Anti-mouse CD49b APC (clone DX5) (BioLegend, catalog number: 108910)
vi. Anti-mouse CD5 APC (clone 53-7.3) (BioLegend, catalog number: 100626)
vii. Anti-mouse CD8a APC (clone 53-6.7) (BioLegend, catalog number: 100711)
viii. Anti-mouse F4/80 APC (clone BM8) (BioLegend, catalog number: 123116)
ix. Anti-mouse FcϵRIα APC (clone MAR-1) (BioLegend, catalog number: 134316)
x. Anti-mouse Gr-1 APC (clone RB6-8C5) (BioLegend, catalog number: 108412)
xi. Anti-mouse Siglec-F APC (clone E50-2440) (BioLegend, catalog number: 155507)
Flow cytometry mastermix staining
i. Anti-mouse CD45 PE/Cyanine7 (clone 30-F11) (BioLegend, catalog number: 103114)
ii. Anti-mouse B220/CD45R APC (clone RA3-6B2) (BioLegend, catalog number: 103212)
iii. Anti-mouse CD11c APC (clone N418) (BioLegend, catalog number: 117310)
iv. Anti-mouse CD3e APC (clone 145-2C11) (BioLegend, catalog number: 100312)
v. Anti-mouse CD4 APC (clone RM 4-5) (BioLegend, catalog number: 100515
vi. Anti-mouse CD49b APC (clone DX5) (BioLegend, catalog number: 108910
vii. Anti-mouse CD5 APC (clone 53-7.3) (BioLegend, catalog number: 100626
viii. Anti-mouse CD8a APC (clone 53-6.7) (BioLegend, catalog number: 100711
ix. Anti-mouse F4/80 APC (clone BM8) (BioLegend, catalog number: 123116)
x. Anti-mouse FcϵRIα APC (clone MAR-1) (BioLegend, catalog number: 134316)
xi. Anti-mouse Gr-1 APC (clone RB6-8C5) (BioLegend, catalog number: 108412)
xii. Anti-mouse Siglec-F APC (clone E50-2440) (BioLegend, catalog number: 155507)
xiii. Anti-mouse CD25 Alexa Fluor 700 (clone PC61) (BioLegend, catalog number: 102024)
xiv. Anti-mouse ST2 PE (clone DIH9) (BioLegend, catalog number: 145304)
xv. Anti-mouse CD90.2 BV711 (clone 53-2.1) (BD Biosciences, catalog number: 740647
Solutions
Wash buffer (see Recipes)
MACS buffer (see Recipes)
FACS buffer (see Recipes)
Complete culture medium (see Recipes)
Lung digestion solution (see Recipes)
MO and EC culture medium (see Recipes)
ILC2 culture medium (see Recipes)
Permeabilization buffer (see Recipes)
Intracellular staining buffer (see Recipes)
Recipes
Wash buffer
PBS-EDTA 2 mM with 10% FBS
MACS buffer
PBS containing 0.5% BSA and 2 mM EDTA
Keep buffer cold (2–8 °C)
FACS buffer
PBS containing 0.5% BSA, 2 mM EDTA, 2 mM sodium azide (NaN3)
Complete culture medium
RPMI medium supplemented with 10% FBS, 1% MEM, 100 U/mL penicillin, 100 U/mL streptomycin, 2 mM L-glutamine, 50 μM of β-mercaptoethanol
Lung digestion solution
Freshly prepare 1.5 mL/sample of RPMI containing 50 μg/mL Liberase and 100 μg/mL DNase I
MO and EC culture medium
RPMI medium supplemented with 10% FBS, 1% MEM, 100 U/mL penicillin, 100 U/mL streptomycin, 2 mM L-glutamine, 50 μM of β-mercaptoethanol, and 10 ng/mL recombinant murine GM-CSF
ILC2 culture medium
RPMI-1640 medium supplemented with 10% FBS, 1% MEM, 100 U/mL penicillin, 100 U/mL streptomycin, 2 mM L-glutamine, 50 μM of β-mercaptoethanol, and 10 ng/mL recombinant mouse IL-2
Permeabilization buffer
PBS containing 0.1% of saponin
Keep buffer cold and sterile after filtration
Intracellular staining buffer
FACS buffer (see Recipe 3) containing 0.1% of saponin
Laboratory supplies
15 and 50 mL conical centrifuge tubes (Corning, catalog numbers: 352096 and 352070)
Cell culture 60 × 15 mm Petri dishes (Thermo Fisher Scientific, catalog number: 150288)
24-well cell culture plates (Falcon, catalog number: 353047)
96-well round (U) bottom plate (Thermo Scientific, catalog number: 163320)
10 mL syringes (Terumo, catalog number: SS+10ES1)
25 G needles (Terumo, catalog number: AN*2516R1)
Falcon® 70 μm cell strainer (Corning, catalog number: 352350)
LD columns (Miltenyi, catalog number: 130-042-901)
Equipment
Swing-bucket centrifuge (Eppendorf, model: 5810R)
Tabletop centrifuge (Eppendorf, model: 5427R)
Inverted light microscope (NIKON, Eclipse TS100)
Fluorescence-activated cell sorter (BD Biosciences, model: Aria IIIu)
Flow cytometer (BD LSR Fortessa X-20)
Laminar flow hood (FASTER SafeFast Classic)
Scalpel (Swann-Morton 210 mm No.3 ref 0933)
Forceps (Bochem Stainless steel 18/10 ref 1141)
GentleMACS dissociator (Miltenyi)
QuadroMACS Separators (Miltenyi)
Hemocytometer (Marienfeld, 0640710)
Software and datasets
FlowJo v10 (FlowJo LLC)
FACSDiVa software
Procedure
In order to get enough lung ECs and ILC2s to perform the co-culture settings, purify lung ECs and ILC2s from five mice. This will allow you to obtain, after each enrichment, 12–15 wells of 5 × 103 lung ECs and 2.5 × 103 ILC2s.
Caution: The entire procedure must be performed under a laminar flow hood to maintain the culture sterile.
Bone marrow MOs isolation
Preparing bone marrow cell suspension
Euthanize the mice and spray the skin with 75% ethanol.
Remove the skin from the lower part of the body.
Remove tissues from legs with scissors.
Separate the femur from the femoral capsule. Be careful not to break the bones; the best is to dislocate the bones rather than cutting them. Place femur and tibia of one leg into a 50 mL tube filled with 20 mL of DMEM on ice.
Take the bones and cut off both ends with a scalpel in a Petri dish.
Using a 10 mL syringe with a 25 G needle, flush the bone marrow cells into a 50 mL tube with wash buffer (see Recipes).
Pipette to obtain a homogeneous cell suspension and filter it through a 70 μm cell strainer into a new 50 mL tube.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant.
Resuspend in 1 mL of RBC lysis buffer. Incubate for 5 min at room temperature (RT) and add 9 mL of wash buffer. Count cells using the hemocytometer.
Note: You should expect approximately 40 × 106–60 × 106 cells with ~94%–98% viability for bone marrow cell collection from both femurs and tibias of an uninfected 8-week-old female C57BL/6 mouse.
Negative selection
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant.
Resuspend in MACS buffer (108 cells/mL) (see Recipes) containing anti-Ly6G and B220 antibodies coupled with APC fluorochrome (see Flow cytometry antibodies 1.a.i) and incubate for 20 min at 4 °C in the dark.
Note: The B220 and Ly6G antibodies target B cells and neutrophils, respectively, that will be further depleted, thereby facilitating FACS sorting.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 1 mL of ice-cold MACS buffer containing purified anti-CD16/32 antibody (diluted 1:500) to block Fc receptors.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant.
Resuspend the cells in MACS buffer (107 cells/100 μL) with the adapted volume of MojoSort APC Nanobeads (10 μL/107 cells) (according to MojoSort Mouse anti-APC Nanobeads protocol). Incubate on ice for 15 min.
Wash the cells by adding MACS buffer up to 4 mL. Centrifuge the cells at 250× g for 5 min at 4 °C. Discard the supernatant.
Resuspend in MACS buffer (5 × 107 cells/500 μL).
Filter cells through 70 μm cell strainer.
Proceed with magnetic negative selection: apply cell suspension onto the prepared LD column (according to Miltenyi LD column protocol). Collect flowthrough containing unlabeled cells after washes with 2 × 1 mL of MACS buffer. The unlabeled cells contain MOs.
MO staining for FACS sorting
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet in FACS buffer (107 cells/mL) (see Recipes) containing antibodies (anti-CD19, CD3, Ly6G, B220, CD11b, Ly6C) for 20 min at 4 °C in the dark. Centrifuge the cells at 250× g for 5 min at 4 °C. Discard the supernatant. Resuspend the pellet in 5 mL of PBS and repeat centrifugation.
Discard supernatant and perform viability staining by adding SYTOX Blue to FACS buffer (1 μL/10 mL/5 × 107 cells). Incubate for 5 min at RT in the dark.
Perform FACS sorting using FACS Aria IIIu. Bone marrow MOs are gated as Ly6G-MHCII-CD3-CD11b+Ly6C+ living cells (Figure 1).
Figure 1. Gating strategy for FACS sorting of bone marrow monocytes (MOs), described as SSC-AlowLy6C+CD11b+ living cells after exclusion of neutrophils (Ly6G+CD11b+), B cells (MHCII+CD11b-), and T cells (CD3+CD11b-) after cell enrichment through MACS-negative selection
Count the cells using the hemocytometer.
Centrifuge at 250× g for 5 min at 4 °C. Discard supernatant and resuspend the cells at 105 cells/100 μL of complete culture medium (see Recipes).
Lung ECs isolation
Preparing lung cell suspension
Euthanize the mice and spray the skin with 75% ethanol.
Catheterize the trachea and add 1 mL of PBS with 10 U/mL of Dispase.
Perfuse through the right ventricle with 5 mL of ice-cold PBS. Dissociate lobes of lung and place them into a 24-well plate with 2 mL of DMEM medium with 10 U/mL of Dispase for 10 min at RT.
Transfer the lung lobes from one mouse into an individual GentleMACS tube containing the lung digestion solution (see Recipes) and process with a gentleMACS dissociator (lung program 01); finally, incubate for 30 min at 37 °C. After digestion, run the lung program 02 with the gentleMACS dissociator.
Filter the cell suspension through a 70 μm cell strainer into a 50 mL tube and top up to 25 mL with wash buffer.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet in 1 mL of RBC lysis buffer. Incubate for 5 min at RT, add 9 mL of wash buffer, and pool all the samples into one 50 mL tube. Count cells using the hemocytometer.
Note: You should expect approximately 6 × 106–12 × 106 cells per lung of a naïve 8-week-old female C57BL/6 mouse.
Negative selection
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in MACS buffer (107 cells/100 μL) with the adapted volume of MojoSort mouse CD45 nanobeads (10 μL/107 cells) according to the manufacturer protocol. Incubate for 15 min at 4 °C.
Wash the cells by adding MACS buffer up to 4 mL. Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant.
Resuspend in MACS buffer (5 × 107 cells/500 μL).
Filter cells through 70 μm cell strainer.
Proceed with magnetic negative selection: apply the cell suspension onto the prepared LD column (follow the Miltenyi LD column protocol). Collect flowthrough containing unlabeled cells after washes with 2 × 1 mL of MACS buffer.
Note: The unlabeled cells contain the non-immune cells.
ECs staining for FACS sorting
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in FACS buffer (107 cells/mL) with antibodies (anti-CD45, CD31, EpCAM) for 20 min at 4 °C in the dark.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet in 5 mL of PBS and repeat centrifugation.
Discard supernatant and perform viability staining by adding SYTOX Blue in FACS buffer (1 μL/1 mL/5 × 107 cells). Incubate for 5 min at RT in the dark.
Proceed to FACS sorting. Lung ECs are gated as CD45-CD31-Epcam+ living cells (Figure 2). Count the cells using the hemocytometer.
Figure 2. Gating strategy for FACS sorting of lung epithelial cells (ECs), described as CD45-CD31-EpCAM+ living cells after cell enrichment though MACS-negative selection (CD45).
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the cells at 5 × 103 cells/50 μL in complete culture medium (see Recipes).
Culture of MOs and lung ECs
Note: The presence of lung ECs with bone marrow MOs will help to obtain MOs in the process of differentiation into AMs.
Add 50 μL (5 × 104 cells) of bone marrow MOs and 50 μL (5 × 103 cells) of lung ECs into a well of a 96-well round-bottom plate.
Add 100 μL of complete culture medium containing 20 ng/mL recombinant murine GM-CSF (for a final concentration of 10 ng/mL) into each well and gently resuspend. Caution: Provide control wells without GM-CSF as well as conditions without ECs.
Incubate at 37 °C with 5% CO2 for three days.
Lung ILC2s isolation
Preparing lung cell suspension
Euthanize the mice and spray the skin with 75% ethanol.
Perfuse through the right ventricle with 5 mL of ice-cold PBS. Transfer the lung lobes into a GentleMACS tube containing lung digestion solution (see Recipes) and process with a gentleMACS dissociator (lung program 01); finally, incubate for 30 min at 37 °C. After digestion, run the lung program 02 with the gentleMACS dissociator. Filter the cell suspension through a 70 μm cell strainer into a 50 mL Falcon tube and top up to 25 mL with wash buffer.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet in 1 mL of RBC lysis buffer. Incubate for 5 min at RT and add 9 mL of wash buffer and pool five samples at a time into one 50 mL Falcon tube. Count cells using the hemocytometer.
Note: You should expect approximately 6 × 106–10 × 106 total cells per lung of a naïve 8-week-old female C57BL/6 mouse.
Negative selection
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet at 108 cells/mL of MACS buffer with lineage marker antibodies (anti-B220, CD11c, CD3, CD4, CD5, CD8α, F4/80, FcϵR1, CD49b, and Siglec-F) conjugated to APC fluorochrome, for 20 min at 4 °C in the dark.
Note: ILC2s represent approximately 1% of the lung leukocytes population; the lineage will target other immune cells such as T cells, B cells, NK cells, and macrophages, and then will improve the FACS sorting.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend the pellet in 5 mL of MACS buffer and repeat centrifugation.
Discard the supernatant and resuspend in MACS buffer (107 cells/100 μL) with the adapted volume of MojoSort APC nanobeads (10 μL/107 cells) (follow manufacturer’s protocol). Incubate on ice for 15 min.
Wash the cells by adding MACS buffer up to 4 mL. Centrifuge the cells at 250× g for 5 min at 4 °C. Discard the supernatant.
Resuspend in MACS buffer (5 × 107 cells/500 μL).
Filter cells through a 70 μm cell strainer.
Proceed with magnetic negative selection: apply cell suspension onto the prepared LD column (follow the Miltenyi LD columns protocol). Collect flowthrough containing unlabeled cells after washes with 2 × 1 mL of MACS buffer.
Caution: The unlabeled cells contain lung ILCs and non-immune cells.
ILC2s staining for FACS sorting
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in FACS buffer (107 cells/mL) with antibodies (anti-B220, CD11c, CD3, CD4, CD5, CD8α, F4/80, FcϵR1, Ly6G, Siglec-F, CD45, CD90.2, ST2, and CD25) for 20 min at 4 °C in the dark. Centrifuge the cells at 250× g for 5 min at 4 °C. Discard the supernatant. Resuspend the pellet in 5 mL of PBS and repeat centrifugation.
Discard supernatant and perform viability staining by adding SYTOX Blue to FACS buffer (1 μL/1 mL/5 × 107 cells). Incubate for 5 min at RT in the dark.
Proceed to FACS sorting using FACS Aria IIIu following manufacturer’s protocol. Lung ILC2s are gated as CD45+Lin-CD90.2+ST2+CD25+ living cells (Figure 3). Count cells using the hemocytometer.
Figure 3. Gating strategy for FACS sorting of lung group 2 lung innate lymphoid cells (ILC2s) after cell enrichment through MACS-negative selection against lineage markers (B220, CD11c, CD3, CD4, CD49b, CD5, CD8α, F4/80, FcϵR1, Gr1 and Siglec-F). ILC2s were identified as Lin-CD45+ST2+CD90.2+CD25+.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend cells at 2.5 × 103 cells/200 μL of ILC2 culture medium (see Recipes). Critical: IL-2 is mandatory to maintain ILC2s in ex vivo culture.
Co-culture of ILC2s with MOs and ECs
This step allows to investigate the role of interactions between ILC2s and MOs differentiating into AMs and to study the potential importance of a targeted ligand–receptor interaction.
Centrifuge the 96-well plate with the bone marrow MOs and lung ECs co-culture at 250× g for 5 min at 4 °C. Gently aspirate the supernatant without touching the pellet.
Add 200 μL of ILC2s (2.5 × 103 cells) in the ILC2 culture medium. Gently mix the cells together. Incubate at 37 °C in 5% CO2 environment for three days.
Caution: Provide control conditions without ILC2s.
Harvest and staining for FACS analysis
Note: A substantial fraction of bone marrow MOs differentiate into AM-like cells, as observed by the expression of CD11c. MO-derived AMs acquire the M2 marker Arg1 when co-cultured with ILC2s from a naïve mouse.
After three days, centrifuge the plate at 250× g for 5 min at 4 °C.
Perform viability staining by resuspending cell pellets in 100 μL/well in PBS with 1:1,500 Zombie AquaTM for 15 min at RT in the dark. Critical: Use fixable viability kit if the assessment requires intracytoplasmic staining such as Arg1.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant.
Resuspend cell pellets in 50 μL/well in FACS buffer with the following antibodies (anti-CD45, CD11c, Siglec-F, MHC-II, CD90, ST2) or any antibodies of targets of your interest, for 20 min at 4 °C in the dark.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 100 μL/well of PFA 2% for at least 15 min at 4 °C in the dark (it may be extended overnight).
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 100 μL/well of permeabilization buffer (see Recipes) and incubate at 37 °C for 15 min.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 100 μL/well of intracellular staining buffer (see Recipes) with antibody against intracellular antigens. Incubate for 30 min at RT in the dark.
Centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 200 μL/well of FACS buffer and repeat the centrifugation.
Discard the supernatant, resuspend in 200 μL/well of FACS buffer, and proceed to FACS analysis using a BD LSR Fortessa X-20 (Figure 4).
Figure 4. Lung ILC2s regulate differentiation and prone a M2 phenotype of MOAMs in vitro. (A) Representative flow cytometry plots of bone marrow monocytes (MOs), bronchoalveolar lavage alveolar macrophage (AMs) and bone marrow MOs differentiated to MOAMs from co-culture with lung epithelial cells (ECs) and lung group 2 lung innate lymphoid cells (ILC2s). (B) Heatmap of differentially expressed genes (False Discovery Rate, FRD) < 0.05; change in expression of over two-fold) in MO-derived macrophages isolated from co-cultures of MOs and ECs with or without ILC2s. Three genes with higher expression in the condition with ILC2s are highlighted.
Harvest for RNA sequencing
Note: For a deeper analysis, transcriptomic analysis can be performed on the ex vivo differentiated MO-derived AMs.
After step E1, centrifuge at 250× g for 5 min at 4 °C and discard the supernatant. Resuspend in 100 μL of FACS buffer with antibodies (anti-Ly6G, Ly6C, CD11c, CD45, CD90.2, ST2, and CD25) for 20 min at 4 °C in the dark. Centrifuge the cells at 250× g for 5 min at 4 °C. Discard the supernatant. Resuspend the pellet in 5 mL of PBS and repeat centrifugation.
Discard supernatant and perform viability staining by adding SYTOX Blue in FACS buffer (1 μL/1 mL/5 × 107 cells/). Incubate for 5 min at RT in the dark.
Proceed to FACS sorting using FACS Aria IIIu by following manufacturer’s protocol. MO-derived AMs are defined as Ly6G-, Ly6C-, autofluorescent, and CD11c+ living cells, and directly sorted in 1 mL of TRIzol before being snap frozen until future analysis (Figure 4).
Data analysis
Samples were sorted on a BD FACS Aria IIIu using FACSDiVa software. Flow cytometry data were analyzed using FlowJo to gate, quantify, and analyze the different populations.
Cells were sorted based on the expression of specific markers:
• Bone marrow MOs were gated as Ly6G-MHCII-CD3-CD11b+Ly6C+ living cells (Figure 1).
• Lung ECs were gated as CD45-CD31-Epcam+ living cells (Figure 2)
• Lung ILC2s were gated as CD45+Lin-CD90.2+ST2+CD25+ living cells (Figure 3).
For the readout, MO-derived AMs were defined as Ly6G-, Ly6C-, autofluorescent, and CD11c+ living cells, and ILC2s as SSC-AlowCD11c-CD90.2+ST2+ living cells.
Detailed information about data analyses appears in the original research article (Loos et al., 2023 in Figure 6 and Figure S4).
Validation of protocol
Loos et al. (2023). Dampening type 2 properties of group 2 innate lymphoid cells by a gammaherpesvirus infection reprograms alveolar macrophages. Science Immunology (Figure 6 and Figure S4).
General notes and troubleshooting
The entire procedure must be performed under a laminar flow hood to maintain sterile conditions and avoid any contamination.
ILC2s and epithelial cells are a minority population in the lung, so a minimum of five individuals per condition should be pooled to obtain enough cells of interest in order to generate triplicates.
Bead enrichment will facilitate FACS sorting, and negative enrichment will avoid early cell activation.
As ILC2 and epithelial cells phenotypes are directly impacted by environmental conditions, the outcome of the experiment largely depends on the environmental conditions to which mice have been subjected, as seen for example with MuHV-4 infection (Loos et al., 2023).
Acknowledgments
The protocol was adapted from the previously published paper: Loos et al. (2023).
This work was supported, in part, by University of Liège (VIRIMPRINT ARC to L.G.), by Fondation Léon Fredericq (grant to P.L.), by the Fonds de la Recherche Scientifique—Fonds National Belge de la Recherche Scientifique (F.R.S./FNRS, “credit de recherche” J007515F to L.G.; “projet de recherche” T.0195.16 to L.G.; research associate support for B.M.; and research fellow for P.L. and C.M.), by Institut MERIEUX starting grant (to L.G.), by EOS joint programme of F.R.S./FNRS Fonds wetenschapellijk onderzoek–Vlaanderen-FWO (EOS ID:30981113 and 40007527) (to L.G.); and by ERC Starting Grant (ERC-StG-2020 VIROME, ID:853608) (to B.M.).
Competing interests
The authors declare no competing interests.
Ethical considerations
All work in the development of this protocol was approved by the Committee on the Ethics of Animal Experiments of ULiège (permit numbers 1845).
References
Ebbo, M., Crinier, A., Vély, F. and Vivier, E. (2017). Innate lymphoid cells: major players in inflammatory diseases. Nat. Rev. Immunol. 17(11): 665–678.
Kerscher, B., Barlow, J. L., Rana, B. M., Jolin, H. E., Gogoi, M., Bartholomew, M. A., Jhamb, D., Pandey, A., Tough, D. F., van Oosterhout, A. J. M., et al. (2019). BET Bromodomain Inhibitor iBET151 Impedes Human ILC2 Activation and Prevents Experimental Allergic Lung Inflammation. Front. Immunol. 10: e00678.
Loos, P., Baiwir, J., Maquet, C., Javaux, J., Sandor, R., Lallemand, F., Marichal, T., Machiels, B. and Gillet, L. (2023). Dampening type 2 properties of group 2 innate lymphoid cells by a gammaherpesvirus infection reprograms alveolar macrophages. Sci. Immunol. 8(80): eabl9041.
Moro, K., Yamada, T., Tanabe, M., Takeuchi, T., Ikawa, T., Kawamoto, H., Furusawa, J., Ohtani, M., Fujii, H., Koyasu, S., et al. (2010). Innate production of TH2 cytokines by adipose tissue-associated c-Kit+Sca-1+ lymphoid cells. Nature 463(7280): 540–544.
Oliphant, C. J., Hwang, Y. Y., Walker, J. A., Salimi, M., Wong, S. H., Brewer, J. M., Englezakis, A., Barlow, J. L., Hams, E., Scanlon, S. T., et al. (2014). MHCII-Mediated Dialog between Group 2 Innate Lymphoid Cells and CD4+ T Cells Potentiates Type 2 Immunity and Promotes Parasitic Helminth Expulsion. Immunity 41(2): 283–295.
Saluzzo, S., Gorki, A. D., Rana, B. M., Martins, R., Scanlon, S., Starkl, P., Lakovits, K., Hladik, A., Korosec, A., Sharif, O., et al. (2017). First-Breath-Induced Type 2 Pathways Shape the Lung Immune Environment. Cell Rep. 18(8): 1893–1905.
Surace, L., Doisne, J. M., Croft, C. A., Thaller, A., Escoll, P., Marie, S., Petrosemoli, N., Guillemot, V., Dardalhon, V., Topazio, D., et al. (2021). Dichotomous metabolic networks govern human ILC2 proliferation and function. Nat. Immunol. 22(11): 1367–1374.
Spits, H. and Di Santo, J. P. (2011). The expanding family of innate lymphoid cells: regulators and effectors of immunity and tissue remodeling. Nat. Immunol. 12(1): 21–27.
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© 2023 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 isolation and culture > Co-culture
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4,819 | https://bio-protocol.org/en/bpdetail?id=4819&type=0 | # Bio-Protocol Content
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This protocol has been updated. See the update notice.
Peer-reviewed
Computational Analysis of Plasma Lipidomics from Mice Fed Standard Chow and Ketogenic Diet
AS Amy L. Seufert *
JH James W. Hickman *
JC Jaewoo Choi
BN Brooke A. Napier
(*contributed equally to this work)
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4819 Views: 1043
Reviewed by: Alka MehraAnand Sharma Sharma Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in eLIFE Oct 2022
Abstract
Dietary saturated fatty acids (SFAs) are upregulated in the blood circulation following digestion. A variety of circulating lipid species have been implicated in metabolic and inflammatory diseases; however, due to the extreme variability in serum or plasma lipid concentrations found in human studies, established reference ranges are still lacking, in addition to lipid specificity and diagnostic biomarkers. Mass spectrometry is widely used for identification of lipid species in the plasma, and there are many differences in sample extraction methods within the literature. We used ultra-high performance liquid chromatography (UPLC) coupled to a high-resolution hybrid triple quadrupole-time-of-flight (QToF) mass spectrometry (MS) to compare relative peak abundance of specific lipid species within the following lipid classes: free fatty acids (FFAs), triglycerides (TAGs), phosphatidylcholines (PCs), and sphingolipids (SGs), in the plasma of mice fed a standard chow (SC; low in SFAs) or ketogenic diet (KD; high in SFAs) for two weeks. In this protocol, we used Principal Component Analysis (PCA) and R to visualize how individual mice clustered together according to their diet, and we found that KD-fed mice displayed unique blood profiles for many lipid species identified within each lipid class compared to SC-fed mice. We conclude that two weeks of KD feeding is sufficient to significantly alter circulating lipids, with PCs being the most altered lipid class, followed by SGs, TAGs, and FFAs, including palmitic acid (PA) and PA-saturated lipids. This protocol is needed to advance knowledge on the impact that SFA-enriched diets have on concentrations of specific lipids in the blood that are known to be associated with metabolic and inflammatory diseases.
Key features
• Analysis of relative plasma lipid concentrations from mice on different diets using R.
• Lipidomics data collected via ultra-high performance liquid chromatography (UPLC) coupled to a high-resolution hybrid triple quadrupole-time-of-flight (QToF) mass spectrometry (MS).
• Allows for a comprehensive comparison of diet-dependent plasma lipid profiles, including a variety of specific lipid species within several different lipid classes.
• Accumulation of certain free fatty acids, phosphatidylcholines, triglycerides, and sphingolipids are associated with metabolic and inflammatory diseases, and plasma concentrations may be clinically useful.
Graphical overview
Keywords: Lipidomics Mass spectrometry Ketogenic diet Free fatty acids Phosphatidylcholines Triglycerides Sphingolipids Circulating lipids
Background
Circulating lipids in the blood, specifically cholesterol and triglycerides (TAGs), are measured in the clinic to determine cardiovascular disease (CVD) risk (Sarwar et al., 2007; Ference et al., 2017). Additional classes of lipids, including free fatty acids (FFAs), phosphatidylcholines (PCs), and sphingolipids (SGs), are associated with metabolic, inflammatory, and infectious diseases; however, reference ranges have not yet been determined. Within lipid classes, specific lipid species may also be useful as biomarkers for early detection of CVD, COVID-19, sepsis, cancer, neurodegenerative diseases, and chronic obstructive pulmonary disease (Kubicek-Sutherland et al., 2017; Kohno et al., 2018; Liu et al., 2020; Casas-Fernández et al., 2022; Ciccarelli et al., 2022). High-throughput lipidomics is a valuable tool for identifying a diversity of lipid species and their relative concentrations within a small plasma sample (< 50 μL). Here, we describe our protocol for analyzing a large dataset, collected via ultra-high performance liquid chromatography (UPLC) coupled to a high-resolution hybrid triple quadrupole-time-of-flight (QToF) mass spectrometry (MS), encompassing 15 different lipid classes, with a focus on four clinically relevant lipid species: FFAs, TAGs, PCs, and SGs (Choi et al., 2015; Seufert et al., 2022). The plasma samples measured were harvested from mice fed a standard chow (SC) or a ketogenic diet (KD) for two weeks. Using this protocol, we were able to determine relative abundances of FFAs, TAGs, PCs, and SGs, and take a snapshot of diet-induced lipidomic profiles within mice. This protocol may be used to analyze any omics dataset.
Currently, common blood lipid panels used in the clinic lack detail regarding specific lipid species found in the blood, measuring only low-density, very-low-density, and high-density lipoprotein cholesterol and total TAGs in order to identify hyperlipidemia, CVD risk and metabolic syndrome (Lee et al., 2023). Although plasma cholesterol and total TAG concentrations are useful in the prevention and identification of hyperlipidemia, CVD, and metabolic syndrome, many other circulating lipids exist that are known to correlate with inflammatory and metabolic dysfunction. For example, dietary saturated fats such as palmitic acid (PA) have recently been shown to sensitize innate immune cells to microbial ligand exposure in vitro and in vivo, and serum PA concentrations positively correlate with insulin resistance in humans (Perreault et al., 2014; Seufert et al., 2022). The methods outlined in this protocol and our corresponding publication (Seufert et al., 2022) show the efficacy of using high-throughput lipidomics to compare relative concentrations of specific lipid species in the blood of SC- and KD-fed mice. Our results highlight the importance of this protocol for future studies in mice or humans that aim to identify lipid changes in the blood due to SFA-enriched diets and supports the advancement of personalized nutrition for humans suffering from metabolic and inflammatory diseases.
UPLC-QToF MS/MS identifies hundreds of lipid species within a single plasma sample; measuring the chromatographic peak area under the curve (AUC) for each lipid allows for relative comparisons within and between samples (Choi et al., 2015). Absolute concentrations may be determined for some lipids; however, specific MS standards at additional costs are required, and absolute concentrations were not measured in this study. Due to the numerous combinations of different headgroups and fatty acids including chain length, number, and position of double bonds, the separation and identification of isobaric lipids that have identical molecular formulas but structural differences are limited (Batarseh et al., 2018). For instance, TAG (16:0/18:1/20:4) and TAG (18:1/18:2/18:2) are isobaric lipids. These lipids are chromatographically separated and identified by MS/MS fragmentation. However, TAG (16:0/18:1/18:2) and TAG (16:0/18:2/18:1) are not chromatographically separated with no difference in MS/MS spectrum because of the same fatty acyl group on different locations of the glycerol backbone (Figure 1). The ion mobility or ozone-induced dissociation techniques can afford information on double bond and location of fatty acyl group on lipids. Lastly, aside from essential FAs that are known to be only derived from exogenous sources (omega-3 and omega-6), lipidomics data does not indicate whether plasma lipids entered the bloodstream directly following digestion of lipids or were produced endogenously by the host. Thus, proper controls are required when studying diet-dependent effects.
Figure 1. Isobaric structure of triglyceride as shown by UPLC-QToF MS/MS: TAG (16:0/18:1/18:2) or TAG (16:0/18:2/18:1). Separation and identification of isobaric lipids with identical molecular formulas is limited due to structural differences that do not show chromatographic separation. (A) Extract ion chromatogram of TAG (16:0/18:1/18:1) or TAG (16:0/18:2/18:1). (B) ToF-MS isotopic pattern of m/z 874.7858. (C) ToF-MS/MS of m/z 874.7858, which shows ammonium adduct in positive ion mode.
Materials and reagents
Biological materials
Age-matched (4–6 weeks) female wildtype BALB/c mice; JAX stock #000651 (see General note 1)
Reagents
Heparin sodium (VWR, catalog number: AAA16198-03)
1× PBS (Thermo Fisher, catalog number: 20012050)
Recipes
Heparin sodium, diluted to 100 IU/mL in 1× PBS, ~5 mL
Laboratory supplies
PicoLab Mouse Diet 20 (Irradiated chow; product 5058), softened with water for first week of acclimation
Compressed CO2 gas in cylinders for euthanizing mice (fill rate: 30%–70% of chamber vol/min)
Standard chow mouse diet (Envigo, TD.08485) (see General note 1)
Ketogenic mouse diet (Envigo, TD.180423) (see General note 1)
BD Vacutainer blood collection tubes (BD, catalog number: 366667)
3 mL BD Luer-Lok syringe with attached needle (25 G × 5/8 in.) (BD, catalog number: 309570)
Axygen Maxy Clear Snaplock microcentrifuge tubes (VWR, catalog number: 10011-700)
Parafilm M wrapping film (Fisher Scientific, catalog number: S37440)
Ice and ice bucket
Equipment
Tabletop centrifuge for microcentrifuge tubes set at 4 °C
Computer with a Microsoft OS (version dependent on PeakView and MultiQuant version utilized)
Euthanasia chamber for mice
Software and datasets
PeakView version 1.2.1 (Sciex) (see General note 2)
MultiQuant version 3.0.2 (Sciex) (see General note 2)
Excel (Microsoft Office 2019) (see General note 2)
R Statistical Software (https://www.r-project.org)
RStudio open-source edition, Boston, MA, USA (https://www.rstudio.com)
GraphPad Prism 9 (www.graphpad.com)
Procedure
Allow mice to acclimate undisturbed to research facility with soft food for one week prior to changing diets.
Separate mice and change diets to SC or KD two weeks prior to plasma collection date.
Note: The KD is soft and requires refrigeration and daily food changes; cages must be cleaned or replaced every three days.
Immediately prior to euthanizing mice, prepare heparin solution (~5 mL) and coat syringes for cardiac punctures by aspirating and expelling the solution with each syringe. The same heparin solution can be used for multiple syringes. After coating, syringes may be placed on a sterile surface resting on needle caps. Ensure that the needles are bevel up and ready for use.
Prepare blood collection tubes and ice bucket and set tabletop centrifuge temperature to 4 °C.
Euthanize mice one at a time with CO2 and prior to cervical dislocation, place mouse supine on bench, and quickly perform cardiac puncture using 3 mL BD Luer-Lok syringe with attached 25 G needle.
It is important that the needle gauge used for this procedure is between 23 and 25 G, in order to avoid hemolysis. Hemolysis is the destruction of red blood cells, and it has been shown to significantly impact levels of certain lipid species in the blood (Burla et al., 2018).
Transfer blood (200–700 μL depending on size of mouse) to BD Vacutainer blood collection tubes and keep on ice.
Transfer blood samples to microcentrifuge tubes and centrifuge at 1,500× g for 20 min at 4 °C.
Collect transparent plasma (supernatant) and transfer to a fresh microcentrifuge tube.
Seal tubes with parafilm and store at -80 °C or ship on dry ice for sample processing to: Jan F. Stevens & Jaewoo Choi. Linus Pauling Institute, Oregon State University, Corvallis, OR, USA (Choi et al., 2015). Frozen plasma (20 μL) was extracted with lipidomics extraction solvent (480 μL, methylene chloride:methanol:isopropanol = 25:10:65, v/v/v + 0.1% BHT), vortexed for 30 s, centrifuged for 10 min at 13,000 rpm at 4 °C. The aliquot (95 μL) was transferred into mass spectrometry analysis tubes and SPLASH LipidoMix (Avanti Lipids) as internal standard mixture (5 μL) was spiked. UPLC was performed using a 1.7 μm particle, 2.1 × 100 mm, CSH C18 Column (Waters, Milford, MA, United States) coupled to a quadrupole TOF mass spectrometer (AB SCIEX, TripleTOF 5600) operated in information-dependent MS/MS acquisition mode. LC and MS conditions were developed as described previously by Choi et al. (2015) with some modifications. For positive ion mode LC-QToF-MS/MS, the mobile phases consisted of (A) 60:40 (v/v) acetonitrile: water with ammonium formate (10 mM) and formic acid (0.1%) and (B) 90:10 (v/v) isopropanol:acetonitrile with ammonium formate (10 mM) and formic acid (0.1% formic acid). For analyses run in the negative ion mode, ammonium acetate (10 mM) was used as the modifier.
Data analysis
Identification and quantification of lipidomics
Process the lipidomics data with PeakView 1.2.1 and quantify lipids with MultiQuant software version 3.0.2 (see General note 3, Choi et al., 2015). Raw MS files (*.wiff) were imported and processed by the program PeakView (Sciex) (Figure 2, Figure 3). PeakView detects spectral features using extract ion chromatogram (XIC) lists from our in-house library of lipids (each defined by a unique chromatographic retention time and accurate mass, MS/MS fragmentation, and isotopic pattern; Figure 3, B and C). Each peak was integrated by MultiQuant (Sciex) software (Figure 4). Peak integration is the quantification step whereby the peak area of an identified lipid is calculated. Peak area is referred to in this protocol as the AUC and this value is proportional to the quantity of the identified lipid.
Save lipidomics data as Excel or .csv files. Normalization can be done in Excel with the following formula: peak area ratio = peak area of identified lipid/peak area of labeled internal standard. Each integrated chromatogram was normalized manually with the use of an internal standard peak purchased from Avanti lipids (SPLASH LipidoMix) (Figure 4).
Figure 2. Lipidomics data processing
Figure 3. Lipid [PC (16:0/18:2)] identification using PeakView software, which explores and interprets qualitative data. (A) Extract ion chromatogram (XIC) of PC (16:0/18:2). (B) XIC manager displays in the table including found mass, mass error, found retention time, formula, adduct, and exact mass. (C) TOF-MS isotopic pattern of m/z 758.5694, which shows PC (16:2/18:0). (D) TOF-MS/MS of m/z 758.5694 as protonated adduct. The m/z 184 represents a unique fragment ion (protonated phosphocholine) corresponding to phosphatidylcholine.
Figure 4. Chromatographic peak area integration using MultiQuant software. (A) Analyte pane includes identified PCs list. (B) Result pane includes each sample name and peak area counts with retention time. (C) Chromatogram review in each sample. The peak can be automatically or manually integrated. AUC values are highlighted in yellow.
Bioinformatic analysis in R
In R, scale each lipid type or class dataset with the scale() function. For mean centering, replace NA values with the mean of the particular variable. For more detailed information on the code see General note 4.
Perform a PCA analysis with the prcomp() function.
PCA analysis is a technique for multivariable data that performs dimension reduction to represent the most important and impactful information as principal components. This allows the data to be visually observed for patterns of similarity (Abdi and Williams, 2010).
Visualize the first two principal components for the dataset for each lipid type with the fviz_pca_ind() function from the factoextra package (Kassambara and Mundt, 2017). Color samples by diet group and create a confidence ellipse around each sample group (Figure 5A).
Set addEllipses = TRUE, ellipse.type = “confidence,” and ellipse.level = 0.95 to create confidence ellipses.
Investigate strong separation between groups in the 2D space indicated by no overlap between the concentration ellipses. Sample groups with low separation for lipid types will have overlapping ellipses and data points.
Investigate specific lipid variables contributing to separation with fviz_pca_biplot() to create a biplot (Wickham, 2016) (Figure 5B).
Choose the top five contributing variables to view with select.var = list(contrib = 5). Adjust the number to increase or decrease the desired number of variables.
Phosphatidylcholine (PC) variables are lipids identified within each plasma sample represented by vectors that are close together and that form small angles with one another because they are positively correlated. Variables point in the direction of the principal component(s) they strongly influence. In the PC biplot, all of the variables point along the PC1 axis towards the KD samples, indicating that the KD samples are relatively more enriched in these PCs. This enrichment strongly contributes to the separation of the KD and SC, as the SC samples group towards the opposite side of PC1, away from where the variables are pointing.
Perform a heatmap analysis with hierarchical clustering using the pheatmap() function on the dataset (Kolde, 2018). Identify lipid types with clear clustering between diet groups (Figure 5C).
Figure 5. Lipidomics data analysis (see General note 4). (A) PCA plot of phosphatidylcholine composition identified via LC-QToF MS/MS between age-matched (6–8 weeks) BALB/c female mice. Dots represent individual mice with colors corresponding to a standard chow (SC) (grey) or ketogenic diet (KD) (orange) with the mean of each diet group surrounded by a 95% confidence ellipse. (B) Biplot labeled with the top five phosphatidylcholines contributing to sample separation in 2D space. (C) Heatmap analysis of phosphatidylcholine composition between individual mice.
Perform t-tests in R or, alternatively, GraphPad Prism to compare significance of lipids between sample diet groups (Table 1).
Table 1. Phosphatidylcholine (PC) data and significance for standard chow (SC)- and ketogenic diet (KD)-fed mice shown in Figure 5. Statistical significance determined by unpaired two-tailed t-test between SC and KD groups, n = 3, AUC = area under the curve.
Phosphatidylcholine (PC) SC (mean AUC) KD (mean AUC) p-value
PC 37:4 107,470 317,878 0.000183788
PC 31:1 2,019 11,590 0.000298222
PC(18:1_20:5) 350,900 1,037,459 0.000400427
PC(18:0_22:6) 1,532,375 5,075,115 0.000436107
PC(14:0_14:0) 853 13,466 0.000506701
PC(18:1_18:1) 9,425,085 46,602,806 0.000613854
PC(p-40:6)/PC(o-40:7) 35,344 72,388 0.000634691
PC(p-38:3)/PC(o-38:4)B 56,172 137,571 0.000665094
PC(18:1_20:3) 8,158,408 25,437,034 0.000693246
PC(18:0_20:4) 11,279,355 33,755,056 0.000713162
PC(18:1_18:2) 2,729,249 10,297,513 0.000744278
PC(16:0_20:3) 3,719,482 14,331,321 0.000806004
PC(p-40:3)/PC(o-40:4) 21,627 43,188 0.000929417
PC(18:0_20:3) 568,891 3,223,095 0.000986705
PC(16:0_18:3) 134,495 426,137 0.001061905
PC(18:2_18:3)B 203,100 527,797 0.001373022
PC(18:0_18:1) 915,812 6,979,001 0.001377977
PC(p-40:5)/PC(o-40:6) 52,554 141,136 0.001658554
PC 42:10 32,839 62,822 0.001987255
PC 30:1 2,889 15,436 0.00199227
PC 31:0 41,226 218,739 0.002324314
PC(16:0_18:2) 30,188,177 53,002,677 0.002454378
PC(18:0_22:5) 78,146 250,710 0.002602027
PC(14:0_16:0) 120,007 663,490 0.003213188
PC(14:0_20:4) 13,280 39,412 0.003952569
PC(p-38:3)/PC(o-38:4)A 20,181 64,890 0.004202996
PC(p-32:1)/PC(o-32:2) 9,363 20,474 0.004275232
PC 38:1 16,263 42,593 0.005217356
PC 37:6 11,239 35,212 0.005279249
PC(16:0_20:4) 28,931,051 53,402,365 0.005752917
PC 37:5 47,580 116,619 0.005997394
PC(p-38:5)/PC(o-38:6) 158,770 307,694 0.008146112
PC(18:2_18:2) 22,306,084 41,033,132 0.008743996
PC(16:0_22:6) 15,035,676 25,864,778 0.010461728
PC(p-34:1)/PC(o-34:2) 40,587 110,459 0.011318898
PC(p-42:5)/PC(o-42:6) 22,624 47,197 0.015107157
PC(16:0_18:1) 14,005,109 28,988,308 0.01571763
PC(18:2_20:4) 11,043,269 19,331,439 0.015779177
PC(18:3_18:3) 7,190 15,091 0.017188207
PC(p-34:2)/PC(o-34:3) 12,575 34,342 0.017422368
PC(14:0_18:2) 42,618 102,328 0.019365582
PC(p-38:4)/PC(o-38:5) 162,550 299,056 0.022117396
PC(p-34:0)/PC(o-34:1) 82,298 132,534 0.023567292
PC(p-36:4)/PC(o-36:5) 128,403 252,282 0.02378729
PC(20:0_18:2) 118,694 224,791 0.025843285
PC(16:0_22:4) 566,758 1,098,135 0.02940295
PC(16:1_18:2) 220,162 292,355 0.033872581
PC(p-38:6)/PC(o-38:7) 69,692 100,671 0.035179669
PC(p-36:1)/PC(o-36:2) 20,699 39,588 0.037242501
PC(18:2_18:3)A 87,828 38,090 0.039675013
Validation of protocol
This protocol was used to generate Figure 3 and associated supplementary files within the following publication: Seufert et al. (2022).
General notes and troubleshooting
Researchers may want to determine which anticoagulant will best fit their experimental design (e.g., heparin vs. EDTA), as some are known to influence lipid extraction and MS analysis (Gonzalez-Covarrubias et al., 2013). Importantly, the same anticoagulant and amount should be used throughout the study in order to avoid potential differences that may occur due to the use of anticoagulants in sample collection.
Alternative mouse models and diets can be used specific to the experimental question.
Newer versions of software may also be used.
Knowledge of MultiQuant software is required.
An in-depth knowledge of R and bioinformatics is required for the data analysis. All code and datasets for Figure 5 can be accessed at https://github.com/hickman6/Lipidomics_Code.
Acknowledgments
Lipidomics data was generated via LC-QToF MS/MS by Dr. Jaewoo Choi at The Linus Pauling Institute, Oregon State University, Corvallis, OR, USA. This protocol was originally used for the following publication: Seufert et al. (2022). This study was supported by National Institute of General Medical Sciences (NIGMS) grant 5R35GM133804-02 to B.A.N. The QTOF mass spectrometer was funded by an instrument grant from the NIH: ABSciex Triple ToF 5600 NIH #1S10RR027878-01.
Competing interests
No competing interests declared.
Ethical considerations
All animal studies were performed in accordance with NIH guidelines, the Animal Welfare Act, and US federal law. All animal experiments were approved by the Oregon Health and Sciences University (OHSU) Department of Comparative Medicine or Oregon State University (OSU) Animal Program Office and were overseen by the Institutional Care and Use Committee (IACUC) under Protocol IDs #IP00002661 and IP00001903 at OHSU and #5091 at OSU. Conventional animals were housed in a centralized research animal facility certified by OHSU.
References
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Batarseh, A. M., Abbott, S. K., Duchoslav, E., Alqarni, A., Blanksby, S. J. and Mitchell, T. W. (2018). Discrimination of isobaric and isomeric lipids in complex mixtures by combining ultra-high pressure liquid chromatography with collision and ozone-induced dissociation. Int. J. Mass Spectrom. 431: 27–36.
Burla, B., Arita, M., Arita, M., Bendt, A. K., Cazenave-Gassiot, A., Dennis, E. A., Ekroos, K., Han, X., Ikeda, K., Liebisch, G., et al. (2018). MS-based lipidomics of human blood plasma: a community-initiated position paper to develop accepted guidelines. J. Lipid Res. 59(10): 2001–2017.
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Controlled Level of Contamination Coupled to Deep Sequencing (CoLoC-seq) Probes the Global Localisation Topology of Organelle Transcriptomes
AS Anna Smirnova
DJ Damien Jeandard
Alexandre Smirnov
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4820 Views: 415
Reviewed by: Alessandro DidonnaMarina Sánchez PetidierRajesh D Gunage
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Original Research Article:
The authors used this protocol in Nucleic Acids Research Dec 2022
Abstract
Information on RNA localisation is essential for understanding physiological and pathological processes, such as gene expression, cell reprogramming, host–pathogen interactions, and signalling pathways involving RNA transactions at the level of membrane-less or membrane-bounded organelles and extracellular vesicles. In many cases, it is important to assess the topology of RNA localisation, i.e., to distinguish the transcripts encapsulated within an organelle of interest from those merely attached to its surface. This allows establishing which RNAs can, in principle, engage in local molecular interactions and which are prevented from interacting by membranes or other physical barriers. The most widely used techniques interrogating RNA localisation topology are based on the treatment of isolated organelles with RNases with subsequent identification of the surviving transcripts by northern blotting, qRT-PCR, or RNA-seq. However, this approach produces incoherent results and many false positives. Here, we describe Controlled Level of Contamination coupled to deep sequencing (CoLoC-seq), a more refined subcellular transcriptomics approach that overcomes these pitfalls. CoLoC-seq starts by the purification of organelles of interest. They are then either left intact or lysed and subjected to a gradient of RNase concentrations to produce unique RNA degradation dynamics profiles, which can be monitored by northern blotting or RNA-seq. Through straightforward mathematical modelling, CoLoC-seq distinguishes true membrane-enveloped transcripts from degradable and non-degradable contaminants of any abundance. The method has been implemented in the mitochondria of HEK293 cells, where it outperformed alternative subcellular transcriptomics approaches. It is applicable to other membrane-bounded organelles, e.g., plastids, single-membrane organelles of the vesicular system, extracellular vesicles, or viral particles.
Key features
• Tested on human mitochondria; potentially applicable to cell cultures, non-model organisms, extracellular vesicles, enveloped viruses, tissues; does not require genetic manipulations or highly pure organelles.
• In the case of human cells, the required amount of starting material is ~2,500 cm2 of 80% confluent cells (or ~3 × 108 HEK293 cells).
• CoLoC-seq implements a special RNA-seq strategy to selectively capture intact transcripts, which requires RNases generating 5′-hydroxyl and 2′/3′-phosphate termini (e.g., RNase A, RNase I).
• Relies on nonlinear regression software with customisable exponential functions.
Graphical overview
Keywords: CoLoC-seq Subcellular transcriptomics RNA localisation Cell fractionation RNase Enzymatic kinetics Nonlinear regression Northern blotting Membrane-bounded organelle Mitochondria
Background
Knowing the localisation topology of transcripts with respect to organelle membranes (inside vs. outside) is critical for the understanding of RNA transactions in various subcellular locations. Selective RNA packaging into viral particles, extracellular vesicles, and ribonucleoproteins (RNPs) attracted much attention over the last two decades (Bresnahan and Shenk, 2000; K. Wang et al., 2010; Arroyo et al., 2011; Routh et al., 2012; Jeppesen et al., 2019; Murillo et al., 2019; Lécrivain and Beckmann, 2020; Gruner and McManus, 2021). Even more intriguing are the intricate interactions between the genetic systems of the nucleus, mitochondria, and plastids inside eukaryotic cells (Woodson and Chory, 2008; L. Levin et al., 2014; Quirós et al., 2016). These organelles possess their own genomes and locally produced transcriptomes. However, in many species, select nuclear-encoded RNAs (primarily tRNAs) enter mitochondria to participate in translation, blurring borders between transcriptomes (Schneider, 2011; Sieber et al., 2011; Jeandard et al., 2019). The scope of such RNA relocation pathways remains insufficiently understood. Therefore, robust genome-wide approaches are required to obtain comprehensive and reliable local transcriptomes.
Confident assignment of RNA localisation topology is challenging, and many approaches have been proposed (Jeandard et al., 2019). In fractionation-based techniques, organelles of interest are purified and treated with a non-specific RNase to degrade contaminant transcripts sticking to their surface. The remaining RNAs, detected by northern blotting, RT-PCR, or RNA-seq, are considered as residing inside the organelles. Although this strategy remains most widely used due to its simplicity and applicability to virtually any species, including non-model, genetically intractable organisms, cell cultures, and tissues (Mercer et al., 2011; Geiger and Dalgaard, 2017), its simple experimental setup turned out to be over-optimistic: some short, structured transcripts embedded in stable RNPs resist degradation, leading to prohibitively high false-positive rates. This caveat is mostly resolved by proximity labelling approaches that use organelle-restricted biochemical tagging of RNA molecules in situ to enable their selective enrichment and identification (Kaewsapsak et al., 2017; Fazal et al., 2019; P. Wang et al., 2019; Zhou et al., 2019; Medina-Munoz et al., 2020; Engel et al., 2021). However, these methods are usually biased against shorter, non-polyadenylated, and lowly abundant transcripts, and require genetic introduction of engineered tagging enzymes targeted to the organelle of interest, which limits their application to model species with tractable genomes and well-characterised protein localisation pathways.
Here, we describe Controlled Level of Contamination coupled to deep sequencing (CoLoC-seq), which marries the accessibility and generality of fractionation-based approaches with the selectivity and robustness of proximity labelling techniques (Jeandard et al., 2023). In a standard CoLoC-seq pipeline (Figure 1, the blue branch), a preparation of intact organelles of interest is split into a suite of samples subjected to a gradient of RNase concentrations. This creates transcript-specific digestion kinetics, amenable to straightforward mathematical modelling, which tells whether a certain RNA fully partakes in the reaction (as expected for contaminants) or if there is a pool of unavailable molecules protected from the RNase. In a parallel Mock CoLoC-seq experiment (Figure 1, the orange branch), the same organelles are first mildly lysed with a detergent to solubilise membranes and then split in a series of identical samples for RNase treatment. Therefore, by measuring the same digestion kinetics in the mildly lysed organelles, one can determine whether the RNA protection is conferred by the organellar membranes or by unrelated factors, such as proteins or intricate structures (such unreactive RNAs are likely false positives).
Successfully tested on human mitochondria, CoLoC-seq outperformed other fractionation-based and proximity labelling approaches, especially in assigning the localisation topology of shorter non-coding RNAs. It can be applied to any RNase-impermeable entity, including eukaryotic organelles, endosymbionts, enveloped and some non-enveloped viruses, and extracellular vesicles.
Figure 1. Overview of the protocol and its main steps. The part in grey covers the isolation of crude mitochondria, blue and orange correspond to the CoLoC and Mock CoLoC procedures, respectively, and green is data acquisition and analysis. The stop signs show the steps where the protocol can be safely interrupted without compromising the outcome of the experiment.
Materials and reagents
Biological materials
The CoLoC-seq was performed on mitochondria isolated from the human Flp-In T-REx 293 cells (Thermo Fisher Scientific, catalog number: R78007). One complete set of CoLoC-seq and Mock CoLoC-seq samples requires approximately 2,500 cm2 of nearly confluent cells (equivalent to ~3 × 108 HEK293 cells) devoid of mycoplasma contamination. For application of CoLoC-seq to other systems and cellular compartments, the optimal amount of starting material should be determined empirically.
Reagents
EDTA (Sigma-Aldrich, catalog number: E4884)
Tris base (Sigma-Aldrich, catalog number: 11814273001)
Sucrose (Sigma-Aldrich, catalog number: S0389)
Sorbitol (Sigma-Aldrich, catalog number: S1876)
NaCl (Sigma-Aldrich, catalog number: S9888)
Bovine serum albumin, lyophilized, fatty acid free (Euromedex, catalog number: 1035-70-C)
NaOH (Sigma-Aldrich, catalog number: 655104)
Bromophenol blue (Sigma-Aldrich, catalog number: B0126)
Deionised formamide (Sigma-Aldrich, catalog number: S4117)
Dulbecco’s phosphate buffered saline (DPBS 1×) (Sigma-Aldrich, catalog number: D5773)
20% sodium dodecyl sulfate (SDS) (Euromedex, catalog number: EU-0660-B)
n-dodecyl-β-D-maltoside (Sigma-Aldrich, catalog number: D4641); store at -20 °C
Glycogen (Sigma-Aldrich, catalog number: G8876); store at 4 °C
TBE 10× (Euromedex, catalog number: ET020-C)
Urea (Sigma-Aldrich, catalog number U5378)
40% acrylamide/bis-acrylamide (19:1) (Carl Roth, catalog number: A516.1); store at 4 °C
Ammonium persulfate (Euromedex, catalog number: EU0009-B)
N,N,N’,N’-Tetramethyl ethylenediamine (TEMED) (Euromedex, catalog number: 50406)
Ethidium bromide 1% (Biosolve BV, catalog number: 05412341); store at 4 °C
SSC buffer 20× (Euromedex, catalog number: BI-D0623-1L)
Denhardt’s solution 50× (Thermo Fisher Scientific, catalog number: 750018); store at -20 °C
TE (Tris-EDTA buffer), pH 7.4 (10×) (Euromedex, catalog number: BI-USD8211-1L)
DNase I (Thermo Fisher Scientific, catalog number: EN0525); store at -20 °C
RNase A, DNase- and protease-free, 10 mg/mL (Thermo Fisher Scientific, catalog number: EN0531); store at -20 °C
SUPERase In RNase inhibitor (Thermo Fisher Scientific, catalog number: AM2694); store at -20 °C
AMPure XP kit (Beckman Coulter, catalog number: A63881)
Bradford assay ROTI Nanoquant (Carl Roth, catalog number: K880.1); store at 4 °C
TRIzol reagent (Thermo Fisher Scientific, catalog number: 15596026); store at 4 °C
γ-[32P]-ATP (10 Ci/L, 3,000 Ci/mmol) (PerkinElmer, catalog number: BLU002A100UC). Should be used within approximately one month (32P half-life is 14.268 days); store at -20 °C
Polynucleotide kinase (PNK) and 10× kinase reaction buffer A (Promega, catalog number: M4101); store at -20 °C
Chloroform (Carl Roth, catalog number: 6340.4)
Isopropanol (Carl Roth, catalog number: CP41.1)
Ethanol (Dutscher, Carlo Erba, catalog number: 3086072-CER)
Solutions
Sterile 0.1 M EDTA (see Recipes)
Sterile EDTA-DPBS (see Recipes)
Tris-HCl 0.1 M, pH 6.7 (see Recipes)
Sucrose 3.3 M (see Recipes)
Sorbitol 3 M (see Recipes)
NaCl 1 M (see Recipes)
NaOH 6% (see Recipes)
Buffer A (see Recipes)
Buffer B (see Recipes)
Buffer C (see Recipes)
Buffer D (see Recipes)
Buffer E (see Recipes)
Buffer F (see Recipes)
Buffer H (see Recipes)
RNA loading buffer (see Recipes)
RNA denaturing polyacrylamide gel (see Recipes)
Ammonium persulfate 10% (see Recipes)
Pre-hybridisation buffer (see Recipes)
Hybridisation buffer (see Recipes)
Washing buffer (see Recipes)
Stripping buffer (see Recipes)
Ethanol 80% (see Recipes)
RNase A dilutions in buffer D (see Recipes)
Glycogen 20 μg/μL (see Recipes)
Ethidium bromide 0.0001% (see Recipes)
Recipes
Sterile 0.1 M EDTA (store at 4 °C)
Stir 37.2 g of EDTA in 800 mL of H2O. Add NaOH to adjust pH to 8. Dilute the solution to 1 L with water. Filter solution through a 0.22 μm filter.
Reagent Final concentration Quantity
EDTA 0.1 M 37.2 g
H2O n/a n/a
NaOH n/a n/a
Total n/a 1,000 mL
Sterile EDTA-DPBS (store at room temperature)
Reagent Final concentration Quantity
DPBS (1×) 1× 9.6 g
EDTA (0.1 M) 2.5 mM 25 mL
H2O n/a 975 mL
Total n/a 1,000 mL
Tris-HCl 0.1 M, pH 6.7 (store at room temperature)
Stir 12.11 g of Tris base in 800 mL of H2O. Add concentrated HCl under the fume hood to adjust pH to 6.7. Dilute the solution to 1 L with water.
Reagent Final concentration Quantity
Tris base 0.1 M 12.11 g
HCl (concentrated) n/a n/a
H2O n/a n/a
Total n/a 1,000 mL
Sucrose 3.3 M (store at 4 °C)
Reagent Final concentration Quantity
Sucrose 3.3 M 112.86 g
H2O n/a n/a
Total n/a 100 mL
Sorbitol 3 M (sterilise by short autoclaving; store at 4 °C)
Reagent Final concentration Quantity
Sorbitol 3 M 54.65 g
H2O n/a n/a
Total n/a 100 mL
NaCl 1 M (store at room temperature)
Reagent Final concentration Quantity
NaCl 1 M 5.84 g
H2O n/a n/a
Total n/a 100 mL
NaOH 6% (store at room temperature)
Reagent Final concentration Quantity
NaOH 6% 0.6 g
H2O n/a 10 mL
Total n/a 10 mL
Buffer A (store at 4 °C)
Reagent Final concentration Quantity
Sorbitol (3 M) 0.6 M 20 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 70 mL
Total n/a 100 mL
Buffer B (store at 4 °C)
Reagent Final concentration Quantity
Sucrose (3.3 M) 1.65 M 50 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 40 mL
Total n/a 100 mL
Buffer C (store at 4 °C)
Reagent Final concentration Quantity
Sucrose (3.3 M) 0.6 M 18 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 72 mL
Total n/a 100 mL
Buffer D (store at 4 °C)
Reagent Final concentration Quantity
Sorbitol (3 M) 0.6 M 20 mL
NaCl (1 M) 200 mM 20 mL
EDTA (0.1 M) 2 mM 2 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 48 mL
Total n/a 100 mL
Buffer E (store at 4 °C)
Reagent Final concentration Quantity
Sorbitol (3 M) 0.6 M 20 mL
EDTA (0.1 M) 5 mM 5 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 65 mL
Total n/a 100 mL
Buffer F (store at 4 °C)
Reagent Final concentration Quantity
Sorbitol (3 M) 0.6 M 20 mL
EDTA (0.1 M) 1 mM 1 mL
Tris-HCl (0.1 M, pH 6.7) 10 mM 10 mL
H2O n/a 69 mL
Total n/a 100 mL
Buffer H (store at 4 °C)
Reagent Final concentration Quantity
Sorbitol (3 M) 0.6 M 0.2 mL
n-dodecyl-β-D-maltoside 1% 10 mg
Tris-HCl (0.1 M, pH 6.7) 10 mM 0.1 mL
H2O n/a 0.7 mL
Total n/a 1 mL
RNA loading buffer
Prepare on RNase-free water and add a few crystals of bromophenol blue until the solution has a deep colour but is still transparent; store at -20 °C in 1 mL aliquots.
Reagent Final concentration Quantity
SDS (20%) 0.025% 0.0125 mL
EDTA (0.1 M) 18 mM 1.8 mL
Deionised formamide n/a 8.1875 mL
Total n/a 10 mL
RNA denaturing polyacrylamide gel
Prepare on RNase-free water. Store at 4 °C and pre-warm before use if urea precipitates.
Reagent Final concentration Quantity
TBE 10× 1× 10 mL
Urea 8 M 48 g
40% acrylamide/bis-acrylamide (19:1) 6% 15 mL
H2O n/a n/a
Total n/a 100 mL
Ammonium persulfate 10% (store at 4 °C for one month)
Reagent Final concentration Quantity
Ammonium persulfate 10% 10 g
H2O n/a n/a
Total n/a 100 mL
Pre-hybridisation buffer (store at room temperature)
Reagent Final concentration Quantity
SSC 20× 6× 300 mL
Denhardt’s solution 50× 5× 100 mL
SDS (20%) 0.2% 10 mL
H2O n/a 590 mL
Total n/a 1,000 mL
Hybridisation buffer (prepare immediately before use)
Reagent Final concentration Quantity
Pre-hybridisation buffer 1× 0.45× 9 mL
TE 10× 0.5× 1 mL
NaCl (1 M) 0.5 M 10 mL
Total n/a 20 mL
Washing buffer (store at room temperature)
Reagent Final concentration Quantity
SSC 20× 5× 250 mL
SDS (20%) 0.1% 5 mL
H2O n/a 745 mL
Total n/a 1,000 mL
Stripping buffer (store at room temperature)
Reagent Final concentration Quantity
SSC 20× 0.01× 0.5 mL
SDS (20%) 0.1% 5 mL
H2O n/a 994.5 mL
Total n/a 1,000 mL
Ethanol 80%
Reagent Final concentration Quantity
Ethanol (100%) 80% 800 mL
H2O 20% 200 mL
Total n/a 1,000 mL
RNase A dilutions in buffer D (final volume 200 μL; prepare immediately before use and keep on ice)
To prepare the working RNase A solution at 10 μg/mL, dilute 2 μL of RNase A (10 mg/mL) in 1,998 μL of buffer D and mix well.
Reagent Final concentration Quantity Buffer D volume
RNase A working solution (10 μg/mL)
0.1 μg/mL
0.2 μg/mL
0.6 μg/mL
1.2 μg/mL
2.0 μg/mL
2.6 μg/mL
3.2 μg/mL
4.0 μg/mL
6.0 μg/mL
2 μL
4 μL
12 μL
24 μL
40 μL
52 μL
64 μL
80 μL
120 μL
198 μL
196 μL
188 μL
176 μL
160 μL
148 μL
136 μL
120 μL
80 μL
Glycogen 20 μg/μL (aliquot and store at -20 °C)
Reagent Final concentration Quantity
Glycogen 20 μg/μL 200 mg
H2O n/a 10 mL
Total n/a 10 mL
Ethidium bromide 0.0001% prepared on RNase-free water (store at 4 °C)
Reagent Final concentration Quantity
Ethidium bromide 1% 0.0001% 10 μL
RNase-free water n/a 100 mL
Total n/a 100 mL
Laboratory supplies
Stericup Quick Release-GV sterile vacuum filtration system 0.22 µm pore size (Merck, catalog number: S2GVU10RE)
Yeast tRNA-derived spike-in transcript, which does not cross-map to the human genome, to enable data normalization:
5′-GAGAAGUAAGCACUGUAAAGGUUUUAGAGCUAGAAAUAGCAAGUUAAAAUAAGGCUAGUCCGUUAUCAACUUGAAAAAGUGGCACCGAGUCGGUGCUUGCCUUGUUGGCGCAAUCGGUAGCGCGUAUGACUCUUAAUCAUAAGGUUAGGGGUUCGAGCCCCCUACAGGGCUCCA-3′
Note: Depending on biological material, compatible spike-in transcript(s) should be used, with a sequence that does not cross-map with the genome of the examined organism. A spike-in transcript can be purchased or synthesised by in vitro T7 transcription (as in this case).
Amersham Hybond-N+ membrane optimised for nucleic acid transfer (Cytiva, catalog number: RPN203B)
Single-use spectrophotometer cuvette 1.6 mL Semi-micro type ClearLine (Dutscher, catalog number: 030101)
Micro Bio-Spin P-6 chromatography columns (Bio-Rad, catalog number: 7326200)
Custom DNA oligonucleotide probes for northern blot hybridisation. The oligonucleotides used for CoLoC-seq of human mitochondria are listed in Jeandard et al. (2023), Table S1.
Equipment
Micropipette PIPETMAN P2, 0.2–2 μL (Gilson, catalog number: F144054M)
Micropipette PIPETMAN P20, 2–20 μL (Gilson, catalog number: F144056M)
Micropipette PIPETMAN P200, 20–200 μL (Gilson, catalog number: F144058M)
Micropipette PIPETMAN P1000, 100–1,000 μL (Gilson, catalog number: F144059M)
Vacuum aspiration system (Integra Bioscience VACUSAFE Aspiration System, Fisher Scientific, catalog number: 11636620)
Refrigerated tabletop centrifuge for 50 mL tubes (Eppendorf 5810R, Eppendorf, catalog number: 5811000015)
Refrigerated tabletop centrifuge for 2 mL tubes (Eppendorf 5427R, Eppendorf, catalog number: 5409000010)
Refrigerated high-speed centrifuge (Avanti J-E, Beckman Coulter, catalog number: 369005) with a F0850 fixed-angle aluminium rotor (Beckman Coulter, catalog number: 364640)
Polycarbonate 50 mL bottles with screw cap for the F0850 rotor (Beckman Coulter, catalog number: 357002)
Waring two-speed blender (The Laboratory Store, catalog number: 8010EB)
Refrigerated ultracentrifuge Optima XPN-100 (Beckman Coulter, catalog number: A94469) with a swinging bucket SW 32 Ti (Beckman Coulter, catalog number: 369694)
Open-top thin-wall polypropylene tubes for a swinging bucket SW 32 Ti (Beckman Coulter, catalog number: 326823)
Water bath (e.g., VWR, catalog number: 76308-830)
Dounce homogenizer (VWR, catalog number: 432-0200)
Spectrophotometer (e.g., Eppendorf BioPhotometer 6131, Marshall Scientific, catalog number: E-BP6131)
Spectrophotometer NanoDrop (Thermo Fisher Scientific, NanoDrop 2000, catalog number: ND-2000)
Block heater (e.g., Stuart, DD Biolab, catalog number: 001150)
Gel electrophoresis chamber (e.g., BT Lab Systems, catalog number: BT206)
Low-current power supply (e.g., Consort EV233, Fisher Scientific, catalog number: 10369312)
Gel documentation system (e.g., E.A.S.Y. Doc Plus, Herolab, catalog number: 2809300)
Wet transfer tank (e.g., BT Lab Systems, catalog number: BT306)
High-current power supply (e.g., PowerPac HC, Bio-Rad, catalog number: 1645052)
UV lamp for RNA cross-linking (e.g., Hoefer UVC 500 Ultraviolet Crosslinker, Amersham Life Science, catalog number: 80-6222-50)
Rotating hybridisation oven (e.g., Problot 12 Hybridization Oven, Labnet, catalog number: H1200A-230V)
Plastic sealing machine (e.g., Manual heat sealer SK-SK 210 series, FALC Instruments, catalog number: 638.1430.20)
Phosphorimager plate (e.g., VWR, catalog number: 28-9564-75)
Exposition cassette (e.g., VWR, catalog number: 29-1755-23)
Light eraser for Phosphorimager plates (e.g., InmoClinc Screen X-ray film viewer, MedicalExpo, catalog number: 16300)
Phosphorimager scanner (e.g., GE Typhoon Trio Imager, GMI, SKU: 8149-30-0017)
Portable Geiger counter (e.g., Mini900 Ratemeter, Thermo Scientific, catalog number: MFG017)
Software and datasets
ImageQuant TL (v. 7.0, GE Healthcare)
Cutadapt (version 2.8) (available at https://pypi.org/project/cutadapt/2.8/)
READemption (version 0.4.3) (available at https://reademption.readthedocs.io/en/latest/)
Human genome sequence (Genome Reference Consortium Human Build 38 patch release 13)
READemption uses segemehl version 0.2.0-418 as the read aligner
Data analysis pipeline on Zenodo (https://doi.org/10.5281/zenodo.6389451)
Integrated Genome Browser (v. 9.1.8) (available at https://bioviz.org/)
Origin 2021b (v9.8.5.212, OriginLab Corporation) or similar nonlinear regression software
Procedure
Cell harvesting and crude fractionation
Human cell harvesting (here, we describe an example procedure for 2,500 cm2 of adherent HEK293 cells cultivated in eleven 225 cm2 flasks):
Gently aspirate the medium using a vacuum aspiration system. To each flask, add 25 mL of sterile EDTA-DPBS and incubate for 20 min at 37 °C.
Gently hit the flasks with a hand to facilitate cells detachment. Collect the cell suspension into six 50 mL tubes.
To collect the maximum of cells, rinse the flasks with 25 mL of EDTA-DPBS by transferring the solution from flask to flask. Divide the obtained cell suspension between the six half-filled 50 mL tubes from the previous step.
Pellet the cells by centrifugation at 600× g for 10 min at room temperature. Discard the supernatant.
Add 15 mL of sterile DPBS at room temperature to each tube and pool cell suspensions in two 50 mL tubes.
Pellet the cells by centrifugation at 600× g for 10 min at room temperature. Discard the supernatant.
Preparation of crude mitochondria (all procedures are carried out on ice with pre-chilled solutions and tubes):
Resuspend the cells in 30 mL of buffer A.
Disrupt the cells at high speed with a pre-chilled laboratory blender three times for 15 s, with 1 min intervals to prevent overheating. Place the homogenate into a 50 mL tube.
Remove cellular debris and nuclei by low-speed centrifugation in a refrigerated tabletop centrifuge at 1,000× g for 3 min at 4 °C. Transfer the supernatant into a new 50 mL tube.
Repeat the procedure described in A2c twice.
Note: The cell debris/nuclei pellet is easily dislodged and dispersed; one should pipette very carefully to limit this undesirable contamination.
Transfer the supernatant into polycarbonate 50 mL bottles with screw cap. Pellet crude mitochondria by centrifugation in a refrigerated high-speed centrifuge at 21,000× g for 30 min at 4 °C.
A brownish pellet will form. Carefully pipette the supernatant out.
Carefully but thoroughly resuspend the mitochondrial pellet in 8 mL of buffer A and split in four 2 mL portions.
Note: It is important to resuspend well the crude mitochondrial pellet and obtain a homogeneous suspension. However, do not overdo it to prevent the lysis of mitochondria.
Prepare four two-cushion sucrose gradients in SW 32 Ti centrifugation tubes. First, place 10 mL of buffer B in the tube. Then, carefully and slowly layer 15 mL of buffer C along the tube wall atop the first cushion using a soft, smoothly going pipette. Avoid disturbing the lower cushion.
Carefully load each 2 mL portion of crude mitochondria atop a two-cushion gradient.
Centrifuge in a refrigerated ultracentrifuge Optima XPN-100 with a swinging bucket SW 32 Ti rotor at 45,000× g for 1 h at 4 °C.
Collect the turbid mitochondria-containing interphase with a pipette.
Combine the interphases from the four tubes in a polycarbonate 50 mL bottle, dilute them with 30 mL of buffer A, and pellet the mitochondria again at 21,000× g for 30 min at 4 °C.
Carefully pipette out the supernatant.
Gently but thoroughly resuspend the mitochondrial pellet in 800 μL of buffer A. Critical: The suspension of mitochondria should be homogeneous. Save 500 μL of the suspension for a CoLoC-seq experiment (section B) and reserve the other 300 μL for a Mock CoLoC-seq experiment (section D). Keep them on ice.
CoLoC procedure
All procedures are carried out at 4 °C with pre-chilled solutions and tubes, unless specified differently; all centrifugations are performed in a refrigerated tabletop centrifuge for 2 mL tubes.
Measure protein concentration in the sample for the CoLoC-seq experiment by Bradford assay.
Note: The spectrophotometer should be calibrated for the Bradford assay, following the manufacturer’s instructions. To this end, one typically uses a series of bovine serum albumin solutions containing 1–10 μg of protein.
Dissolve 2 μL of the mitochondrial suspension in 40 μL of 6% NaOH. For the blank, use 2 μL of buffer A.
Add 760 μL of MilliQ water and mix thoroughly.
Add 200 μL of the Bradford solution, mix well, and transfer the suspension to a disposable plastic spectrophotometer cuvette.
After 60 s of incubation at room temperature, read the blank OD at 595 nm. Then read the mitochondrial protein sample OD at the same wavelength and determine its protein concentration using the standard curve. From 2,500 cm2 of nearly confluent HEK293 cells, one typically obtains 5–10 mg of crude mitochondria.
Adjust the mitochondrial suspension to 1.6 mg of protein per millilitre using buffer A.
Split the resulting suspension of mitochondria in a series of identical 80 μL samples. Ten samples must be sufficient to create an informative RNA depletion curve.
Prepare two series of 80 μL RNase A dilutions in buffer D with concentrations ranging from 0 to 6 μg/mL (e.g., 0, 0.1, 0.2, 0.6, 1.2, 2.0, 2.6, 3.2, 4.0, and 6.0 μg/mL). Use one series for the CoLoC and the other for the Mock CoLoC procedure.
Pre-warm the RNase A dilutions on water bath at 25 °C for 1 min.
Mix the 80 μL mitochondrial samples with the corresponding 80 μL RNase A dilutions and incubate on water bath at 25 °C for 10 min.
Dilute the reactions with ice-cold buffer E to 1.6 mL.
Centrifuge at 16,000× g for 20 min at 4 °C.
Resuspend the pellets thoroughly in 100 μL of buffer F.
RNA extraction
Caution: RNA extraction should be performed in a fume hood and with personal protection.
Add 1 mL of TRIzol and shake samples for 10 s (do not vortex). Pause point: Once TRIzol is added to the samples, they can be stored at -80 °C; otherwise, proceed to step C2.
Note: TRIzol is stored at 2–25 °C; however, it should be warmed up to room temperature before use.
Incubate samples for 5 min at room temperature.
Add 0.2 mL of chloroform and cap the tubes securely.
Shake the tubes vigorously by hand for 15 s.
Incubate for 3 min at room temperature.
Centrifuge the samples at 14,000× g for 15 min at 4 °C.
Carefully transfer the upper aqueous phase containing RNA to a new tube by angling the tube at ~45° and pipetting the solution out with a P200 micropipette. Critical: Take up the aqueous phase as completely as possible but avoid touching the protein-containing interphase. Contamination of RNA samples with protein may result in low-quality and/or artefactually degraded RNA.
Add 2 μL of 20 μg/μL glycogen to the samples to facilitate RNA precipitation.
Add 0.5 mL of isopropanol to the samples and mix thoroughly.
Let the RNA precipitate at -20 °C for 1 h. Pause point: The samples can be stored at -80 °C.
Centrifuge at 14,000× g for 20 min at 4 °C. Carefully decant the supernatant into a clean tube.
Note: RNA pellets sometimes do not stick well to the tube wall. Decanting the supernatant to another tube prevents the pellet loss; if needed, simply transfer the supernatant with the floating pellet back and repeat the centrifugation.
Wash the pellet with 0.5 mL of 80% ethanol.
Centrifuge at 14,000× g for 10 min at 4 °C. Decant the supernatant as previously.
Wash the pellet with 0.5 mL of 100% ethanol.
Centrifuge at 14,000× g for 10 min at 4 °C.
Decant the supernatant, as previously. Dry the pellets by leaving the tubes open on the bench for 5–10 min.
Fully dissolve RNA in 45 μL of RNase-free water. Pause point: Samples of isolated RNA can be stored at -80 °C.
Mix each sample of extracted RNA with 90 ng of a spike-in transcript to enable subsequent data normalization.
Add to each sample 5 μL of the 10× DNase I reaction buffer, 1 U of DNase I, and 20 U of SUPERase•In. Incubate for 30 min at 37 °C to digest residual DNA.
Re-extract RNA with TRIzol, as described in steps C1–C7. Precipitate it, as in steps C9–C16.
Note: No further glycogen addition is required, since the previously added glycogen partitions with RNA during extraction. Re-dissolve the RNA pellet in 20 μL of RNase-free water.
Measure RNA concentration in each sample with NanoDrop. Expected RNA concentrations are in the range of 50–200 ng/μL. Pause point: RNA samples can be stored at -80 °C.
Mock CoLoC procedure
Mix 300 μL of the reserved crude mitochondrial extract from step A2n with 300 μL of ice-cold buffer H.
Lyse the mitochondria on ice with 50 strokes in a small Dounce homogenizer.
Clear the lysate by centrifugation at 16,000× g for 20 min at 4 °C. Transfer the supernatant to a new tube.
Note: The remaining pellet must be relatively small, and the supernatant must be of yellow-to-brown colour, indicating good solubilisation of the mitochondrial contents. If the pellet is still big and the supernatant is colourless, repeat the lysis adding more buffer H.
Repeat step D3 to remove any remaining insoluble material.
Measure protein concentration in the cleared lysate by Bradford assay, as described in step B1.
Adjust the lysate to protein concentration of 1.6 mg/mL with ice-cold buffer A.
Split the resulting solution in a series of identical 80 μL samples in 2 mL tubes.
Perform RNase treatment in the same way as for the regular CoLoC procedure (steps B4–B6).
Centrifuge the reactions at 16,000× g for 40 min at 4 °C.
Add 1.44 mL of TRIzol. Pause point: The samples can be stored at -80 °C. Otherwise, proceed to step D11.
Incubate the samples at room temperature for 5 min. Add 320 μL of chloroform and extract RNA, as described in steps C4–C21.
Northern blotting
Caution: Work with radioactive material in a specially equipped lab using personal protection.
Note: While the northern blotting protocol described here is based on radioactive probes, viable non-radioactive alternatives also exist, e.g., irNorthern, using near-infrared fluorescence (Miller et al., 2018), and digoxigenin-labelled probes, exploiting western-like chemiluminescence detection (Höltke et al., 1992).
Take 1 μg of the 0 RNA sample (treated with 0 μg/mL RNase A). Take the same volume of all remaining samples in a series. (For example, if the 0 sample has the RNA concentration of 200 ng/μL, you should take 5 μL of each sample in this series, independently of the RNA concentration of the other samples.) (Note: RNA yield naturally decreases in samples treated with higher RNase A concentrations. Keeping the same volume fraction enables correct dosage for all samples of a series. Measuring the level of the spike-in RNA will further ensure equal sample loading.) Mix the RNA 1:1 (by volume) with RNA loading buffer. Caution: The RNA loading buffer contains formamide and should be manipulated under the hood.
Prepare a big (15–20 cm) and thick (1.5 mm) 6% RNA denaturing polyacrylamide gel by polymerising the gel solution by addition of 1/100 volume of 10% ammonium persulfate and 1/1,000 volume of TEMED.
Denature the samples at 95 °C for 5 min in a block heater.
Install the gel slot in the electrophoresis chamber and add a sufficient amount of 1× TBE buffer.
Just before loading the samples, thoroughly wash the pockets from urea 10 times by vigorous pipetting with a P1000 pipette.
Keep the samples on the bench for 1 min and load them immediately after the last wash of the pockets on the gel using a smoothly going pipette with a long thin tip.
Note: It is important to load the sample firmly and quickly on the very bottom of the well, avoiding spurts and bubbles.
Close the chamber and run the gel at 10 V/cm and 30 mA until bromophenol blue is approximately 3 cm from the bottom.
Carefully disassemble the sandwich. Stain the gel in 200 mL of 0.0001% ethidium bromide for 5 min. Visualise the ethidium fluorescence and record the resulting image with the help of a suitable UV-based gel documentation system.
Note: Minimise exposure of the gel to UV light to avoid RNA crosslinking.
Assemble a northern blot sandwich according to manufacturer’s instructions for a wet transfer tank. For example, use three sheets of Whatman paper with dimensions matching those of the gel. Moisten them with 0.5× TBE and apply them one upon the other, atop the sponge on the anode (+) side of the cassette. After each sheet, use a roller to gently chase air bubbles from the sandwich. Then, apply a Hybond-N+ membrane for nucleic acids transfer (moistened in 0.5× TBE) atop the Whatman sheets. Then, carefully place the gel on the membrane and cover it with three more moistened Whatman sheets. Close the sandwich.
Place the cassette in the corresponding wet transfer chamber filled with cold, freshly prepared TBE 0.5× buffer and perform transfer at 75 V and up to 3 A for 1.5 h in a cold room, with gentle magnet stirring.
Note: While 1.5 h is generally enough for a complete transfer, this step can be prolonged up to 3 h, at the experimenter’s convenience.
Disassemble the sandwich and air dry the membrane on a sheet of absorbing paper. Crosslink RNA to the membrane for 3 min in Hoefer UVC 500 Ultraviolet Crosslinker. Pause point: The dry membrane can be stored at 4 °C.
Place the membrane in a hybridisation glass tube, add 20 mL of pre-hybridisation buffer, and incubate in a slowly rotating hybridisation oven for 30 min at 65 °C. Make sure that the membrane is uniformly unfolded in the tube and does not fold back on itself (if this happens, simply invert the tube), and that all areas subject to hybridisation are in maximally identical conditions and uniformly covered with the liquid.
Replace the pre-hybridisation buffer with the hybridisation buffer and add 3–5 μL of the purified labelled probe.
Note: For 5′-[32P]-oligonucleotide labelling, see section F.
Incubate the membrane with the probe for at least 2 h (normally, over day or overnight) at 42 °C with continuous rotation in the hybridisation oven.
Note: If the probed transcript is of low abundance, the amount of the probe can be increased. If the probe is long or highly structured, it should be denatured and put on ice, as described in step F1, prior to its addition to hybridisation buffer. If the probe is very long (> 40 nt), it is recommended to increase the incubation temperature to 50–60 °C to avoid non-specific hybridisation. The optimal hybridisation temperature can be adjusted for each oligo individually, with 42 °C performing well for most probes.
To wash the blot, discard the hybridisation mix to a dedicated radioactive waste container, add 20 mL of the washing buffer, and rotate the tube in the hybridisation oven for 15 min at the same temperature. Caution: Do not discard radioactive solutions in the sink.
Discard the washing buffer in a dedicated radioactive waste container. Remove the membrane from the glass tube.
Dry the membrane on a sheet of absorbing paper on the bench. Seal it in a plastic film.
Expose the membrane overnight (or longer, up to a week, if the signal is expected to be weak) with a phosphorimager plate in an appropriate cassette. Shorter expositions are only needed if the signal is very strong (measurable with a portable counter) and becomes quickly saturated.
Scan the phosphorimager plate on Typhoon or a similar device using appropriate instrument settings.
Erase the residual signal from the plate with a light eraser.
Note: The phosphorimager plate bleaching is also recommended immediately before exposition (step E18) to reduce background.
To remove the previous probe for further hybridisations, rotate the membrane in a glass tube in the hybridisation oven in 20–30 mL of stripping buffer for 30 min at 63–65 °C. If the probe is very long and/or the signal is strong, remove it by rocking the membrane with an excess of stripping buffer in a water bath at 80 °C for 30 min. Then, proceed with the next hybridisation from step E13.
Oligonucleotide labelling
Caution: Work with radioactive material in a specially equipped lab using personal protection.
Prepare a 20 μL labelling reaction by mixing 13 μL of water, 1 μL of a 10 μM solution of the DNA oligonucleotide to label, 2 μL of 10× PNK buffer, 3 μL of [32P]-γ-ATP, and 1 μL of PNK.
Note: If the probed transcript is very abundant, one can take 1 μL of [32P]-γ-ATP and 15 μL of water instead (the rest of the mix will be the same). If the probe is very long (> 40 nt) or highly structured, mix 13 μL of water and 1 μL of a 10 μM solution of the DNA oligo and denature it at 95 °C for 1 min, then immediately put on ice for 1 min and quickly add the remaining components of the labelling mix, as described above. This usually increases labelling efficiency.
Incubate the reaction mixture at 37 °C for 30 min to 1 h. Caution: Shield the heating block with a Plexiglas screen to protect yourself and others from exposure to radioactivity.
Oligonucleotide purification:
Briefly vortex a Micro Bio-spin P-6 chromatography column.
Shake the drops down, break the bottom plug off, and insert the column in the corresponding collecting tube.
Open the upper lid, close it again to push air bubbles through, and open once again.
Let the liquid in the column drop out by gravity flow for 3–4 min on the bench. Discard the flow-through.
Centrifuge the column with the collecting tube at 1,000× g in a tabletop centrifuge at room temperature for 2 min.
Discard the collecting tube.
Carefully, without touching the resin, apply the labelling solution to the column and insert it in a clean 1.5 mL collection tube.
Centrifuge at 1,000× g in a tabletop centrifuge at room temperature for 4 min. One should normally obtain approximately 20 μL of clean labelled probe. Critical: Depending on the supplier, [32P]-γ-ATP solutions sometimes contain a colorant to facilitate their tracking. This one should stay on the column. In the case where the eluate is still coloured, repeat the procedure with another column.
Measure the probe solution with a portable counter: it should normally be over the scale and beep.
Note: Store the labelled oligonucleotide at -20 °C in a dedicated freezer in the radioactivity lab. Before each use, verify with a portable counter that the probe is still sufficiently radioactive.
Library preparation & RNA-seq
Note: cDNA library preparation and RNA-seq can be outsourced. In Jeandard et al. (2023), Figure S2 shows the key library preparation steps that ensure the selective sequencing of intact transcripts, as briefly summarised below.
Remove caps with RNA 5′-pyrophosphohydrolase.
Ligate the 5′-adapter to 5′-phosphorylated ends.
Ligate the 3′-adapter to 3′-hydroxyl ends.
Perform the first-strand cDNA synthesis using M-MLV reverse transcriptase with a 3′-adapter-annealing primer.
PCR-amplify the resulting cDNA with a high-fidelity DNA polymerase and barcoded TruSeq primers (15 cycles).
Purify cDNA with the AMPure XP kit.
Perform cDNA fragmentation and end repairing and proceed with another round of adapter ligation and PCR amplification.
Pool the cDNA samples equimolarly and perform size-selection on an agarose gel in the range 10–220 nt (excluding the flanking sequences).
Sequence the pool on an Illumina NextSeq 500 instrument (75 nt single-end reads) or similar.
Note: Although the fragmentation step may be expected to destroy the strand-specificity of the protocol, our mapping results (Jeandard et al., 2023) showed that the first 5′-adapter ligation (step G2) largely determines the strandedness of the reads, which permits unambiguous transcript assignment and quantification. Other strategies to enforce the preservation of the strand information can also be implemented (J. Z. Levin et al., 2010; Dar et al., 2016).
Data analysis
The first important information about the success of a CoLoC experiment comes from the analysis of ethidium bromide–stained gels and northern blots. Results of these steps help to evaluate the quality of the RNA samples and make a decision on their suitability for subsequent library preparation (Figure 2). The northern blot signal from the full-size transcript in each sample is analysed by ImageQuant TL or similar densitometry software and normalised by the signal of the spike-in RNA in the same sample. This enables direct measurement of intact transcript levels across the gradient of RNase A concentrations. A well-behaved, informative CoLoC/Mock CoLoC series:
contains at least 9–10 samples with no signs of artefactual degradation (unspecific, irregular RNA degradation is sometimes visible as the appearance of a smear or random bands),
shows a robust signal for the full-size transcript at least in the 0 sample (treated with 0 μg/mL RNase A), which permits reliably measuring the starting level of the RNA of interest,
has a monotonously decreasing pattern for at least some of the probed full-size transcripts, as the RNase concentration increases, indicating that the selected RNase concentration range provides sufficient activity and resolution to plot an informative digestion kinetics.
Note: We strongly recommend probing for a wider variety of transcripts differing in size (50–1,500 nt), structure (loosely structured vs. tightly folded), and localisation, including both bona fide resident RNAs and a few abundant contaminants. If their behaviour corresponds to the expectations of the CoLoC model (gradual disappearance for contaminants; plateauing for residents), it is usually a good indication that such samples can be used for a genome-wide analysis by RNA-seq.
Figure 2. Evaluating the quality of CoLoC samples from ethidium bromide staining and northern blot data. A. This ethidium bromide–stained gel shows an example of a well-behaved series of CoLoC samples. Addition of RNase A provokes gradual bulk RNA degradation, which gets more pronounced as the RNase concentration increases. B. This ethidium bromide–stained gel provides an example of a failed experiment: addition of RNase A has little-to-no effect on bulk RNA; the RNase activity turned out to be insufficient to create meaningful digestion dynamics. C. These northern blots illustrate two successful outcomes of a CoLoC experiment. The upper panel shows an RNase-resistant transcript: its level remains relatively unchanged across the entire gradient of RNase concentrations. The lower panel features an RNase-sensitive RNA, progressively digested with increasing RNase concentrations. D. These northern blots show two examples of less successful experiments. In the upper panel, the transcript level evolves in a fuzzy, indeed random pattern, preventing the fitting of any reasonable kinetics model. In the lower panel, the signal in the 0 sample is too low and cannot be measured with confidence. Therefore, one cannot determine the starting level of this RNA and reliably scale its profile.
Important steps of RNA-seq data treatment are described in Materials and Methods of Jeandard et al. (2023). Briefly, the sequencing reads were pre-processed with cutadapt version 2.8 (Martin, 2011) to trim adapter sequences. Read alignment and gene feature quantification were done with READemption version 0.4.3 (Förstner et al., 2014), using segemehl version 0.2.0-418 (Otto et al., 2014) as the read aligner. All libraries were aligned to the Human genome (Genome Reference Consortium Human Build 38 patch release 13) retrieved from RefSeq (O’Leary et al., 2016). The parameters used for alignment, coverage calculation, and feature quantification can be found in the scripts deposited at Zenodo (https://doi.org/10.5281/zenodo.6389451). Entries for repetitive genes were compounded and their reads were summed up. A cut-off of 30 reads was applied to all 0 samples to ensure reliable initial level measurement. Read counts in each library were normalised by the corresponding number of reads mapping to the spike-in RNA, as described for northern blotting. Read distributions can be visualised in Integrated Genome Browser (v. 9.1.8), Integrative Genomics Viewer, or other similar software. Due to the specifics of the library preparation strategy, the reads typically cluster at the 5′ ends of transcripts. By contrast, if we saw that the overwhelming majority of reads for a certain mRNA- or lncRNA-encoding locus aligned to embedded tRNA-, snRNA-, 7SL-, 5S rRNA-, or mtDNA-like (NUMTs) sequences (usually found in introns), we excluded such genes from the analysis as obvious cross-mapping artefacts.
The quantitative data obtained from northern blots or RNA-seq permits deducing the rate at which RNase A digests the transcript and the relative size of the unreactive pool, protected from the nuclease, using a kinetics model described in detail in Jeandard et al. (2023) in Materials and Methods:
f(A)=(1-P0) e-k'iA+P0 (1)
where f(A) is the relative proportion of the ith transcript remaining after treatment with A μg/mL of RNase A, P0 is the initial protected proportion of the ith transcript (i.e., the part of the ith transcript pool that is unavailable to RNase A, as measured at 0 μg/mL of RNase A), and k’i is the effective digestion rate constant for the ith transcript. Because f(A) is expressed in relative units, the starting transcript level (0 sample) must be normalised to 1, and the remaining samples of the series must be scaled accordingly. To fit the reaction model into the data, we used a customised nonlinear regression function in Origin 2021b, but any other similar software can be used too. Caution: Do not linearize the Equation 1 with the intention to use linear regression instead! Such an approach is misleading as it results in incorrect error modelling, leading to overoptimistic or, on the contrary, strangely poor output statistics. Nonlinear regression software is now widely accessible and quite intuitive. A good general primer in nonlinear regression and associated topics can be found in Motulsky (2010).
Begin by trying to fit the full model (called Model 1 in Jeandard et al., 2023) that has two parameters: the effective digestion rate constant k’ and the initial proportion of RNA protected from digestion P0. It is useful to naturally constrain k’ to be non-negative (k’ ≥ 0) and P0 to be in the range [0,1]. If the software has such an option, it is also recommended to specify as the initial P0 estimate the lowest measured level of the transcript (typically observed in the sample, treated with the highest RNase concentration). This considerably accelerates fitting and increases chances that the fit will converge on a biochemically meaningful combination of parameters reflecting the actual state of the system.
Several regression outcomes are possible (Figure 3). In a successful experiment, the large majority of transcripts yield converging fits, i.e., a certain pre-specified level of the sum of squares of residuals has been reached. This means that k’ and P0 can be reliably estimated for most kinetics (Figure 3A). If this is not the case, the regression software reports a non-converging fit (Figure 3B), which might be either due to low quality of the data (points are too much scattered and do not form any obvious pattern) or because the Model 1 is too complex. In the first case, the data for this specific transcript are, unfortunately, unusable. In the second case, fitting a simpler, nested model may rescue the analysis (see the discussion of this model below):
f(A)=e-k'iA (2)
One should pay attention to—and correctly interpret—the three groups of statistics associated with each fit. The first one is determination coefficient R2. When R2 = 1, the fit is perfect: the model goes through every data point. When R2 = 0, the model is actually a straight horizontal line. (It is also possible in nonlinear regression for R2 to be negative, meaning that the model fits data more poorly than the horizontal line. Such cases are exceptionally rare in CoLoC-seq. See, for example, Figure 3B.) Caution: It is a common misconception to conclude that low R2 means poor model fit! The quality of fitting can only be evaluated by looking at the residuals, and if the regression software says that the fit converged, it means that the residuals are small enough. Therefore, the fit can actually converge on a horizontal line (R2 = 0) as the best model fitting the data (Figure 3C). This case is typical for resident or RNase-resistant transcripts: their level is unaffected by increasing RNase concentration and, within the limits of the random error, remains constant (i.e., the graph is a horizontal line).
Figure 3. Real-life examples of CoLoC-seq data and their analysis by nonlinear regression, performed in Origin 2021b. A. Well-behaved series showing a typical RNase-sensitive transcript. P0 is nearly 0 and insignificant; k’ is high and significant. Dependency is low, indicating that P0 and k’ have been estimated independently from each other; R2 is high; the fit converged. B. Example of a failed series: the data points are too much scattered; as a result, the fit did not converge. C. This successful series shows a typical behaviour of a transcript protected from RNase: with the exception of a couple of outliers, the data points align close to the horizontal line with P0 ≈ 1. k’ is insignificant, indicating that no appreciable degradation of this transcript occurred. R2 is nearly 0 (i.e., horizontal line), but the fit converged well. D. This well-behaved series shows a transcript that is partially protected from RNase. Both P0 and k’ are significantly non-zero, indicating that the population of this RNA was digested to some extent and plateaued at the level of 0.224 (i.e., 22.4% of this transcript pool is estimated to be protected from RNase). E. Fitting the Model 1 (solid line) into this series resulted in highly entangled and unreliable P0 and k’ estimates (dependency 0.729), suggesting that the two variables are significantly collinear, and the model is unnecessarily complex. Fitting the simpler Model 2 (which does not have the P0 parameter; dashed line) permitted a more reliable estimation of k’. Note that the two models yield almost indistinguishable fits; however, the Model 2 is more parsimonious, which explains its success in estimating k’. See Jeandard et al. (2023), Table S2 for further examples.
The second group of statistics concerns the best-fit values of the parameters and the associated uncertainty measures. The parameter of primary interest is P0. When P0 ≈ 0, there is no sizable protected pool for the ith RNA, meaning that it fully participates in the reaction (Figure 3A). Therefore, under the CoLoC-seq setup, such an RNA is a regular, degradable contaminant. We operationally consider that all transcripts with P0 < 0.1 are potential contaminants (see Figure 3C in Jeandard et al., 2023). By contrast, when P0 is considerably larger (P0 > 0.1, Figure 3D), a significant proportion (> 10%) of the ith RNA does not participate in the reaction, i.e., it is somehow protected from RNase A (with P0 = 1 meaning that 100% of the transcript is protected). How exactly protected depends on the results of CoLoC-seq vs. Mock CoLoC-seq experiments. If the transcript shows high P0 in CoLoC-seq but not in Mock CoLoC-seq, where the organellar membranes have been dissolved, one can consider this as evidence that a pool of this transcript genuinely resides inside the organelle, and its protection was due to the membranes. By contrast, if both CoLoC-seq and Mock CoLoC-seq return high P0 values, this means that the transcript in question is intrinsically resistant to RNase A, and its protection has nothing to do with the organellar membranes. As discussed on several examples in Jeandard et al. (2023), such transcripts are most likely false positives.
The best-fit value of k’ may also be of interest. For example, when k’ ≈ 0 (Figure 3C), it means that the transcript in question practically does not undergo degradation, and the initial Model 1 degenerates to the simplistic Model 3, i.e., a horizontal line:
f(A) = 1 (3)
The best-fit values should be interpreted along with the accompanying uncertainty statistics. These are often returned as standard errors (SE) or, more conveniently, confidence intervals (CI). Narrow SE or CI mean that the corresponding parameters (P0 or k’) are estimated with precision. Importantly, when the CI for P0 includes 0, it means that P0 is not significantly different from 0, i.e., there is no strong evidence for the existence of a protected pool for this transcript. When the CI for k’ includes 0, it means that the digestion rate is insignificant, that there is no strong evidence that RNase A actually cleaves this RNA (i.e., the transcript is protected). Under the CoLoC-seq setup, regular, degradable contaminants typically show low P0 values, with CIs including 0, and relatively high k’ values, with CIs far from 0 (Figure 3A). By contrast, protected transcripts feature relatively high P0 values, with CIs far from 0; their k’ values can vary, depending on the proportion of the ith RNA available for degradation (Figure 3C and 3D).
The best-fit values of parameters are often accompanied by P-values conveying similar information (the null hypothesis being that the true value of the parameter is 0). A low P-value suggests that the parameter is likely non-zero. Under the CoLoC-seq setup, degradable contaminants often show high P-values for P0 and low P-values for k’ (Figure 3A). By contrast, resident transcripts should normally have low P-values for P0 (Figure 3C, D). A high P-value (and a CI including 0) means that the corresponding parameter is not useful, i.e., the complete Model 1 can be simplified to the nested Model 2 (simple pseudo-first order decay) or the Model 3 (no decay).
The last part of regression statistics, which is very important to take in consideration, is the dependency between parameters. Dependency is the degree of entanglement between the estimated parameters. In some situations, especially where the RNase-mediated digestion is very slow or even inexistant, P0 and k’ become significantly collinear, and the regression software struggles to fit them separately since small changes in either of them yield nearly equivalent results (Figure 3E). In Jeandard et al. (2023), we arbitrarily used the dependency cutoff of 0.3, above which we considered P0 and k’ to be too much entangled to speak about their independent fitting. This threshold can be reviewed when applying CoLoC-seq to other systems. Just like the uncertainty parameters discussed above, high dependency means that the Model 1 is unnecessarily complex, and one of the parameters must be dropped.
Since the Model 3 is always implicitly tested by the regression software (see the discussion of R2 above), one can only omit P0 from the model, thus yielding the Model 2. Therefore, if by using the complete Model 1 the fit does not converge, or one of the estimated parameters is not significant, or there is strong interdependency between the parameters, one should try to fit the simpler Model 2 into the data. This usually helps to rescue for analysis the profiles of the majority of remaining transcripts and especially those with a high level of protection from RNase A (i.e., the transcripts of highest interest for CoLoC-seq). The Model 2 has only one explicit parameter, k’. Interpretation of the results of this regression follows the same reasoning as for the Model 1, with high significant k’ corresponding to degradable transcripts, while low insignificant k’ means protection (one can assign such transcripts a nominal P0 of 1). The decision making is summarized in Table 1.
We recommend using at least two biological replicates for CoLoC-seq and Mock CoLoC-seq. If the replicates are very similar to each other, it makes sense to combine them for model fitting and thereby increase the power and precision of the regression, while preserving the information about biological variability of the original samples. The similarity of the replicates can be evaluated at several levels: (i) by plotting read counts for the same transcript from different replicates at the same RNase concentration, or (ii) by fitting the Model 1 separately for each replicate and plotting together the P0 values from different replicates (see for an example Figure S6 in Jeandard et al., 2023). The latter test is more stringent.
Validation of protocol
We evaluated the feasibility and performance of CoLoC-seq on the well-studied mitochondrial transcriptome of human embryonic kidney 293 (HEK293) cells (Jeandard et al., 2023). To this end, two biological replicate series of CoLoC-seq and Mock CoLoC-seq samples were analysed by northern blotting and RNA-seq. The replicates were very similar to each other and yielded highly correlated P0 values (Figure S6 in Jeandard et al., 2023), confirming the intra-method reproducibility. The RNA levels and the fitted P0 values obtained from RNA-seq and northern blotting measurements also showed excellent agreement (Figure 2 and Figure S5 in Jeandard et al., 2023), indicating that CoLoC-seq correctly captures quantitative information about the kinetic behaviour of analysed transcripts. No significant bias at the level of transcript abundance, sequence, or structure was observed (Figure S7 in Jeandard et al., 2023). However, due to limitations of the standard RNA-seq protocol, the representation of tRNAs in the RNA-seq libraries was generally biased against extensively modified species (Figure S8 in Jeandard et al., 2023).
The behaviour of transcripts in the CoLoC-seq and Mock CoLoC-seq setups was significantly different (Figures 3 and 4 and Figure S6 in Jeandard et al., 2023), indicating that the mitochondrial membranes do provide shelter for a subset of RNA molecules. As expected, the mitochondrial DNA-encoded mitochondrial RNAs were mostly resistant to RNase A in the CoLoC-seq setup but rapidly degraded in Mock CoLoC-seq, confirming that they are genuinely present inside the organelles. By contrast, the vast majority of nuclear DNA-encoded transcripts, such as the abundant 5.8S rRNA and U6 snRNA, were rapidly degraded in both cases, confirming that they are surface-attached degradable contaminants (Figures 2–4 in Jeandard et al., 2023). Of note, a few short, highly structured, and protein-bound noncoding transcripts, such as 5S rRNA and the RNA components of RNases P and MRP, plateaued at an intermediate level in both the CoLoC-seq and the Mock CoLoC-seq experiments. This indicates that they remained, to a large extent, resistant to RNase degradation even when the mitochondrial membranes had been dissolved and may, therefore, represent recalcitrant contaminants. We also identified a few RNA Pol III transcripts, such as Y RNAs, SNAR-A, and tRNAs, as likely partially mitochondria-localised (Figure 4 and Figure S8 in Jeandard et al., 2023), which was further corroborated by biochemical and smFISH assays (Figure 5 in Jeandard et al., 2023).
Table 1. Decision making criteria to interpret CoLoC-seq regression data
Model 1 (complete)
CoLoC-seq Mock CoLoC-seq Interpretation
P0 P0 CI includes 0? P0 P-value k’ k’ CI includes 0? k’P-value P0 P0 CI includes 0? P0 P-value k’ k’ CI includes 0? k’P-value
≈0 Usually yes* Usually high* High No Low ≈0 Usually yes* Usually high* High No Low Degradable contaminant
Far from 0 but < 1 No Low High No Low ≈0 Usually yes* Usually high* High No Low Partially resident inside the organelle
≈1 No Low ≈0 Yes High ≈0 Usually yes*
Usually
high*
High No Low Fully resident inside the organelle
Far from 0 but < 1 No Low High No Low Far from 0 but < 1 No Low High No Low Partially RNase-resistant transcript
≈1 No Low ≈0 Yes High ≈1 No Low ≈0 Yes High RNase-resistant transcript
Model 2 (pseudo-first order decay)
k’ k’ CI includes 0? k’P-value k’ k’ CI includes 0? k’P-value Interpretation
High No Low High No Low Degradable contaminant
≈0 Yes High High No Low Fully resident inside the organelle
≈0 Yes High ≈0 Yes High RNase-resistant transcript
*The CI width and the P-value largely depend on the sample size, and at a high enough n it is quite common to obtain statistically significant results even for very small P0 values (close to 0). In such situations, one should rely more on the absolute value of P0: if it is very small, it is reasonable to consider such a transcript as fully degradable (i.e., contaminant, under the CoLoC-seq setup), even if the uncertainty measures seem to be "highly significant". In Jeandard et al. (2023), given the limited precision of RNA level measurements by northern blotting and by RNA-seq, we arbitrarily chose the P0 cut-off of 0.1. This means that at least 10% of the transcript needs to be excluded from the reaction to speak about a significant level of protection. See also the discussion of unusually small yet significant k’ values in Table S2 in Jeandard et al. (2023).
Since the mitochondrial transcriptome of human cells has become a popular benchmark for subcellular transcriptomics (Mercer et al., 2011; Kaewsapsak et al., 2017; Fazal et al., 2019; P. Wang et al., 2019; Zhou et al., 2019), we could directly compare the CoLoC-seq performance with that of other genome-wide methods (especially, the most robust proximity labelling techniques). We found that CoLoC-seq performed equally well on long transcripts (rRNAs, mRNAs, lncRNAs) and significantly outperformed alternative methods on shorter transcripts (tRNAs, snRNAs, snoRNAs, scaRNAs etc.), which are generally poorly covered by proximity labelling approaches.
General notes and troubleshooting
General notes
One should take into account two important points when choosing the RNase for CoLoC-seq experiments: (i) it should behave as a kinetically perfect enzyme, i.e., its catalysis must be limited only by diffusion (Park and Raines, 2003) and (ii) it should produce 5′-hydroxyl and 2′- or 3′-phosphate termini. The first property enables a straightforward use of the CoLoC-seq kinetics model, which implies that diverse transcripts in the sample are cleaved independently, without significant RNase-sequestration effects; therefore, the RNase concentration can be considered constant, and the entire reaction becomes pseudo-first order. The second property permits selective sequencing of intact transcripts (as required by the CoLoC-seq model, which only looks at remaining intact RNA; see Eq. 1): a single cleavage by such an RNase generates a terminus incompatible with standard adaptor ligation (see the section G. Library preparation & RNA-seq). (The caveat here is that one cannot study transcripts with natural 5′-OH or 2′- or 3′-phosphorylated ends by RNA-seq. However, they remain analysable by northern blotting.) Other highly active RNases generating 5′-hydroxyl and 2′- or 3′-phosphate termini (micrococcal nuclease, RNase I, RNase T1) could in principle be used too. However, we and others found them to be overall less well performing and more idiosyncratic than RNase A [Yang, 2011; Aryani and Denecke, 2015; Jeandard et al., 2023 (Figure S1)].
The exact RNase concentrations used in CoLoC-seq depend on the specific activity of the enzyme batch and the nature of the studied biological material. They should be adjusted individually for every new application. It is essential that the selected concentration range enables the observation of gradual digestion dynamics of contaminant transcripts without compromising the quality of the final RNA samples (Figure 2A, 2B). One should strive to get particularly high resolution at low RNase concentrations, where a small change in RNase typically results in a big change in the remaining transcript level (Figure 3A, 3D). This facilitates k’ fitting and thereby increases the precision of the P0 estimate.
CoLoC-seq can be applied to nearly any membrane-bounded organelles (see, for example, the previously published isolation protocols for chloroplasts and apicoplasts: Kunst, 1998; Botté et al., 2018) and other entities known or suspected to contain RNA (viruses, endosymbiotic organisms, extracellular vesicles). Since extracellular RNA can be packaged in and protected by membranous vesicles, such as exosomes, but also by free RNPs, researchers that wish to adapt CoLoC-seq to such systems may need to incorporate a proteinase K pre-treatment step to expose RNP-embedded contaminants and enable their subsequent digestion by RNase (Arroyo et al., 2011; Hill et al., 2013; Mateescu et al., 2017; Jeppesen et al., 2019; Murillo et al., 2019; Gruner and McManus, 2021).
Finally, due to intrinsically low susceptibility of miRNAs to RNase-mediated degradation (especially when they are associated with proteins, as it is typically the case), we discourage using CoLoC-seq (or indeed other RNase-based approaches) to infer their localisation topology (Arroyo et al., 2011; Aryani and Denecke, 2015).
Troubleshooting (Table 2)
Table 2. Troubleshooting
Problem Possible cause Solutions
Presence of unspecific RNA degradation in samples (smear, additional random bands)
• RNase contamination in water.
• RNase contamination from air.
• RNase contamination of pipette filters.
• Prepare all solutions on RNase-free water.
• Work in a dedicated RNase-free environment.
• Change pipette filters.
Low RNA yield
• Insufficient starting material.
• Poor organelle yield.
• Increase the amount of the starting material.
• Try an alternative organelle isolation protocol with a higher yield.
Uneven pattern across samples upon ethidium bromide staining or northern blotting (e.g., Figure 2D)
• Organelle pellet was unevenly or insufficiently resuspended between samples.
• Non-uniform RNA isolation.
• Insufficiently solubilised RNA pellets.
• Resuspend the organelle pellet carefully but thoroughly before splitting it into the sample series. Use a narrower pipette tip, if required.
• Make sure that the aqueous phase is taken up uniformly between samples. Always process samples in the same order to make their treatment maximally identical.
• If you suspect that the RNA pellet is not fully dissolved (often due to protein contamination), re-extract all samples with TRIzol.
Poor digestion dynamics showing either insignificant (e.g., Figure 2B) or, on the contrary, too rapid RNA degradation across samples
• Too low/high RNase activity or concentration range.
• Suboptimal reaction conditions.
• Increase/decrease the amount of RNase.
• Change the enzyme batch.
• Increase/decrease reaction temperature (between 0 and 37 °C).
• Increase/decrease reaction time (1–15 min).
• Try to adjust the salt concentration, specific divalent cations, or pH, based on known RNase preferences.
• We recommend, before attempting the complete CoLoC-seq experiment, to perform test digestions in total cell or organelle lysates (as described in part D of the protocol), using a variety of enzymes and reaction conditions (see Figure S1 in Jeandard et al., 2023).
In the CoLoC-seq setup, bona fide resident transcripts are degraded as if they were contaminants • Too harsh isolation protocol compromised the integrity of the organelles. • Use a milder cell disruption and/or organelle isolation protocol.
In the CoLoC-seq setup, all RNAs look at least partially protected
• Cells were not sufficiently disrupted.
• Cell debris contaminated the organelle prep.
• Use additional/stronger disruption and check the material under a light microscope.
• Add an additional low-speed centrifugation step to further remove cell debris.
• Take up the supernatant after the low-speed centrifugation more cleanly, discarding the lower part of the supernatant together with the debris pellet.
Poor RNA yield in the Mock CoLoC-seq setup • Insufficient lysis of the organelles.
• Use a stronger/more concentrated non-ionic detergent.
• Apply extra mechanical force (Dounce homogeniser, syringe, sonication; but beware of RNA shearing!).
Acknowledgments
This work was supported by the University of Strasbourg through the IdEx Unistra [ANR-10-IDEX-0002], the SFRI-STRAT_US project [ANR 20-SFRI-0012], EUR IMCBio [ANR17-EURE-0023], and IdEx – Attractivité [ANR-10-IDEX-0002-02 to Al.S.], and by the Centre National de la Recherche Scientifique (CNRS). The CoLoC-seq method was first published in Jeandard et al. (2023).
Competing interests
There are no conflicts of interest or competing interests.
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Supplementary information
Scripts used for read alignment, coverage calculation, and feature quantification of human mitochondrial CoLoC-seq data can be found on Zenodo (https://doi.org/10.5281/zenodo.6389451).
The CoLoC-seq and Mock-CoLoC-seq sequencing data published in Jeandard et al. (2023) were deposited in NCBI Gene Expression Omnibus (GEO) under accession number GSE183167 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE183167).
Article Information
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© 2023 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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Peer-reviewed
Immunoprecipitation of Reporter Nascent Chains from Active Ribosomes to Study Translation Efficiency
RC Roberta Cacioppo
CL Catherine Lindon
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4821 Views: 687
Reviewed by: Dipak Kumar PoriaRanen AvinerRitu Gupta
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Original Research Article:
The authors used this protocol in eLIFE May 2023
Abstract
The study of translation is important to the understanding of gene expression. While genome-wide measurements of translation efficiency (TE) rely upon ribosome profiling, classical approaches to address translation of individual genes of interest rely on biochemical methods, such as polysome fractionation and immunoprecipitation (IP) of ribosomal components, or on reporter constructs, such as luciferase reporters. Methods to investigate translation have been developed that, however, require considerable research effort, including addition of numerous features to mRNA regions, genomic integration of reporters, and complex data analysis. Here, we describe a simple biochemical reporter assay to study TE of mRNAs expressed from a transiently transfected plasmid, which we term Nascent Chain Immunoprecipitation (NC IP). The assay is based on a plasmid expressing an N-terminally Flag-tagged protein and relies on the IP of Flag-tagged nascent chains from elongating ribosomes, followed by quantitative reverse transcription polymerase chain reaction (RT-qPCR) quantification of eluted mRNA. We report that elution of mRNA following IP can be achieved by treatment with puromycin, which releases ribosome-mRNA complexes, or with purified Flag peptide, which instead releases nascent chain-ribosome-mRNA complexes. In the example described in this protocol, untranslated regions (UTRs) of a gene of interest were used to flank a FlagVenus coding sequence, with the method allowing to infer UTR-dependent regulation of TE. Importantly, our method enables discrimination of translating from non-translating mRNAs. Additionally, it requires simple procedures and standard laboratory equipment. Our method can be used to test the effect of regulators, such as microRNAs or therapeutic drugs or of various genetic backgrounds, on translation of any user-selected mRNA.
Key features
• The novel NC IP protocol builds upon a previously published method for detection of mRNA-binding proteins (Williams et al., 2022).
• The NC IP protocol is adapted for detecting mRNA actively undergoing translation.
• The method uses mammalian cell culture but could be adapted to multiple organisms, including budding yeast (S. cerevisiae).
Graphical overview
Design of the Nascent Chain Immunoprecipitation (NC IP) reporter and assay. Left. The construct carries a 3× Flag tag at the N-terminal end of Venus protein (FlagVenus). In this example, the reporter is adapted to study untranslated regions (UTR)-dependent expression by flanking FlagVenus coding sequence with UTRs of Aurora kinase A (AURKA) mRNA (depicted reporters refer to Cacioppo et al., 2023, Figure 3). The depicted reporters carry mutations in the proximal (p) or distal (d) polyadenylation signal (PAS). Right. Following reporter transfection, ribosomes are locked onto reporter mRNA by treating cells with cycloheximide (CHX), which prevents ribosome run-off and additional rounds of elongation, before cell lysis and immunoprecipitation (IP) of FlagVenus nascent chains via anti-Flag beads. Reporter mRNAs are then eluted, isolated, and quantified by RT-qPCR.
Keywords: Reporter assay UTR regulation mRNA translation Transient transfection Nascent chain immunoprecipitation One-step RT-qPCR
Background
Translation is the process of protein synthesis using messenger RNAs (mRNAs) as template molecules, occurring in ordered steps (Blanchet and Ranjan, 2022), typically modulated by 5′ and 3′ untranslated regions (UTRs) (Hinnebusch et al., 2016; Mayr, 2019). The gold standard method for measurements of translation efficiency (TE) is ribosome profiling (Ingolia et al., 2009). However, this method allows for genome-wide analysis only, whereas, in some cases, specific investigation of TE of individual mRNAs might be of interest. A typical approach to understanding TE of specific mRNAs is to complement measurements of protein abundance with measurements of mRNA stability. Luciferase reporters are often used, with the luciferase coding sequence flanked by UTR(s) of interest. This approach, however, ignores influences of protein maturation and degradation rates. Alternatively, a measure of TE is provided by quantifying target mRNAs recovered after fractionation of polysomes or immunoprecipitation (IP) of ribosomal components. However, although highly translating mRNAs are loaded with a high number of ribosomes (Guttman et al., 2013; Ingolia et al., 2014; Liang et al., 2018), it is highly debatable whether a high number of ribosomes on transcripts is per se a robust indicator of high TE specifically of coding regions. Untranslated regions may instead be loaded with ribosomes actively engaged in translation (Ingolia et al., 2014); alternatively, transcripts may be bound by ribosomes while lacking coding capacity (Guttman et al., 2013). Additionally, neither approach provides information regarding which part of the endogenous mRNA is undergoing translation. This is relevant given that a multiplicity of mRNAs contain upstream open reading frames (Renz et al., 2020). Advanced imaging- and single cell–based methods have been developed (Biswas et al., 2019). Genetically encoded reporters enable visualization of translation but require co-transfection of multiple factors and intricate cloning for the addition of extensive features to reporter UTRs, e.g., arrays of stem–loops. Also, the requirement for sophisticated imaging platforms and complex data analysis are some other limitations of these recent methods.
Here, we describe a simple yet effective alternative method that we recently introduced to assess TE of mRNAs of interest as dictated by UTRs (Cacioppo et al., 2023). Our assay is based on IP of Flag-tagged nascent chains, ensuring exclusion of non-translating mRNAs, which has already been proved an efficient approach (Raue et al., 2007; Aviner et al., 2013; Zhang et al., 2013). We have found the method particularly useful to measure the role of 3′UTRs, and it could be expanded to probe the role of regulators such as miRNAs, or of various genetic backgrounds, on translation. In addition, since our assay measures TE of transfected reporters, the translation of any specific segments of a given mRNA can be assessed independently. However, although our assay requires simple protocol steps, standard laboratory equipment, and straightforward data analysis, it is more laborious than luciferase assays and not suitable for handling a large number of samples. In addition, contrarily to the polysome fractionation and ribosomal components IP methods, our assay cannot assess TE of multiple mRNAs simultaneously. Nonetheless, we believe that our assay is a valuable addition to the tools available to researchers for the study of translation of mRNAs of interest.
Materials and reagents
Biological materials
Human osteosarcoma epithelial cell line (U2OS) or other cell lines
Plasmids expressing 3× Flag-tagged reporter protein flanked by UTRs of interest from Cacioppo et al. (2023).
RT-qPCR forward (CTGACCCTGAAGCTGATCT) and reverse (GCATGGCGGACTTGAAGAAG) primers targeting the Venus coding sequence on the reporter mRNA
Reagents
Cell culture medium [DMEM supplemented with 10% fetal bovine serum (FBS), 200 μM GlutaMAX-1, 100 U/mL penicillin, 100 μg/mL streptomycin, and 250 ng/mL fungizone] or other media as appropriate for the selected cell line
Phosphate-buffered saline (PBS) (Severn Biotech, catalog number: 207410)
Trypsin/EDTA solution (VWR, catalog number: LZBE02-007E)
Neon transfection system 100 μL kit (Thermo Fisher Scientific, catalog number: MPK10025)
Hanks’ balanced salt solution (HBSS) (Hyclone, catalog number: 10739674)
RNaseZap (Sigma, catalog number: R2020)
UltraPure DNase/RNase-free distilled water (Thermo Fisher Scientific, catalog number: 10977035)
Protector RNase inhibitor (Roche, catalog number: 3335399001)
EDTA (Invitrogen, catalog number: 10458654)
Complete Mini EDTA-free protease inhibitor (Roche, catalog number: 11836170001)
Tris-HCl pH 7.5 (Fisher Scientific, catalog number: BP1757-100)
Lithium chloride (LiCl) (Sigma, catalog number: L7026)
Tris-buffered saline (TBS) (Sigma, catalog number: T5912)
Cycloheximide (CHX) (Sigma, catalog number: 01810)
Triton X-100 (VWR, catalog number: 28817.295)
Anti-Flag M2 magnetic beads (Sigma, catalog number: M8823)
Puromycin (Alfa Aesar, catalog number: J67236.XF)
3× Flag peptide (Sigma, catalog number: F4799)
Ethanol (absolute) (Sigma, catalog number: 24105-2.5L-M)
Luna Universal One-Step RT-qPCR kit (NEB, catalog number: E3005S)
Monarch RNA Cleanup kit (NEB, catalog number: T2040L)
Solutions
3× Flag peptide elution buffer (manufacturer’s instructions)
Lysis buffer (see Recipes)
Wash buffer (see Recipes)
Puromycin elution buffer (see Recipes)
Recipes
Critical: Prepare all solutions in RNase-free conditions.
Note: All solutions should be prepared fresh. However, buffers can be stored at 4 °C, provided that some reagents are added just before the relative experimental step (denoted with “*”).
Lysis buffer
Reagent Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 100 mM 250 μL
LiCl (8 M) 500 mM 156 μL
EDTA (0.5 M) 10 mM 50 μL
CHX (20 mg/mL)* 0.1 mg/mL 12.5 μL
Triton X-100 (10%) 0.1% 25 μL
Protease inhibitor* 1× Dissolve ¼ tablet directly in lysis buffer
Protector RNase inhibitor (40 U/μL)* 0.1 U/μL 6.25 μL
H2O n/a 2 mL
Total n/a 2.5 mL
Wash buffer
Reagent Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 10 mM 30 μL
LiCl (8 M) 600 mM 225 μL
EDTA (0.5 M) 1 mM 6 μL
CHX (20 mg/mL)* 0.1 mg/mL 15 μL
Protector RNase inhibitor (40 U/μL)* 0.1 U/μL 7.5 μL*
H2O n/a 2,717 μL
Total n/a 3 mL
Puromycin elution buffer
Reagent Final concentration Quantity
Tris-HCl (1 M, pH 7.5) 10 mM 6 μL
LiCl (8 M) 600 mM 45 μL
EDTA (0.5 M) 1 mM 1.2 μL
Protector RNase inhibitor (40 U/μL)* 0.1 U/μL 1.5 μL*
Puromycin (200 μM) 20 mM 6 μL
H2O n/a 540 μL
Total n/a 600 μL
Laboratory supplies
Ice
Tissue culture dishes (15 cm) (Nunc, catalog number: 168381)
Tissue culture dishes (10 cm) (Nunc, catalog number: 150350)
Sterile pipette filter tips (10, 200, 1,000 μL) (Starlab, catalog number: S1122)
10 mL plastic pipettes (Fisher Scientific, catalog number: 11839660)
50 mL centrifuge tubes (Greiner, catalog number: 227285)
Sterile and RNase-free 1.5 mL centrifuge tubes
Cell scrapers (Fisher, catalog number: 11597692)
96-well reaction plates (Thermo Fisher Scientific, catalog number: 4346906)
Optical adhesive films (Thermo Fisher Scientific, catalog number: 4360954)
Equipment
Pipettes (10 μL, 200 μL, 1,000 μL) (Gilson)
Pipetboy (Integra)
Centrifuge (Thermo Fisher Scientific, catalog number: 15868722)
MiniSpin centrifuge (Eppendorf, catalog number: 5452000060)
Neon Transfection System (Thermo Fisher Scientific, catalog number: MPK5000)
Neon Transfection System Pipette (Thermo Fisher Scientific, catalog number: MPP100)
Micro-volume spectrophotometer NanoDropTM Lite (Thermo Fisher Scientific)
Tube rotator (Fisherbrand, catalog number: 88861050)
Magnetic rack (Invitrogen)
StepOnePlus Real-Time PCR System (Applied Biosystems, catalog number: 4376600)
Ice buckets
4 °C fridge and -20 °C freezer
Software and datasets
Microsoft Excel (Microsoft Corporation) for analysis of RT-qPCR data and creation of graphs. Can be substituted by R or any other software for data analysis.
Procedure
The protocol here presented is designed for analysis of three conditions in a single experiment. If a different number of conditions is required, adapt volumes accordingly. A flowchart of the procedures is shown in Figure 1. See General note 1.
Figure 1. Overview of the protocol steps
Seed 1 × 106 cells onto a 15 cm dish per condition such that cells are confluent after 24 h
Electroporation
Note: Carry out all procedures under a laminar flow hood and follow standard tissue culture procedures.
Prepare materials for the electroporation.
Place medium, trypsin, and HBSS solutions at 37 °C.
Place the electroporation tube into the holder in the Neon Electroporation machine.
Pipette 3 mL of E2 electrolytic buffer from Neon transfection system kit into the electroporation tube.
Prepare 3 × 1.5 mL tubes containing 1 mL of cell medium and keep at 37 °C. These are required later to enable recovery of cells following electroporation.
Prepare 3 × 1.5 mL tubes each containing 3–5 μg of plasmid DNA.
Prepare 3 × 1.5 mL tubes containing 1 mL of PBS. These are required later to rinse the electroporation pipette tip before moving to the next electroporation condition.
Select the transfection program: voltage 1,150 V, width 30 ms, 2 pulses.
Attach a gold electroporation tip to the electroporation pipette.
Detach cells from 15 cm plates.
Remove medium and wash cells in 10 mL of PBS.
Remove PBS and add 1.5 μL of warm trypsin.
Incubate at 37 °C for 3–5 min and then add 15 mL of warm medium.
Transfer cells to a 50 mL Falcon tube.
Centrifuge at 200× g for 3 min at room temperature.
Wash cells in 10 mL of PBS and centrifuge again at 200× g for 3 min.
Remove PBS and wash again in 1 mL of PBS, transferring cells to 1.5 mL tubes.
Centrifuge using MiniSpin at 300× g for 1 min at room temperature.
Remove PBS and equilibrate cells in HBSS by resuspending cell pellet in 1 mL of warm HBSS.
Centrifuge using MiniSpin at 300× g for 1 min.
Resuspend cells in 330 μL of warm HBSS and transfer into appropriate tube with plasmid DNA.
Note: Electroporation is performed in three rounds per condition. For this, use 100 μL per electroporation round plus excess to avoid formation of bubbles during pipetting.
Caution: Air bubbles could cause arcing during electroporation.
Use the electroporation pipette to transfer cells to the Neon electroporation tube. Perform electroporation. Transfer electroporated cells to 1.5 mL tube with 1 mL of warm medium. Gently invert tube and place at 37 °C. Caution: When pipetting electroporated cells into tube with warm medium, avoid contact between the pipette tip and the medium as this could cause arcing due to presence of FBS in the medium.
Repeat step B11 two more times for the same condition.
Before moving to the next condition, rinse the electroporation pipette tip in PBS multiple times.
Centrifuge electroporated cells using MiniSpin at 300× g for 1 min.
Resuspend in 1 mL of warm medium, plate in 10 cm dishes (one dish per condition) with final 10 mL of medium and incubate at 37 °C for 24 h.
Clean and dispose of electroporation kit components according to the manufacturer’s instructions. Critical: At sections C–F, strictly follow standard procedures for handling RNA. Treat work surfaces and pipettes with 70% ethanol and RNaseZap. Carry out all procedures on ice and always keep samples and solutions on ice. See General note 2.
Cell lysis
Prepare solutions.
35 mL of PBS + CHX 0.1 mg/mL
2.5 mL of lysis buffer
1.2 mL of TBS 1×
Add CHX at a final concentration of 0.1 mg/mL to electroporated cells and incubate at 37 °C for 15 min.
Place plate on ice and wash cells once quickly with 10 mL of ice-cold PBS + CHX 0.1 mg/mL.
Scrape quickly in 1 mL of ice-cold PBS + CHX 0.1 mg/mL while keeping plate on ice.
Transfer cells into 1.5 mL tubes and centrifuge using MiniSpin at 300× g for 5 min at 4 °C.
Resuspend cell pellets in 200 μL of ice-cold lysis buffer and incubate on ice for 15 min.
Centrifuge using MiniSpin at 12,000× g for 15 min at 4 °C.
Transfer 180 μL of supernatants (Input) into new 1.5 mL tubes.
Note: Transfer less than 200 μL of supernatant to avoid collection of pelleted material.
Make a 10 μL aliquot for RNA isolation. Keep at 4 °C and perform RNA isolation (see section E) at the same time as the eluted RNA (see section D, steps 9–10).
Immunoprecipitation
Note: Use 40 μL of beads slurry per condition. Beads need to be mixed to achieve a homogenous suspension. Handle beads according to the manufacturer’s instructions. Use a magnetic rack to separate beads from solutions.
Transfer 120 μL of beads slurry to a 1.5 mL tube. Remove solution in which beads were stored.
Wash beads twice in 600 μL of ice-cold TBS 1×.
Note: Gently invert tube 4–5 times to wash beads.
Add 750 μL of ice-cold lysis buffer and split equally into 3 × 1.5 mL tubes.
Add 170 μL of Input samples to respective tube containing beads.
Incubate tubes at 4 °C overnight gently rotating at 10–30 rpm.
Prepare solutions.
3 mL of wash buffer.
600 μL of puromycin elution buffer or 600 μL of 3× Flag peptide elution buffer.
Discard supernatants (flowthrough).
Wash beads twice with 0.5 mL of ice-cold wash buffer.
Note: Gently invert tube 4–5 times to wash beads.
Perform elution. See General note 3.
Elution by puromycin: add 200 μL of puromycin elution buffer to each tube containing beads and incubate at 4 °C for 30 min gently rotating at 10–30 rpm. Collect Elution samples and keep on ice for RNA isolation to be performed straight after.
or
Elution by 3× Flag peptide: add 100 μL of 3× Flag peptide elution buffer to each tube containing beads and incubate at 4 °C for 30 min gently rotating at 10–30 rpm. Collect eluates and repeat step. Collect eluates again and pool with previous respective eluates (total volume of elution samples 200 μL). Keep on ice for RNA isolation to be performed straight after.
Perform RNA purification as in section E.
RNA isolation
Isolate RNA using Monarch RNA Cleanup kit according to the manufacturer’s instructions. These include a step of DNA digestion. Elute in 30 μL of ice-cold nuclease-free H2O.
Assess RNA concentration and purity using the NanoDrop.
Add EDTA at final concentration between 0.1 and 1 mM to samples.
Note: This is optional. However, addition of EDTA may reduce the activity of any contaminating RNases. Make sure that EDTA is compatible with both the RNA isolation method and the reagents for downstream RT-qPCR.
Pause Point: Purified Input and Elution samples can be stored at -80 °C long term.
One-step RT-qPCR
See General note 4.
Thaw RT-qPCR kit components, primers, and purified RNA samples on ice.
Prepare a reaction master mix according to the manufacturer’s instructions. For three conditions, prepare the equivalent of 20 reactions (includes one extra reaction to account for pipetting errors).
Note: Quickly vortex and spin all reaction reagents before use.
Distribute master mix to 19 wells of a 96-well plate.
Distribute each RNA sample to three wells.
Add a volume of H2O equivalent to the volume of RNA samples in a single well to the well corresponding to the no-template control reaction. Critical: Avoid formation of bubbles while pipetting during steps F3–F4. This might interfere with the amplification reaction.
Cover plate with optical adhesive film.
Quickly spin the plate to collect drops at the bottom of wells and to remove potential bubbles.
Cover plate with darkening foil while setting up RT-qPCR machine.
Load plate onto the RT-qPCR machine and start reaction. Perform one-step RT-qPCR according to the manufacturer’s instructions.
Data analysis
Translation efficiency (TE) is calculated using the percent input method, which compares the amount of target mRNA measured in the Elution fraction to the total amount of the target mRNA in the Input fraction. For this, first the ∆Cq is calculated as follows:
∆Cq = Cq(Elution) - [Cq(Input) - Log2(DF)]
where DF is dilution factor (e.g., if 5% of total Input is used for RT-qPCR, the DF is 20). TE of reporter mRNA is then calculated as:
% Input = 100 × 2-∆Cq
Representative data can be found in Figure 2. For each experiment, three technical replicates and three repeats of the experiment should be performed. Appropriate controls for the experiment should be used, like using an untagged reporter protein (see Cacioppo et al., 2023) or untransfected cells.
Representative data
Figure 2. Quantification by RT-qPCR of indicated reporter mRNAs in an experiment designed to test the role of different 3′ untranslated regions (UTR) configurations (D, L, S) of Aurora kinase A (AURKA), eluted via puromycin (left) or 3× Flag peptide (right). Results representative of three repeats of the experiment. See Cacioppo et al., 2023 for details of the experiment.
Validation of protocol
This protocol was validated in Cacioppo et al. (2023). eLife. Differential translation of mRNA isoforms underlies oncogenic activation of cell cycle kinase Aurora A.
General notes and troubleshooting
General notes
Other standard procedures for transfection (e.g., lipofectamine transfection) and cell lysis can be used. Number of cells to transfect per condition should be adapted to transfection efficiency or to the estimated expression levels of the reporter protein and determined empirically for the selected cell type. The cultures should be sub-confluent on the day of harvesting to avoid contact inhibition of translation. Furthermore, in some cell types and under particular states (e.g., genotoxic or proteotoxic stress), ribosomes may exist in stalled forms, suggesting that association of mRNA with ribosomes may not be indicative of active translation in these cases.
Make sure to optimise conditions. As for any procedure that involves handling and quantifying RNA, empirical tests may be required to find optimal conditions. Alternative methods of RNA isolation and RT-qPCR kits could be used, but conditions must be optimised accordingly.
To read about the action of puromycin, consult Blobel and Sabatini (1971) and Aviner (2020). It is important to note that, although being cost efficient and potentially eluting cleaner RNA, puromycin-induced release of ribosomes may be more variable upon conditions as salt, temperature, ribosome state, presence of CHX, or nucleic or amino acid sequence motifs, than 3× Flag-mediated elution. For this, it is recommended to initially test both elution methods in parallel, in order to find the more suitable for one’s own experimental aims and conditions.
It is highly recommended to perform RT-qPCR as soon as possible, to avoid alterations in the composition of the RNA extracts due to eventual RNA degradation. The Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines (Bustin et al., 2009) should be followed when performing RT-qPCR experiments. One-step RT-qPCR can be replaced by sequential cDNA synthesis and RT-qPCR.
To control for efficiency of IP and for optimization purposes, RT-qPCR could be performed with primers to housekeeping genes (e.g., ACTB, GAPDH). In addition, RNA can be isolated from the Flowthrough, as well, and analysed in parallel with Input and Elution samples.
The assay could be adapted to immunoprecipitate nascent chains using other tags different from Flag tag. Use of small peptide tags is recommended to minimise possible influences on translation efficiency.
It is important to note that cell lysates typically contain a higher ratio of mature to nascent reporter protein. Although we have no evidence of this, it may be possible that the mature reporter protein interacts with ribosomes or mRNA outside the context of active translation (e.g., in RNA granules), and this could interfere with the interpretation of results. Furthermore, reporter expression levels will depend on transfection yield, overall translation rates, and time from transfection to harvesting. Therefore, the conditions should be determined empirically. A small-scale transfection experiment could be performed, harvesting samples at multiple time points, and measuring expression by anti-Flag immunoblotting.
Troubleshooting (see Table 1)
Table 1. Troubleshooting
Issue Probable cause Possible Solution
Low RNA yield RNase contamination
• Ensure all instruments, workspaces, and reagents are RNase-free.
• Change gloves routinely.
• Use filtered pipette tips.
• Pre-chill all tubes and buffers before use.
• Increase volume of RNase inhibitor.
• Perform procedures under a laminar flow hood.
Low IP efficiency
(see General note 5)
Low electroporation efficiency
• Make sure to warm medium and solutions at 37 °C.
• Make sure cells are not stressed, damaged, or contaminated by Mycoplasma. Cell density should not be too low or too high and fresh cultures should be used.
• Make sure plasmids have high degree of purity and that a volume < 1/10 of volume of the cell suspension is used.
• Electroporation conditions (voltage, width, pulses) might need to be optimised, especially if using a different cell line.
Immunoprecipitated target RNA is too low
• Combine Elution RNA samples from multiple technical replicates.
• Increase amount of beads slurry.
• Increase stringency of washes.
• Increase duration of the elution step.
Low purity of Input/Eluted RNA
• Increase volume of RNase inhibitor.
• Make sure samples are not contaminated by genomic DNA.
Non-specific binding to beads
• Increase stringency of washes.
• Decrease volume of lysate used.
• Coat beads with bovine serum albumin prior to IP.
Issues with RT-qPCR
• Make sure primers are designed to be optimal. Primer pairs should have amplification efficiency of 90%–110%.
• Make sure pipetting is consistent and accurate.
• Follow RT-qPCR kit manufacturer’s instructions on maximum amount of template RNA to use per reaction.
• General RT-qPCR troubleshooting applies.
Acknowledgments
We are grateful to members of the Lindon Lab and to the staff of Department of Pharmacology for technical support. This work was supported by Biotechnology and Biological Sciences Research Council (BBSRC) (grant no. BB/R004137/1) to CL. RC is supported by David James Studentship from the Department of Pharmacology. The protocol was inspired by a method developed by Williams et al., 2022. The corresponding original research article is Cacioppo et al. (2023). Graphical overview was created with BioRender.com.
Competing interests
There are no conflicts of interest or competing interests.
References
Aviner, R. (2020). The science of puromycin: From studies of ribosome function to applications in biotechnology. Comput. Struct. Biotechnol. J. 18: 1074–1083.
Aviner, R., Geiger, T. and Elroy-Stein, O. (2013). Novel proteomic approach (PUNCH-P) reveals cell cycle-specific fluctuations in mRNA translation. Genes Dev. 27(16): 1834–1844.
Biswas, J., Liu, Y., Singer, R. H. and Wu, B. (2019). Fluorescence Imaging Methods to Investigate Translation in Single Cells. Cold Spring Harbor Perspect. Biol. 11(4): a032722.
Blanchet, S. and Ranjan, N. (2022). Translation Phases in Eukaryotes.In: Entian, KD. (Ed.). Ribosome Biogenesis (pp. 217–228). Methods in Molecular Biology. Humana, New York.
Blobel, G. and Sabatini, D. (1971). Dissociation of Mammalian Polyribosomes into Subunits by Puromycin. Proc. Natl. Acad. Sci. U. S. A. 68(2): 390–394.
Bustin, S. A., Benes, V., Garson, J. A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M. W., Shipley, G. L., et al. (2009). The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments. Clin. Chem. 55(4): 611–622.
Cacioppo, R., Akman, H. B., Tuncer, T., Erson-Bensan, A. E. and Lindon, C. (2023). Differential translation of mRNA isoforms underlies oncogenic activation of cell cycle kinase Aurora A. eLife 12: e87253.
Guttman, M., Russell, P., Ingolia, N. T., Weissman, J. S. and Lander, E. S. (2013). Ribosome Profiling Provides Evidence that Large Noncoding RNAs Do Not Encode Proteins. Cell 154(1): 240–251.
Hinnebusch, A. G., Ivanov, I. P. and Sonenberg, N. (2016). Translational control by 5’-untranslated regions of eukaryotic mRNAs. Science 352(6292): 1413–1416.
Ingolia, N. T., Ghaemmaghami, S., Newman, J. R. S. and Weissman, J. S. (2009). Genome-Wide Analysis in Vivo of Translation with Nucleotide Resolution Using Ribosome Profiling. Science 324(5924): 218–223.
Ingolia, N. T., Brar, G. A., Stern-Ginossar, N., Harris, M. S., Talhouarne, G. J., Jackson, S. E., Wills, M. R. and Weissman, J. S. (2014). Ribosome Profiling Reveals Pervasive Translation Outside of Annotated Protein-Coding Genes. Cell Rep. 8(5): 1365–1379.
Liang, S., Bellato, H. M., Lorent, J., Lupinacci, F. C. S., Oertlin, C., van Hoef, V., Andrade, V. P., Roffé, M., Masvidal, L., Hajj, G. N. M. and Larsson, O. (2018). Polysome-profiling in small tissue samples. Nucleic Acids Res. 46(1): E3.
Mayr, C. (2019). What Are 3’ UTRs Doing?. Cold Spring Harbor Perspect. Biol. 11(10): a034728.
Raue, U., Oellerer, S. and Rospert, S. (2007). Association of Protein Biogenesis Factors at the Yeast Ribosomal Tunnel Exit Is Affected by the Translational Status and Nascent Polypeptide Sequence. J. Biol. Chem. 282(11): 7809–7816.
Renz, P. F., Valdivia-Francia, F. and Sendoel, A. (2020). Some like it translated: small ORFs in the 5′UTR. Exp. Cell. Res. 396(1): 112229.
Williams, T. D., Cacioppo, R., Agrotis, A., Black, A., Zhou, H. and Rousseau, A. (2022). Actin remodelling controls proteasome homeostasis upon stress. Nat. Cell Biol. 24(7): 1077–1087.
Zhang, Y., Wölfle, T. and Rospert, S. (2013). Interaction of Nascent Chains with the Ribosomal Tunnel Proteins Rpl4, Rpl17, and Rpl39 of Saccharomyces cerevisiae. J. Biol. Chem. 288(47): 33697–33707.
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4,822 | https://bio-protocol.org/en/bpdetail?id=4822&type=0 | # Bio-Protocol Content
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Peer-reviewed
Fractionation of Native Protein Complexes from Mammalian Cells to Determine the Differential Proteasome Activity and Abundance
AF Ana Paula Zen Petisco Fiore
CV Christine Vogel
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4822 Views: 791
Reviewed by: Neha NandwaniPetru-Iulian Trasnea Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in bioRxiv Apr 2022
Abstract
Eukaryotic cells have different types of proteasomes that differ in size. The smallest proteolytically active particle is the 20S proteasome, which degrades damaged and oxidized proteins; the most common larger particle is the 26S proteasome, which degrades ubiquitylated proteins. The 26S proteasome is formed by a 20S particle capped with one or two regulatory particles, named 19S. While proteasome particles function in the cytoplasm, endoplasmic reticulum, and nucleus, our understanding of their abundance and activity in different cellular compartments is still limited. We provide a three-step protocol that first involves detergent-based fractionation of the cytoplasmic and nuclear compartments, maintaining the integrity and activity of proteasome complexes. Second, the protocol employs native gel separation of large multiprotein complexes in the fractions and a fluorescence-based in-gel quantitation of the activity and different proteasome particles. Finally, the protocol involves protein in-gel denaturation and transfer to a PVDF membrane. Western blotting then detects and quantifies the different proteasome particles. Therefore, the protocol allows for sensitive measurements of activity and abundance of individual proteasome particles from different cellular compartments. It has been optimized for motor neurons induced from mouse embryonic stem cells but can be applied to a variety of mammalian cell lines.
Key features
• Protocol for fractionation of active nuclear and cytoplasmic proteasome complexes.
• Native electrophoresis and fluorescence-based in-gel activity assay, which allows the visualization and quantification of active complexes within the acrylamide gel matrix.
• In-gel protein denaturation followed by transfer of complexes to PVDF membrane, which allows the analysis of complexes’ abundance using antibodies.
Graphical overview
Keywords: Native protein complexes fractionation 20S and 26S proteasomes Activity assay Induced neurons Electrophoresis
Background
The proteasome is an essential component of ATP-dependent proteolysis in eukaryotic cells and is responsible for the degradation of most cellular proteins (Goldberg, 2003; Rousseau and Bertolotti, 2018). It is therefore essential for the maintenance of proteostasis. Since the loss of proteostasis is a hallmark of several diseases, the proteasome has become an attractive drug target (Schmidt and Finley, 2014). Several proteasome inhibitors have either been approved for clinical use or are currently in clinical trials (Buac et al., 2013).
There are two major proteasome subunits within the cell, in addition to other less common subunits (Dahlmann, 2016). The smaller subunit comprises the 20S proteasome (700 kDa), which degrades damaged and oxidized proteins, without the need for ubiquitylation. In comparison, the larger subunit, named 26S (2,000–3,000 kDa) proteasome, selectively degrades ubiquitylated proteins. The 26S proteasome is formed by one 20S proteasome capped with one or two regulatory subunits named 19S (Tomko and Hochstrasser, 2013). The barrel-shaped 20S proteasome consists of four stacked heteroheptameric rings: two α-rings on the ends that sandwich a pair of β-rings in between. The six proteolytically active sites are located within the β-rings (Tomko and Hochstrasser, 2013). Entry of ubiquitylated proteins into the 20S is controlled by the 19S subunit, which has specific ubiquitin receptors (Martinez-Fonts et al., 2020). The abundance and activity of the different proteasome subunits are extensively controlled, involving a network of chaperones and regulators (Marshall and Vierstra, 2019; Schnell et al., 2022).
Proteasome-based degradation has first been detected in the cytoplasm and the endoplasmic reticulum (von Mikecz, 2006; Maghames et al., 2018). However, later work also identified highly active degradation of ubiquitylated proteins in the nucleus by the nuclear proteasome (von Mikecz, 2006). Most recent evidence points to AKIRIN2 as a highly conserved protein that controls the nuclear import of proteasomes (de Almeida et al., 2021). These developments highlight the importance of understanding the differential abundance and activity of proteasome subunits in the cytoplasm and nucleus of eukaryotic cells.
This protocol represents a significant advancement in cellular fractionation techniques, as it enables the separation of cellular components while preserving the integrity of protein complexes. We adapted two previously published protocols that allow fractionation of native proteins using detergents (DeCaprio and Kohl, 2020) and a native separation of complexes under nondenaturing conditions with an in-gel proteasome activity assay (Yazgili et al., 2021). Firstly, the plasma membrane is selectively permeabilized using digitonin, which effectively solubilizes it without disrupting intracellular membranes that contain lower levels of cholesterol. Digitonin binds to the membrane, forming pores by complexing with membrane cholesterol and other sterols (Tang et al., 2021). Next, the cytoplasmic lysate is isolated, and the remaining pellet is washed. This step removes contaminating lipidic membranous organelles, resulting in a highly enriched nuclear fraction (Ridsdale et al., 2006; Malhotra, 2008). The different proteasome subunits are then separated using gel separation techniques under native conditions that preserve their functional activity. Both the loading and running buffers used in this step contain ATP, providing an available energy source. Moreover, the buffers contain DTT, which protects against non-proteasomal protease activities. To assess proteasome activity, the protocol utilizes fluorogenic peptides known as Suc-LLVY-AMC, which act as substrates for the 20S and 26S proteasome subunits (Goldberg, 2003). When the peptides are cleaved by the proteasome, the fluorophore called AMC is released, generating a fluorescent signal that can be measured, including as an in-gel activity assay (Yazgili et al., 2021).
After performing gel electrophoresis, the gel content is denatured, and the content is transferred onto a PVDF membrane to facilitate further analysis using specific antibodies. The abundance of the 20S core subunit of the proteasome can be determined by detecting Psma1-7, while the presence of the 19S regulatory subunit is indicated by Psmc3. The colocalization of Psmc3 and Psma1-7 antibodies provides confirmation of the assembly of the fully functional 26S proteasome subunit. The intensity of the bands observed on the PVDF membrane reflects the relative abundance of the proteins, serving as a reliable indicator of the respective subunit quantities.
Overall, this protocol offers a robust method for isolating and quantifying active proteasome subunits, enabling a comprehensive analysis of their functional state.
Materials and reagents
Materials
Cell cultures in plates or suspension (here: neurons differentiated from mouse embryonic stem cells)
0.45 μm sterile filter (Corning, catalog number: 430514)
Cell lifter (Sigma, catalog number: SILA0008)
1.5 mL tubes (Eppendorf, catalog number: 022363204)
Pipette tips (USA Scientific, catalog number: various)
Glass slides (Corning, catalog number: 12-553-10)
Coverslips (Menzel Gläser, catalog number: 630-2118)
NU-PAGE 3%–8% tris-acetate (Thermo Fisher, catalog number: EA0378)
PVDF membrane (Millipore, catalog number: IPFL00010)
Micropipettes (Gilson, catalog number: various)
Microcentrifuge (Eppendorf, catalog number: 5427R)
Sonicator (Covaris, catalog number: S220)
Mini gel tank (Thermo Fisher, catalog number: A25977)
Power supply (Bio-Rad, catalog number: 1645050)
Mini Trans-Blot Cell (Bio-Rad, catalog number: 1703930)
Fluorescence imaging system Gel Doc (Bio-Rad) with the filter for ethidium bromide
NIR imaging system (Odyssey Clx, LiCOR Biosciences)
Cold room at 4 °C
Shaker (Boekel Scientific, catalog number: 260350)
Reagents
ddH2O
Phosphate buffered saline (PBS) (Thermo Fisher Scientific, catalog number: 14190250)
PIPES (Sigma-Aldrich, catalog number: P6757)
NaCl (Fisher Scientific, catalog number: BP350)
MgCl2 hydrate (Sigma-Aldrich, catalog number: M2670)
EDTA (EMD Millipore, catalog number: 324503)
Digitonin (EMD Millipore, catalog number: 300410)
Protease inhibitors (Roche, catalog number: 11697498001)
NaOH (Sigma-Aldrich, catalog number: L4509)
Trypan blue (Invitrogen, catalog number: 15250061)
Igepal (Sigma-Aldrich, catalog number: I8896)
DC Protein quantification kit (Bio-Rad, catalog number: 5000111)
Orange G (Sigma-Aldrich, catalog number: 861286)
Glycerol (Sigma-Aldrich, catalog number: G2025)
Tris (Sigma-Aldrich, catalog number: T6066)
DTT (Cayman Chemical Company, catalog number: 700416)
ATP (Sigma-Aldrich, catalog number: A2383)
Boric acid (Sigma-Aldrich, catalog number: B6768)
MgCl2 (Sigma-Aldrich, catalog number: M2670)
Tris-HCl pH 7.5 (Thermo Fisher Scientific, catalog number: 15567027)
Suc-LLVY-AMC (Bachem, catalog number: 4011369.0100)
Sodium dodecyl sulfate (SDS) (Fisher Scientific, catalog number: BP166-500)
Na2CO3 (Sigma-Aldrich, catalog number: S7795)
2-mercaptoethanol (Sigma-Aldrich, catalog number: M6250)
Glycine (Sigma-Aldrich, catalog number: G8898)
Methanol (Sigma-Aldrich, catalog number: 179337)
Tris Buffered Saline (TBS 20×) (VWR, catalog number: J640)
Tween 20 (Sigma-Aldrich, catalog number: P7949)
Blocking buffer in TTBS (LiCOR Biosciences, catalog number: 927-60001)
Core proteasome units (20S), Psma1-7 antibody (1:1,000) (Enzo Life Sciences, catalog number: BML-PW8195-0100)
Regulatory proteasome units (26S), Psmc3 antibody (1:1,000) (Cell Signaling Technology, catalog number: 25430S)
Mouse secondary antibody (1:10,000) (LiCOR Biosciences, catalog number: 925-32210)
Rabbit secondary antibody (1:10,000) (LiCOR Biosciences, catalog number: 926-68073)
Solutions
4× PIPES buffer (see Recipes)
Cytoplasmic extraction buffer (pH 6.8) (see Recipes)
Nuclear extraction buffer (pH 6.8) (see Recipes)
4× Loading buffer (pH 7.3) (see Recipes)
8× TBE buffer pH 8.3 (see Recipes)
Native gel running buffer (pH 8.3) (see Recipes)
250× ATP/MgCl2 (see Recipes)
Reaction buffer for in-gel proteasome activity assay (see Recipes)
Denaturation buffer (see Recipes)
Tris-glycine transfer buffer (see Recipes)
T-TBS (see Recipes)
Recipes
4× PIPES buffer
Reagent Final concentration Quantity
NaCl 400 mM 5.8 g
PIPES 40 mM 3 g
1 M NaOH 1 M enough to dissolve PIPES
MgCl2 hydrate 12.5 mM 0.64 g
Final volume 250 mL
Dissolve NaCl in 150 mL of ddH2O. Dissolve PIPES in a small volume of NaOH (start with 2 mL and carefully increase, only if necessary). Mix the PIPES solution with NaCl solution. Add MgCl2 hydrate to the mixture. Adjust the final volume to 250 mL. Filter through a 0.45 μm sterile filter. Store at 4 °C or aliquot and freeze (for longer storage).
Cytoplasmic extraction buffer (pH 6.8)
Reagent Final concentration Quantity
4× PIPES buffer 1× 25 mL
Digitonin 0.015% 18.75 mg
EDTA 3 mM 0.6 mL
Protease inhibitors 1 tablet for 10 mL of buffer
Final volume 100 mL
Dissolve digitonin in 10 mL of 4× PIPES buffer. Add the remaining 4× PIPES buffer, EDTA, and 20 mL of ddH2O. Allow the mixture to cool at 4 °C. Adjust the pH to 6.8 and add ddH2O for a total volume of 100 mL. The cytoplasmic extraction buffer can be kept at 4 °C for up to one month. The protease inhibitors should be added fresh to the mixture.
Nuclear extraction buffer (pH 6.8)
Reagent Final concentration Quantity
4× PIPES buffer 1× 25 mL
Igepal 0.1% 100 μL
EDTA 3 mM 0.6 mL
Protease inhibitors 1 tablet for 10 mL of buffer
Final volume 100 mL
Add 4× PIPES buffer, Igepal, EDTA, and 20 mL of ddH2O. Allow the mixture to cool at 4 °C. Adjust the pH to 6.8 and add ddH2O for a total volume of 100 mL. The nuclear extraction buffer can be kept at 4 °C for up to one month. The protease inhibitors should be added fresh to the mixture.
4× Loading buffer (pH7.3)
Reagent Final concentration Quantity
Orange G 0.01% 1 g
Glycerol (87%) 43.50% 5 mL
Tris (pH 7.5, 500 mM) 250 mM 5 mL
Final Volume 10 mL
Dissolve all the chemicals for a final volume of 10 mL in ddH2O; the buffer should be kept at 4 °C.
8× TBE buffer pH 8.3
Reagent Final concentration Quantity
Tris base 712 mM 86.4 g
Boric acid 712 mM 44 g
EDTA Na2 16 mM 5.6 g
Final volume 1 L
Native gel running buffer (pH 8.3)
Reagent Final concentration Quantity
8x TBE buffer 1× 100 mL
ATP 413 μM 200 mg
MgCl2 (1 M) 2 mM 1.6 mL
DTT (1 M) 0.5 mM 400 μL
Final volume 800 mL
Dissolve all chemicals in 600 mL of ddH2O, adjust the pH to 8.3, and complete with ddH2O to a final volume of 800 mL. This solution can be prepared in advance and kept at 4 °C; however, ATP, MgCl2, and DTT need to be added freshly to the mixture.
250× ATP/MgCl2
Reagent Final concentration Quantity
MgCl2 hydrate 2.5 M 1.0165 g
ATP 250 mM 0.3026 g
Final volume 2 mL
Dissolve all the chemicals for a final volume of 2 mL in ddH2O; the buffer should be kept at -80 °C or prepared fresh.
Reaction buffer for in-gel proteasome activity assay
Reagent Final concentration Quantity
Tris (pH 7.5, 100 mM) 1× 12.5 mL
250× ATP/MgCl2 1× 100 μL
DTT (1 M) 1 mM 25 μL
Suc-LLVY-AMC (2 mM, dissolved in DMSO) 48 μM 600 μL
Final Volume 25 mL
Dissolve all chemicals in 25 mL of ddH2O. The reaction buffer can be prepared in advance and kept at 4 °C. However, ATP and DTT should be added freshly to the mixture, while the proteasome substrate Suc-LLVY-AMC should be added immediately before incubation of the gels in reaction buffer.
Denaturation buffer
Reagent Final concentration Quantity
SDS 2% 10 g
Na2CO3 66 mM 3.5 g
β-mercaptoethanol 1.50% 7.5 mL
Final volume 500 mL
In a fume hood, dissolve all chemicals in 500 mL of ddH2O.
Tris-glycine transfer buffer
Reagent Final concentration Quantity
Tris 48 mM
Glycine 39 mM
SDS 1.3 mM
Methanol 20% 200 mL
Final volume 1 L
Dissolve Tris and glycine in 600 mL of ddH2O and adjust to pH 8.3. Add SDS and methanol. Adjust the volume to 1 L with ddH2O. The transfer buffer should be kept at 4 °C until use.
T-TBS
Reagent Final concentration Quantity
20× TBS 1× 100 mL
Tween 20 0.10% 2 mL
Final volume 2 L
Dilute 10× TBS in 1.7 L of ddH2O, add the Tween 20, and adjust with ddH2O to a final volume of 2 L.
Time considerations
Native protein fractionation: 2–3 h.
Native gel electrophoresis: 4 h.
Activity assay: 45 min.
Transfer to membrane and immunoblot: 2 days.
Software
Image StudioTM Acquisition Software (LiCOR Biosciences)
Fiji ImageJ (https://imagej.net/software/fiji/)
Procedure
Native protein fractionation
This protocol is described for one replicate of 600,000 cells maintained in one 10 cm dish.
Maintain cells as needed. Here, we maintained cells in a 10 cm dish following published protocols (Mazzoni et al., 2013; An et al. 2019). All steps below were developed for starting material of 600,000 cells. The density/confluency of cells is not expected to alter intracellular proteasome subunits’ availability.
On the day of collection, aspirate the media from the plate and wash the plate once with 5 mL of ice-cold PBS.
Add 500 μL of cytoplasmic extraction buffer and, using the cell lifter, scrape cells from plates. The suspension should be kept on ice until 90%–95% of the cells are lysed.
Determine cell lysis by using a small amount of the suspension, staining with trypan blue, and examining under an optical microscope. Briefly, take 10 μL of the cell suspension and mix it with an equal volume of trypan blue solution. Mix gently to ensure uniform staining. Allow the cell–trypan blue mixture to incubate for 3–5 min at room temperature and cover the mixture with a coverslip. Examine the cells under a microscope and distinguish between unstained and stained blue cells. Successful cell lysis will result in stained cells due to membrane rupture. We suggest the lysis to be assessed by microscopy 10–15 min after scraping the cells.
In our hands, 95% lysis is reached after approximately 30 min. It is expected that a similar period is enough to reach a proper lysis for other cell types in culture.
Centrifuge lysed cells at 480× g for 10 min at 4 °C. A white pellet enriched in nuclei will appear at the bottom of the tube.
Transfer the supernatant (cytoplasmic fraction) to a fresh tube and snap-freeze. Store cytoplasmic fraction at -80 °C until further use.
Wash the nuclei pellet three times with 200 μL of cytoplasmic extraction buffer and centrifuge at 480× g for 10 min at 4 °C.
Resuspend the pellet in 50 μL of nuclear extraction buffer, pipetting the pellet up and down.
Keep suspension on ice for 30 min. Resuspend the pellet after 10 min.
Sonicate the suspension at 4 °C. Here, the sonicator was programmed to 5% duty factor, three cycles of sonication for 10 s each, with 30 s intervals between each cycle.
Centrifuge at 6,780× g for 10 min at 4 °C.
Transfer the supernatant (nuclear fraction) to a fresh tube and snap-freeze. Store nuclear fraction at -80 °C until use.
Native gel electrophoresis and activity assay
Thaw samples on ice.
Determine total protein concentration with the DC Protein assay kit, following the manufacturer’s instructions. This is a detergent-compatible quantification kit. If using a different kit, ensure its compatibility with detergents in the lysis buffers.
Add the respective lysis buffer to achieve a final protein concentration of 0.5 μg/μL for the cytoplasmic fraction and 0.2 μg/μL for the nuclear fraction.
Add 5 μL of the 4× loading buffer to 15 μL for each fraction.
Separate samples in NU-PAGE at 150 V and 4 °C for 4 h in native gel running buffer. The long run time is due to the large size of the proteasome subunits.
Incubate the gel in the reaction buffer at room temperature for 30 min using a shaker.
Protect gels from light during the reaction.
Handling the gel properly is critical for the success of this part of the analysis. Please check critical notes for tips on proper handling.
Detect the fluorescence at A360 excitation/A460 emission (ethidium bromide filter) using Gel Doc (Figure 1).
Figure 1. Confirmation of fractionation success and localization of Psma1-7 and Psmc3 in induced spinal motor neurons by SDS-PAGE/immunoblot: the cytoplasmic or nuclear Psma1-7 and Psmc3 were detected in induced spinal motor neurons treated with either DMSO (-) or CPA (+). To ensure accurate fractionation, cytoplasmic markers Synaptophysin (Syn) and α-tubulin, as well as the nuclear marker LaminA/C (Lam A/C), were also detected as fractionation controls.
Transfer of proteins to PVDF membrane and abundance analysis by immunoblotting
Denature proteins in the gel by incubation in the denaturation buffer for 10 min at room temperature in a shaker.
Transfer proteins to a low background PVDF membrane in TRIS-glycine transfer buffer at 40 mAV for 16 h.
Activate the PVDF membrane by incubating for 5 min in 20% methanol.
Following activation, equilibrate the membrane and gel for 5 min in the transfer buffer in a shaker prior to use.
Block membranes in blocking buffer at room temperature for 1 h.
Incubate the membranes in blocking buffer with primary antibodies (Psma1-7 and Psmc3) at 4 °C overnight.
Wash membranes with T-TBS five times for 10 min each.
Incubate membranes with mouse and rabbit secondary antibodies in TBS at room temperature for 1 h. We used a secondary antibody before scanning, as the two primary antibodies were from different species.
Scan membrane images with the gel imaging system.
Quantify images (band densitometry) with ImageJ software according to the manufacturer’s instructions. Normalize measurements to the sum of the band intensities on the entire gel.
Data analysis
We confirmed the success of the fractionation by SDS/immunoblotting, as shown in Figure 1. We detected the cytoplasmic or nuclear localization of Psma1-7 and Psmc3. As fractionation control, we detected the cytoplasmic markers Synaptophysin (Syn) and α-tubulin and the nuclear marker LaminA/C (Lam A/C).
Differentiated spinal motor neurons were treated with DMSO (vehicle, control) and cyclopiazonic acid (20 nM CPA, proteotoxic stress) for 12 h. We prepared the cytoplasmic and nuclear extracts as described above and determined proteasome activity and abundance.
The first lane (1) in Figure 2A shows proteasome subunits from control cells, and the second lane (2) describes proteasome subunits from cells treated with CPA. The band intensity signifies subunit activity as measured by proteolysis of the fluorescent substrate.
Figure 2B shows immunoblots of the respective gels against Psmc3 and Psma1-7, which were performed after the activity assay, as described above.
The right panels in Figure 2A/B show the quantified results. In the experiment, statistical significance (p-value < 0.05) was achieved with five replicate experiments with respect to changes in proteasome activity and abundance under stress. The activity of the 20S subunit in the nuclear extract was below the detection limit.
Figure 2. Characterization of proteasome activity and particles abundance in induced spinal motor neurons under stress conditions. A. Proteasome in-gel activity assay of fractionated extracts from induced spinal motor neurons treated with control (1) or stress inductor (2) for 12 h. Proteins were gel-separated under non-denaturing conditions before measuring the activity of the different proteasome particles by the proteolysis of the substrate peptide Suc-LLVY-AMC. The graphs next to the blots show the quantitative results. Intensities measured in the gels were normalized by calculating the ratio between the value measured for individual bands and the sum of all measurements taken for the entire gel. B. Immunoblotting protein extracts from samples described in Figure 2A. Proteins were gel-separated under non-denaturing conditions and transferred to a membrane to quantify the abundance of different proteasome particles using specific antibodies: Psmd2 (red) for single and double-capped 26S particles and Psma1-7 (green) for 20S particles. The graphs next to the blots show the quantitative results. Intensities measured in the gels were normalized by calculating the ratio between the value measured for individual bands and the sum of all measurements taken for the entire gel.
Validation of protocol
This protocol was devised and implemented to assess cytoplasmic and nuclear proteasome activity and the assembly of two types of motor neurons with different sensitivities to misfolded protein stress. The activity and abundance assays were performed in two biological replicates in five independent experiments. The findings of this analysis are represented in Figure 7C and 7D of the currently available preprint at https://doi.org/10.1101/2022.04.10.487765 (Petisco Fiore et al., 2022).
General notes and troubleshooting
General notes
The protocol works well even with limited material: induced motor neurons are small, providing only 5 and 1 μg of protein for the cytoplasmic and nuclear fraction, respectively, from a single 10 cm dish (replicate). The protein amount was sufficient for the proteasome activity and abundance measurements described.
The success of fractionation should be confirmed by SDS-PAGE/immunoblotting, using cytoplasmic and nuclear lysates to detect markers. We use specific antibodies for Lamin A/C (nuclear marker), α-tubulin (cytoplasm marker), and Synaptophysin (neuronal cytoplasmic marker).
Critical parameters and troubleshooting
Insufficient cell lysis decreases the amount and purity of the cytoplasmic and nuclear fractions.
Gently and periodically pipette during cell lysis to enhance plasma membrane disruption. It aids in achieving efficient cell lysis.
Igepal 0.1% is used for lysing the nuclei during the protocol while avoiding disruption of protein complexes. Exceeding this concentration can lead to undesired complex disruption.
The protocol includes ATP and DTT supplements to be added freshly even when using buffers prepared on the previous day.
The extracts should be thawed on ice to prevent the disassembly of protein complexes and loss of proteasome activity.
Suc-LLVY-AMC for activity assays in the protocol should be added freshly to the activity buffer for optimal results.
A high-quality gel used ensures accurate visualization and analysis of protein complexes. Pre-cast native gels are recommended for convenience and consistency.
A spatula for handling soft native gels can be used to prevent folding or ripping the gel when transferring it to the transilluminator for imaging.
Spraying distilled water onto the imaging plate helps to move the gel and prevents it from sticking and tearing during manipulation.
Acknowledgments
This protocol resulted from the adaptation of two published protocols: DOI:10.1101/pdb.prot098582 (DeCaprio and Kohl, 2020) and DOI: 10.1016/j.xpro.2021.100526 (Yazgili et al., 2021). C.V. acknowledges funding from the National Institutes of Health (R35 GM127089) and the Chan Zuckerberg Initiative.
Competing interests
The authors disclose no conflict of interests.
References
An, D., Fujiki, R., Iannitelli, D. E., Smerdon, J. W., Maity, S., Rose, M. F., Gelber, A., Wanaselja, E. K., Yagudayeva, I., Lee, J. Y., et al. (2019). Stem cell-derived cranial and spinal motor neurons reveal proteostatic differences between ALS resistant and sensitive motor neurons. eLife 8: e44423.
Buac, D., Shen, M., Schmitt, S., Rani Kona, F., Deshmukh, R., Zhang, Z., Neslund-Dudas, C., Mitra, B. and Dou, Q. P. (2013). From Bortezomib to other Inhibitors of the Proteasome and Beyond. Curr. Pharm. Des. 19(22): 4025–4038.
Dahlmann, B. (2016). Mammalian proteasome subtypes: Their diversity in structure and function. Arch. Biochem. Biophys. 591: 132–140.
DeCaprio, J. and Kohl, T. O. (2020). Differential Detergent Lysis of Cellular Fractions for Immunoprecipitation. Cold Spring Harb. Protoc. 2020(2): pdb.prot098582.
de Almeida, M., Hinterndorfer, M., Brunner, H., Grishkovskaya, I., Singh, K., Schleiffer, A., Jude, J., Deswal, S., Kalis, R., Vunjak, M., et al. (2021). AKIRIN2 controls the nuclear import of proteasomes in vertebrates. Nature 599(7885): 491–496.
Goldberg, A. L. (2003). Protein degradation and protection against misfolded or damaged proteins. Nature 426(6968): 895–899.
Maghames, C. M., Lobato-Gil, S., Perrin, A., Trauchessec, H., Rodriguez, M. S., Urbach, S., Marin, P. and Xirodimas, D. P. (2018). NEDDylation promotes nuclear protein aggregation and protects the Ubiquitin Proteasome System upon proteotoxic stress. Nat. Commun. 9(1): e1038/s41467-018-06365-0.
Malhotra, V. (2008). Regulated assembly of proteins and lipids at the Golgi to generate membrane fission activity. Chem. Phys. Lipids 154: S3.
Marshall, R. S. and Vierstra, R. D. (2019). Dynamic Regulation of the 26S Proteasome: From Synthesis to Degradation. Front. Mol. Biosci. 6: e00040.
Martinez-Fonts, K., Davis, C., Tomita, T., Elsasser, S., Nager, A. R., Shi, Y., Finley, D. and Matouschek, A. (2020). The proteasome 19S cap and its ubiquitin receptors provide a versatile recognition platform for substrates. Nat. Commun. 11(1): e1038/s41467-019-13906-8.
Mazzoni, E. O., Mahony, S., Closser, M., Morrison, C. A., Nedelec, S., Williams, D. J., An, D., Gifford, D. K. and Wichterle, H. (2013). Synergistic binding of transcription factors to cell-specific enhancers programs motor neuron identity. Nat. Neurosci. 16(9): 1219–1227.
von Mikecz, A. (2006). The nuclear ubiquitin-proteasome system. J. Cell Sci. 119(10): 1977–1984.
Petisco Fiore, A. P. Z., Maity, S., An, D., Rendleman, J., Iannitelli, D., Choi, H., Mazzoni, E. and Vogel, C. (2022). New molecular signatures defining the differential proteostasis response in ALS-resistant and -sensitive motor neurons. bioRxiv: e487765.
Ridsdale, A., Denis, M., Gougeon, P. Y., Ngsee, J. K., Presley, J. F. and Zha, X. (2006). Cholesterol Is Required for Efficient Endoplasmic Reticulum-to-Golgi Transport of Secretory Membrane Proteins. Mol. Biol. Cell 17(4): 1593–1605.
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In-gel proteasome assay to determine the activity, amount, and composition of proteasome complexes from mammalian cells or tissues. STAR Protoc. 2(2): 100526.
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Successful Transfection of MicroRNA Mimics or Inhibitors in a Regular Cell Line and in Primary Cells Derived from Patients with Rheumatoid Arthritis
SW Si Wang
JX Jing Xu
YG Yuanxu Guo
YC Yongsong Cai
WZ Wenhua Zhu
LM Liesu Meng
CJ Congshan Jiang
SL Shemin Lu
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4823 Views: 510
Reviewed by: Navnita Dutta Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Frontiers in Immunology Mar 2021
Abstract
The transfection of microRNA (miRNA) mimics and inhibitors can lead to the gain and loss of intracellular miRNA function, helping us better understand the role of miRNA during gene expression regulation under specific physical conditions. Our previous research has confirmed the efficiency and convenience of using liposomes to transfect miRNA mimics or inhibitors. This work uses miR-424 as an example, to provide a detailed introduction for the transfection process of miRNA mimics and inhibitors in the regular SW982 cell line and primary rheumatoid arthritis synovial fibroblasts (RASF) cells from patients by using lipofection, which can also serve as a reference to miRNA transfection in other cell lines.
Key features
• MiRNA mimics and inhibitors transfection in regular SW982 cell line and primary RASF cells.
• Treatment and culture of RASF primary cells before transfection.
Using liposomes for transfection purposes.
Keywords: miRNA Mimic and inhibitor Lipofection Primary RASF cell
Background
Transfection is a specialized technique for introducing exogenous genes into cells. With the deepening of research on gene and protein functions, transfection has become a common basic method in laboratory work. Transfection can be roughly divided into three methods: physical, chemical, and biological mediation. Electroporation, microinjection, and gene gun are very promising physical methods. There are also many chemical methods, such as the classical calcium phosphate coprecipitation method and the lipofection method. Biological methods include primitive protoplast transfection and various virus-mediated transfection techniques (Kim and Eberwine, 2010).
Lipofection refers to the cationic liposome with a positive charge on the surface, which can interact with the phosphate group of nucleic acid through electrostatic interactions to encapsulate DNA or RNA molecules, forming lipid complexes. It can also be adsorbed by negatively charged cell membranes on the surface and then enter cells through fusion or endocytosis. Lipofection is suitable for transfect DNA or RNA into suspension or adherent cultured cells, and is currently one of the most convenient and efficient transfection methods in the laboratory (Felgner et al., 1987).
MicroRNAs (miRNAs) are a class of non-coding small RNAs composed of 18–25 nucleotides. Mostly, they regulate the expression of various protein cod genes at the post-transcriptional level, by inhibiting translation or inducing mRNA degradation, and have a significant impact on physiological processes such as cell differentiation, proliferation, apoptosis, and hematopoiesis. Hence, abnormal expression of miRNA is pathological and can result in the development and progression of multiple diseases (Bartel, 2004).
Some studies have found multiple miRNA expression disorders in rheumatoid arthritis (RA) patients, suggesting that an abnormal miRNA expression may be the molecular mechanism of the disease. These studies involve various cellular biological behaviors, as well as the regulation of signaling pathways and the study of target genes. With the expansion of research fields, the important role of miRNA in RA is gradually receiving increased attention. For example, miR-18a was found to be an inhibitor of nuclear factor kappa-B (NF-κB) in rheumatoid arthritis synovial fibroblasts (RASF) (Trenkmann et al., 2013); miR-19a is associated with the expression of Toll-like receptors in RASF (Philippe et al., 2012); and miR-23b is related to the expression of various inflammatory cytokines in RASF (Zhu et al., 2012).
As a key cell type in the research of pathogenesis of rheumatoid arthritis, the excessive proliferation and limited apoptosis of RASFs are considered to be the pathological basis of RA, which can directly promote joint destruction and cartilage damage by enhancing the production of matrix degrading enzymes (van der Helm-van Mil and Huizinga, 2008; Bartok and Firestein, 2010). Mimics and inhibitors of miRNAs were transfected into the regular SW982 cell line and the primary RASF cells from patients to study the molecular mechanism of these miRNAs during gain and loss of their function.
Materials and reagents
Biological materials
SW982 cell line (ATCC, catalog number: HTB-93)
RASF primary cell (origin)
Reagents
LipofectamineTM 3000 (Invitrogen, catalog number: L300001)
Mimics and inhibitors (GenePharma company, catalog number: B02001, B03001)
0.25% Trypsin-EDTA (HyClone, catalog number: SH30042.01B)
Fetal bovine serum (FBS) (Sangon biotechnology, catalog number: E51002)
DMEM high glucose culture medium (HyClone, catalog number: SH30022.01B)
Penicillin-Streptomycin 100× (P/S) (HyClone, catalog number: SV30010)
Saline (CR double-crane, catalog number: H20054037)
Phosphate buffered saline (PBS) (Biosharp, catalog number: BL601A)
DNase (Sigma-Aldrich, catalog number: AMPD1)
Hyaluronidase (Sigma-Aldrich, catalog number: 37259-53-3)
Type I collagenase (Sigma-Aldrich, catalog number: SCR103)
TRIzol reagent (Invitrogen, catalog number: 15596-026)
Pierce RIPA buffer (Thermo, catalog number: 89900)
NaOH (Xilong chemistry, catalog number: 7697-37-2)
HCI (Xilong chemistry, catalog number: 121-23-1)
Solutions
SW982 and RASF cell culture medium (see Recipes)
Recipes
SW982 and RASF cell culture medium
Reagent Final concentration Volume
DMEM high glucose medium n/a 450 mL
Fetal bovine serum 10% 50 mL
P/S 0.1% 0.5 mL
Total n/a 500.5 mL
Adjust the pH with NaOH or HCl to 7.2–7.4.
Serum-free DMEM medium refers to the medium without the addition of FBS and antibiotics (the basal medium).
Laboratory supplies
100 mm sterile polystyrene culture dish (Nunc, catalog number: 150340)
6-well sterile polystyrene cell culture plate (Nunc, catalog number: 150239)
15 mL sterile centrifuge tube (KIRGEN, catalog number: KG2611)
1.5 mL centrifuge tube (KIRGEN, catalog number: KG2211S)
Ophthalmic scissors (Shinva, catalog number: ZC056R)
Hemacytometer (Bio-Rad, catalog number: 1450011)
Cell culture flask (Nunc, catalog number: 156340)
Equipment
Cell culture incubator: 37 and 5% CO2 (Thermo, catalog number: THM#3427)
Centrifuge (Eppendorf, catalog number: 5810R)
Biological safety cabinetry (Biobase, catalog number: BSC-1500IIB2-X)
Microscope (Nikon, catalog number: Eclipse E200)
Procedure
Cell preparation
SW982 cells are regularly cultured in DMEM high glucose medium supplemented with 10% FBS and 0.1% P/S and incubated at 37 °C in humid conditions with 5% CO2. RASF cells were isolated from synovial tissues of the knee joints from four RA patients; Table 1 displays the specific information of patients. The procedures for RASF isolation are as follows; a simple flowchart for cell extraction is shown in Figure 1.
Table 1. Patients’ information
Number Gender Age (years) RF (IU/mL) ESR (mm/h) CRP (mg/L) anti-CCP positive ratio
1 Female 37 18.62 90 8.7 18.5
2 Female 60 71.97 113 34.5 3.2
3 Female 45 195.6 87 42.96 110.4
4 Female 66 61.5 31 10.22 33.6
RF: rheumatoid factor
ESR: erythrocyte sedimentation rate
CRP: C-reactive protein
CCP: cyclic citrullinated peptide
Figure 1. Schematic diagram of synovial fibroblasts extraction process. Created with BioRender.com.
Cut off excess adipose tissue from the RA synovial tissue specimen, rinse the synovial tissue with sterile saline until there are no obvious blood stains, select suitable tissues, place it in a sterile culture dish, and use ophthalmic scissors to cut it as small as possible.
Add 5 mL of saline, soak the tissues, and then transfer it into a 15 mL centrifuge tube. Centrifuge at 500× g for 5 min and discard the supernatant.
Add 5 mL of 0.25% trypsin-EDTA for resuspension, type I collagenase (final concentration 1 mg/mL), hyaluronidase (final concentration 0.15 mg/mL), and DNase (final concentration 0.15 mg/mL). Mix them well, digest at 37 for 60–90 min, and shake the centrifuge tube occasionally.
After tissues’ digestion, add 5 mL of culture medium and resuspend. Let it stand until the remaining tissue block in the 15 mL centrifuge tube settles at the bottom and transfer the supernatant to another 15 mL centrifuge tube for centrifugation to obtain cell pellets.
Wash the pellets with 5 mL of culture medium, resuspend, centrifuge, and discard the supernatant. Add 4 mL of culture medium again and repeat the resuspension centrifugation step to collect as many cells as possible.
Count the collected cells with a hemacytometer. Seed 4 million cells per 25 cm2 of the culture bottle. The newly isolated cells are highly diverse. In addition to RASF, there are lymphocytes, osteoclasts, macrophages, and macrophage-like synovial cells. Therefore, the abundant cell density at the very beginning is crucial for the growth of synovial cells.
Place the culture flask inoculated with cells in a 5% CO2 cell incubator at 37 and observe the cell adhesion for any contamination on the next day. On the third day, change the fluid and discard the other types of cells or dead cells, which did not adhere to the bottom of the flask. Passage the cells when the confluence reaches 90%. Primary synovial fibroblasts in good condition are shown in Figure 2.
Figure 2. The expected cell morphology of rheumatoid arthritis synovial fibroblast primary cells. Scale bar: 1 μm.
Cell seeding
Wash thoroughly with PBS buffer three times. After discarding all the liquid, add 1 mL of trypsin, gently move the culture flask to make it even, and incubate it at 37 for approximately 2 min. Observe the cell condition and avoid excessive digestion, which may affect the cell morphology.
Add 4 mL of complete culture medium to terminate the reaction, gently detach the cells from the bottom with a pipette, transfer the cells to a 15 mL centrifuge tube, centrifuge at 1,000× g for 5 min to collect cell pellets, discard the supernatant, add 1 mL of the culture medium, and fully resuspend the cells.
According to the required number of cells in the experiment, inoculate SW982 cells and RASF cell in a 6-well plate overnight and make sure they grow to 80% confluence.
Preparation of the transfection reagents
For each well in the 6-well plate, add 5 μL of LipofectamineTM 3000 reagent and mix well using a pipette in a 1.5 mL centrifuge tube containing 250 μL of serum-free DMEM medium. Incubate at room temperature for 5 min.
Add miRNA mimics or inhibitors and mix them in another centrifuge tube containing 250 μL of serum-free DMEM medium. The sequence information of mimics and inhibitors is shown in Table 2.
Table 2. MiRNA mimics and inhibitors sequence information
Name Product Sequence (from 5′ to 3′)
miRNA NC Mimic Sense UUCUCCGAACGUGUCACGUTT
Antisense ACGUGACACGUUCGGAGAATT
miR-424-5p Mimic
Sense CAGCAGCAAUUCAUGUUUUGAA
Antisense CAAAACAUGAAUUGCUGCUGUU
miRNA NC Inhibitor CAGUACUUUUGUGUAGUACAA
miR-424-5p Inhibitor UUCAAAACAUGAAUUGCUGCUG
NC: negative control
Combine the two mixtures from steps 1 and 2, gently mix 15 times, and incubate at room temperature for 20 min.
Prepare the cells during the incubation period of the previous step. Wash the cells in the 6-well plate with PBS and add 1.5 mL of serum-free culture medium. After the incubation of the transfection reagent, add 500 μL of the mixture from the previous step to each well, mix thoroughly, and incubate in a CO2 incubator. Forty-eight hours later, replace the medium with the complete SW982 and RASF cell culture medium.
Troubleshooting
The temperature of LipofectamineTM 3000 may affect transfection efficiency; therefore, when mixed with LipofectamineTM 3000, the culture medium should be kept at room temperature.
The number and status of cells have an impact on transfection. Resuscitated cells and primary cells should be transmitted to at least the third generation, and the number of cells should be stable before use.
After adding LipofectamineTM 3000 in the experimental steps, the mixing should be gentle to avoid damaging the liposome particles and affecting the transfection efficiency.
The transfection effect depends on the cell state, concentration of transfectants, transfection reagents, operation of transfection processes, etc. Therefore, a pilot study for subsequent transfection efficiency verification is always needed to determine whether they are effective for each individual study. We are usually confident to validate the gain or loss of function for miRNAs and their target genes following transfection with the miRNA mimics or inhibitors for 48 h.
Data analysis
Data were analyzed using SPSS 21. Experimental data were expressed as mean ± standard error of the mean (SEM). Differences between the groups were analyzed using the Mann-Whitney test. p < 0.05 was considered statistically significant (* p < 0.05).
Validation of protocol
The validation of transfection efficiency can be achieved by detecting the intracellular miRNA level following the transfection of exogenous miRNA mimics or inhibitors. Use TRIzol reagent to collect the cell lysate and perform RT-qPCR to verify transfection efficiency. In our published results (Wang et al., 2021), miRNA expression can increase up to a thousand times from its previous intracellular level after transfection of the mimics, and expression of some miRNAs can also be significantly reduced after transfection with inhibitors (Figure 3). This indicates the success of the lipofection method, which is simple and highly reproducible. Validation of target genes is achieved by western blotting, using RIPA buffer to collect protein lysate. After protein quantification, gel electrophoresis, and antibody incubation, the results of target gene expression changes were obtained; this part of the results can be referenced in our published article (Wang et al., 2021) and in Figure 4.
Figure 3. RT-qPCR results showing the RNA expression level of miR-424 in rheumatoid arthritis synovial fibroblasts (RASF) and SW982 cell line following the transfection of its mimics or inhibitors for 48 h. A, B. Expression of miR-424 in 10 nM mimic (A) or 100 nM inhibitor (B)-treated RASF. C, D. Expression of miR-424 in 10 nM mimic (C) or 100 nM inhibitor (D)-treated SW982 cells. Scale bar: mean ± SEM of samples from four patients (A and B) or from three independent cell experiments (C and D) p < 0.05, *: vs. negative control (NC), using Mann-Whitney test.
Figure 4. Western blotting results showing the protein expression of validated miR-424 target gene in SW982 cell line following miR-424 mimic transfection for 48 h. The protein expression of DICER1 (A), CCND1 (B), and CCND3 (C) in SW982 cells following miR-424 mimic transfection. Scale bar: mean ± SEM from three independent cell experiments. p < 0.05, *: vs. negative control (NC), using Mann-Whitney test.
General notes and troubleshooting
General notes
MiRNA mimics and inhibitors were synthesized by the company. Stock solutions at 20 μM were prepared before use and stored at -20 .
NC group with miRNA should be included. All experiments should be carried out in triplicates as a minimum.
According to our previous experience, for miRNA transfection, the final concentration of the 10 nM mimic substance is enough for most miRNA experiments; for inhibitors, a 100 nM should be ok. The optimal concentration could be obtained through pilot experiments.
Acknowledgments
We are grateful for the National Science Foundation of China (grant No. 81671629, 81701619, 81970029, and 81902249) and Shaanxi Province Natural Science Foundation (Project No. 2021JQ-024 and 2018JM7057).
Competing interests
These experimental procedures were adapted from our previous work published in Wang, S. et al., 2021. The authors declare that there are no known competing financial interests or personal relationships that could have appeared to influence this work.
References
Bartel, D. P. (2004). MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116(2): 281–297.
Bartok, B. and Firestein, G. S. (2010). Fibroblast-like synoviocytes: key effector cells in rheumatoid arthritis. Immunol. Rev. 233(1): 233–255.
Felgner, P. L., Gadek, T. R., Holm, M., Roman, R., Chan, H. W., Wenz, M., Northrop, J. P., Ringold, G. M. and Danielsen, M. (1987). Lipofection: a highly efficient, lipid-mediated DNA-transfection procedure. Proc. Natl. Acad. Sci. U. S. A. 84(21): 7413–7417.
Kim, T. K. and Eberwine, J. H. (2010). Mammalian cell transfection: the present and the future. Anal. Bioanal. Chem. 397(8): 3173–3178.
Philippe, L., Alsaleh, G., Suffert, G., Meyer, A., Georgel, P., Sibilia, J., Wachsmann, D. and Pfeffer, S. (2012). TLR2 Expression Is Regulated by MicroRNA miR-19 in Rheumatoid Fibroblast-like Synoviocytes. J. Immunol. 188(1): 454–461.
Trenkmann, M., Brock, M., Gay, R. E., Michel, B. A., Gay, S. and Huber, L. C. (2013). Tumor Necrosis Factor α-Induced MicroRNA-18a Activates Rheumatoid Arthritis Synovial Fibroblasts Through a Feedback Loop in NF-κB Signaling. Arthritis Rheumatol. 65(4): 916–927.
van der Helm-van Mil, A. H. and Huizinga, T. W. (2008). Advances in the genetics of rheumatoid arthritis point to subclassification into distinct disease subsets. Arthritis Res. Ther. 10(2): 205.
Wang, S., Xu, J., Guo, Y., Cai, Y., Ren, X., Zhu, W., Geng, M., Meng, L., Jiang, C., Lu, S., et al. (2021). MicroRNA-497 Reduction and Increase of Its Family Member MicroRNA-424 Lead to Dysregulation of Multiple Inflammation Related Genes in Synovial Fibroblasts With Rheumatoid Arthritis. Front. Immunol. 12: e619392.
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Fluorescence in situ Localization of Pri-miRNAs in Isolated Arabidopsis thaliana Nuclei
TG Tomasz Gulanicz
AZ Agnieszka Zienkiewicz
KZ Krzysztof Zienkiewicz
AK Anna Kasprowicz-Maluski
ZS Zofia Szweykowska-Kulinska
AJ Artur Jarmolowski
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4824 Views: 559
Reviewed by: Durai SellegounderEduardo ListikShreya Mukhopadhyay
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Original Research Article:
The authors used this protocol in Nature Plants Apr 2022
Abstract
Here, we present an approach combining fluorescence in situ hybridization (FISH) and immunolabeling for localization of pri-miRNAs in isolated nuclei of A. thaliana. The presented method utilizes specific DNA oligonucleotide probes, modified by addition of digoxigenin-labeled deoxynucleotides to its 3′ hydroxyl terminus by terminal deoxynucleotidyl transferase (TdT). The probes are then detected by immunolabeling of digoxigenin (DIG) using specific fluorescent-labeled antibodies to visualize hybridized probes. Recently, we have applied this method to localize pri-miRNA156a, pri-miRNA163, pri-miRNA393a, and pri-miRNA414 in the nuclei isolated from leaves of 4-week-old A. thaliana. The present approach can be easily implemented to analyze nuclear distribution of diverse RNA classes, including mRNAs and pri-miRNAs in isolated fixed cells or nuclei from plant.
Keywords: Fluorescent in situ hybridization Immunolabeling Digoxigenin Pri-miRNA Confocal microscopy
Background
Microscopic imaging allows to observe and analyze the localization of diverse RNA molecules in their native cellular environment. It was found to be essential for understanding their turnover and function in different types of cells. Conventional fluorescence in situ hybridization (FISH) techniques have been designed for visualization of RNAs and relay on using fluorescently labeled DNA oligonucleotides (Hu et al., 2014). However, FISH methods are often challenging to detect RNAs expressed at low levels in the cells, such as pri-miRNAs, mainly due to low efficiency of resulting fluorescence signal. Here, we describe a method to detect pri-miRNA molecules in isolated nuclei of A. thaliana by combining FISH together with immunolocalization of digoxigenin (Figure 1). This method has been shown to be more sensitive and effective than conventional FISH (Bhat et al., 2020; Gonzalo et al., 2022; Stepien et al., 2022), and can successfully be applied to detect pri-miRNAs in plant and animal cells.
Figure 1. Overview of combined fluorescence in situ hybridization (FISH) and immunolabeling approach to detect pri-miRNAs in situ. Prior to the FISH, DNA oligonucleotides were labeled with digoxigenin at their 3′-ends by terminal transferase (TdT) (1), and the nuclei were isolated following the procedure described in (2). Pri-miRNAs were next localized by applying FISH (3) combined with the immunolocalization of digoxigenin attached to 3′-ends of the probes (4 and 5). The results were registered with confocal microscope (6). DIG: digoxigenin, TdT: terminal transferase, pAb: primary antibody, sAb: secondary antibody
Materials and reagents
Reagents
Roche Terminal Transferase 8000 U (Merck, catalog number: 3333566001)
ChromaTideTM Alexa FluorTM 488-5-dUTP (Thermo Fisher, catalog number: C11397)
Roche DIUTPS-RO (DIG-11-dUTP) (Digoxigenin-11-dUTP, alkali-stable) (Merck, catalog number: 11093088910)
Hoechst 33342, trihydrochloride, trihydrate, 100 mg (Thermo Fisher, catalog number: H1399)
ProLongTM Gold Antifade Mountant (Thermo Fisher, catalog number: P36930)
Paraformaldehyde, 16% w/v aq. soln., methanol free (Thermo Fisher, catalog number: 043368.9M)
Formamide (Merck, catalog number: FX0420)
Sodium chloride (Merck, catalog number: 1.16224)
Potassium chloride (Merck, catalog number: 7447-40-7)
Magnesium chloride (Merck, catalog number: 7786-30-3)
Sodium citrate (Merck, catalog number: 1613859)
Sodium phosphate, 0.5 M, pH 7.0 buffer (Thermo Fisher, catalog number: J63791.AK)
Saline sodium citrate (SSC) 20× concentrate (Merck, catalog number: S6639)
Ethylenediaminetetraacetic acid (EDTA) (Merck, catalog number: E9884-100G)
Bovine serum albumin (BSA) (Merck, catalog number: A9418-5G)
Ficoll® 400 (Merck, catalog number: F2637-5G)
Polyvinylpyrrolidone (Merck, catalog number: P0930-50G)
BSA, acetylated (Thermo Fisher, catalog number: AM2614)
Spermine (Merck, catalog number: S3256-1G)
Anti-digoxigenin antibody (Merck, 11333062910)
Goat anti-mouse secondary antibody, Alexa Fluor Plus 488 (Thermo Fisher, catalog number: A55058)
Goat anti-mouse secondary antibody, Alexa Fluor Plus 555 (Thermo Fisher, catalog number: A32727)
RNase A (Thermo Fisher, catalog number: EN0531)
Tris-HCl at pH 7.5 (Thermo Fisher, catalog number: 15567027)
Tween 20 (Merck, catalog number: P1379)
Sucrose (Merck, catalog number: S0389)
Phosphate buffered saline (PBS) (Merck, catalog number: P4417)
Triton X-100 (Merck, catalog number: 9036-19-5
Solutions
Hybridization buffer (see Recipes)
Denhardt’s solution 100× (50 mL) (see Recipes)
LB01 buffer (10 mL) (see Recipes)
Sorting buffer (10 mL) (see Recipes)
Equipment
Humid chamber (e.g., StainTray Slide Staining, Polysciences, catalog number: 25502-1)
Polylysine adhesion slides (e.g., Roth, catalog number: ET10.1)
Cover glasses for microscope slides (e.g., Merck, Brand cover glasses, catalog number: BR470045)
CellTrics 30 µm (Sysmex, catalog number: CTS220112)
Confocal microscope, Leica DMI 6000B, TCS TrinoTCS SP8
Plan Apochromat DIC H oil immersion lens (63×/1.4)
Software
ImageJ, Fiji (https://imagej.net/software/fiji/)
Procedure
Nuclei isolation
Note: Nuclei isolation was performed according to the method described in Pontvianne et al. (2007 and 2012).
Collect 2–3 leaves from 4-week-old A. thaliana plants and fix them for 20 min at room temperature in 4% formaldehyde solution in Tris buffer (10 mM Tris-HCl at pH 7.5, 10 mM EDTA, 100 mM NaCl). Leaves should be completely immersed.
Notes:
For 4% fixative preparation, dilute 10 mL of 16% methanol-free formaldehyde in 30 mL of Tris buffer.
Nuclei can also be isolated from seedlings, following the same protocol.
WARNING: Formaldehyde is a volatile and toxic carcinogen and should only be used in a fume hood. Gloves, safety glasses, and protective clothing should also be worn when working with it.
Wash the leaves twice for 10 min in Tris buffer.
Chop the leaves with a razor blade in 0.5 mL of LB01 buffer to obtain a soup in a Petri dish.
Filter the resulting lysate through a 30 μm cell strainer.
Add 12 μL of sorting buffer per 3 μL of filtered nuclei suspension obtained in step 3.
Spread the mixture on adhesion slides covered with Polylysine using a pipette and leave to dry at room temperature (can be kept overnight).
Probe preparation
Dissolve DNA oligonucleotides with double-distilled H2O to a final concentration of 100 μM.
Prepare terminal transferase (TdT) working solution: 5 mM CoCl2, 1× TdT Reaction buffer, 0.1 mM DIG-11-dUTP, 0.1 mM dATP, and 400 U of TdT per reaction in PCR tube.
Notes:
Optionally, you can supply a reaction buffer additionally with Alexa Fluor 5-dUTP (final concentration: 0.2 mM).
Instead of DIG-11-dUTP, Biotin-11-dUTP could be used.
Combine 1 μL of DNA oligonucleotide with 9 μL of TdT working solution and incubate it for 40 min at 37 °C in a PCR tube.
Notes:
Incubation is performed without shaking. After incubation, the reaction mixture could be stored at -20 °C. In our studies, probes were working after three months of storage.
For our studies, the probes were provided by Metabion company (Planegg, Germany).
Fluorescence in situ hybridization (FISH)
Prepare permeabilization buffer: 0.1% Triton X-100 in 1× PBS.
Incubate the sample in permeabilization buffer for 10 min at room temperature. The buffer should cover the whole sample area.
Wash the sample twice with 1× PBS.
Prepare hybridization buffer (Niedojadlo et al., 2012).
For FISH, dilute 1 μL of the probe mixture (section B) in 100 μL of hybridization buffer (final probe concentration: 100 pM/μL).
Note: The final probe concentration in hybridization buffer may vary for different RNAs and should be adjusted experimentally.
Add 100 μL of hybridization buffer to the sample on the slide and cover with a slide cover.
Place the slide in a humid hybridization chamber in the 28 °C incubator and leave overnight. The slides should be prevented from drying during hybridization. To obtain this, a small volume of PBS or water should be placed inside the chamber to keep a stable humidity.
Note: Incubation is performed with a gentle shaking (0.6 rpm)
After incubation, wash the slide using the following order: 4× SSC for 5 min, 2× SSC for 5 min, 1× SSC for 5 min, and 1× PBS for 10 min.
Immunolabeling
Dilute primary anti-digoxigenin antibody in 1× PBS containing 0.05% acetylated BSA diluted 1:100.
Cover the sample with 100 μL of antibody diluent (containing anti-digoxigenin antibody from step 1) and place the slide cover. Incubate the sample in a humid chamber overnight at 10 °C.
Remove primary antibody from the sample and wash with 1× PBS for 5 min.
Dilute the secondary antibody labeled with fluorescence dye in 1× PBS containing 0.05% acetylated BSA (diluted 1:1,000).
Place 100 μL of secondary antibody solution on the sample, cover it with a slide cover glass, and incubate in a humid chamber for 3 h at 37 °C.
Remove the secondary antibody from the sample and wash it two times with 1× PBS for 5 min.
Note: The time and temperature of incubation, as well as antibody concentration, should be defined experimentally for different antibodies used.
Nuclei labeling and sample mounting
Dilute Hoechst 33342 at 1:1,000 in 1× PBS and add 100–200 μL to the sample. Incubate for 10 min at room temperature.
Note: For nuclei labeling, DAPI (4′,6-diamidino-2-phenylindole) diluted 1:1,000 in 1× PBS buffer can also be used.
Remove the Hoechst 33342 solution from the slide and wash it once with 1× PBS for 5 min.
Cover the sample with 20 μL of ProLong Gold Antifade reagent, carefully place the coverslip, and press down firmly with the finger to remove air bubbles and uniformly distribute the antifade reagent over the samples.
Leave the sample to dry at room temperature, protecting it from the light.
Data analysis
In our studies, the results were registered with the Leica SP8 confocal microscope using a diode 405 laser, an argon/ion laser with a wavelength of 488 nm, and a diode laser DPSS 561 that emitted light with a wavelength of 561 nm with an optimized pinhole, long exposure time (200 kHz) and 63× (numerical aperture, 1.4). Images were collected sequentially in blue (Hoechst 33342), green (Alexa 488 fluorescence), and red (Alexa 555) channels. The low laser power (0.4%–5% of maximum power) and a single-channel collection were set to minimize bleed-through between fluorescence channels. For bleed-through analysis, Leica SP8 software was used.
The presented approach was previously applied to localize pri-miRNA156a and pri-miRNA163 in the nuclei isolated from leaves of 4-week-old A. thaliana (Figure 2).
Figure 2. Detection of pri-miR156a and pri-miR163 using digoxigenin-labeled probes and antibodies targeting digoxigenin. The probes hybridizing to intron sequence located downstream of stem-loop structure were applied. (A) Schematic illustration of stem-loop structure of pri-miRNA; probe is depicted in red. (B) Detection of pri-miRNA156a (left image, green) and pri-miRNA163 (right image, green). DNA was stained with Hoechst (blue). Scale bar = 2.5 μm.
To confirm that the observed nuclear spots were due to hybridization of probes to pri-miRNA, we treated the isolated nuclei with RNase A (100 μg/mL in PBS for 30 min) and then applied the probe recognizing the intronic sequence of pri-miR156a (Figure 3B). We have not observed hybridization signals in nuclei treated with RNase A. Similarly, we have not observed fluorescence signal if a sense probe matching the pri-miR156a intron was applied (Figure 3C). Additionally, we have calculated a fluorescence intensity by Fiji software to distinguish the positive fluorescence signal of hybridization from the background.
Figure 3. Detection (fluorescence in situ hybridization, FISH) of pri-miR156a (red) using the intron-recognizing probe (Intron antisense probe), in the nuclei treated with RNase A and using the probe with the sequence identical with a fragment of intron (Intron sense probe). The fluorescence intensity is plotted along the yellow line shown on each merged image. Scale bar = 2.5 μm.
Recipes
Hybridization buffer
Reagent Final concentration Quantity
Formamide 30% (v/v) 600 μL
20× SSC 4× 400 μL
100× Denhardt’s solution 5× Denhardt’s buffer 100 μL
Sodium phosphate buffer (0.5 M) 50 mM 200 μL
EDTA (0.5 M) 1 mM 4 μL
H2O n/a 696 μL
Total n/a 2 mL
Denhardt’s solution 100× (50 mL)
Reagent Final concentration Quantity
Bovine serum albumin (BSA) 2% (w/v) 1 g
Ficoll 400 2% (w/v) 1 g
Polyvinylpyrrolidone 2% (w/v) 1 g
H2O n/a 50 mL
Total n/a 50 mL
LB01 buffer (10 mL)
Reagent Final concentration Quantity
Tris-HCl (1M) 15 mM 150 μL
EDTA 2 mM 20 μL
Spermine 0.5 mM 5 μL
Potassium chloride (KCl) (1M) 80 mM 800 μL
Sodium chloride (NaCl) (1M) 20 mM 200 μL
Triton (10%) 0.1% 1 mL
Sorting buffer (10 mL)
Reagent Final concentration Quantity
Tris-HCl (1 M) 100 mM 1 mL
Potassium chloride (KCl) (1 M) 50 mM 500 μL
Magnesium chloride (MgCl2) (1 M) 2 mM 20 μL
Tween 20 (10%) 0.05 mM 50 μL
Sucrose 5% 0.5 g
Bovine serum albumin (BSA) 3% 0.3 g
Acknowledgments
This work was supported by grants from the Polish National Science Centre (UMO-2019/32/T/NZ1/00508, UMO-2016/23/N/NZ1/00010, UMO-2013/10/A/NZ1/00557) and the Initiative of Excellence–Research University (05/IDUB/2019/94) at Adam Mickiewicz University, Poznan, Poland, to A.J.
Competing interests
There are no conflicts of interest or competing interests.
References
Bhat, S. S., Bielewicz, D., Gulanicz, T., Bodi, Z., Yu, X., Anderson, S. J., Szewc, L., Bajczyk, M., Dolata, J., Grzelak, N., et al. (2020). mRNA adenosine methylase (MTA) deposits m6A on pri-miRNAs to modulate miRNA biogenesis in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 117(35): 21785–21795.
Gonzalo, L., Tossolini, I., Gulanicz, T., Cambiagno, D. A., Kasprowicz-Maluski, A., Smolinski, D. J., Mammarella, M. F., Ariel, F. D., Marquardt, S., Szweykowska-Kulinska, Z., et al. (2022). R-loops at microRNA encoding loci promote co-transcriptional processing of pri-miRNAs in plants. Nat. Plants 8(4): 402–418.
Hu, L., Ru, K., Zhang, L., Huang, Y., Zhu, X., Liu, H., Zetterberg, A., Cheng, T. and Miao, W. (2014). Fluorescence in situ hybridization (FISH): an increasingly demanded tool for biomarker research and personalized medicine. Biomarker Res. 2(1): e1186/2050-7771-2-3.
Niedojadlo, K., Piecinski, S., Smolinski, D. J., Bednarska-Kozakiewicz, E. (2012) Transcriptional activity of Hyacinthus orientalis L. female gametophyte cells before and after fertilization. Planta 236: 153–169.
Pontvianne, F., Matía, I., Douet, J., Tourmente, S., Medina, F. J., Echeverria, M. and Sáez-Vásquez, J. (2007). Characterization of AtNUC-L1 Reveals a Central Role of Nucleolin in Nucleolus Organization and Silencing of AtNUC-L2 Gene in Arabidopsis. Mol. Biol. Cell 18(2): 369–379.
Pontvianne, F., Blevins, T., Chandrasekhara, C., Feng, W., Stroud, H., Jacobsen, S. E., Michaels, S. D. and Pikaard, C. S. (2012). Histone methyltransferases regulating rRNA gene dose and dosage control in Arabidopsis. Genes Dev. 26(9): 945–957.
Stepien, A., Dolata, J., Gulanicz, T., Bielewicz, D., Bajczyk, M., Smolinski, D. J., Szweykowska-Kulinska, Z. and Jarmolowski, A. (2022). Chromatin-associated microprocessor assembly is regulated by the U1 snRNP auxiliary protein PRP40. Plant Cell 34(12): 4920–4935.
Article Information
Copyright
© 2023 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 > RNA > RNA detection
Plant Science > Plant molecular biology > RNA
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This protocol has been corrected. See the correction notice.
Peer-reviewed
Isolation of Epithelial and Stromal Cells from Colon Tissues in Homeostasis and Under Inflammatory Conditions
CM Clara Morral *§
RG Reem Ghinnagow *
TK Tatiana Karakasheva
YZ Yusen Zhou
AT Anusha Thadi
NL Ning Li
BY Benjamin Yoshor
GS Gloria E. Soto
CC Chia-Hui Chen
DA Daniel Aleynick
SW Sarah Weinbrom
MF MaryKate Fulton
YU Yasin Uzun
MB Meenakshi Bewtra
JK Judith R. Kelsen
CL Christopher J. Lengner
KT Kai Tan
AM Andy J. Minn
KH Kathryn E. Hamilton
(*contributed equally to this work, § Technical contact)
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4825 Views: 2442
Reviewed by: Kazem NouriArundhati Mehta Anonymous reviewer(s)
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Abstract
Inflammation of the gastrointestinal tract is a prevalent pathology in diseases such as inflammatory bowel disease (IBD). Currently, there are no therapies to prevent IBD, and available therapies to treat IBD are often sub-optimal. Thus, an unmet need exists to better understand the molecular mechanisms underlying intestinal tissue responses to damage and regeneration. The recent development of single-cell RNA (sc-RNA) sequencing-based techniques offers a unique opportunity to shed light on novel signaling pathways and cellular states that govern tissue adaptation or maladaptation across a broad spectrum of diseases. These approaches require the isolation of high-quality cells from tissues for downstream transcriptomic analyses. In the context of intestinal biology, there is a lack of protocols that ensure the isolation of epithelial and non-epithelial compartments simultaneously with high-quality yield. Here, we report two protocols for the isolation of epithelial and stromal cells from mouse and human colon tissues under inflammatory conditions. Specifically, we tested the feasibility of the protocols in a mouse model of dextran sodium sulfate (DSS)-induced colitis and in human biopsies from Crohn’s patients. We performed sc-RNA sequencing analysis and demonstrated that the protocol preserves most of the epithelial and stromal cell types found in the colon. Moreover, the protocol is suitable for immunofluorescence staining of surface markers for epithelial, stromal, and immune cell lineages for flow cytometry analyses. This optimized protocol will provide a new resource for scientists to study complex tissues such as the colon in the context of tissue damage and regeneration.
Key features
• This protocol allows the isolation of epithelial and stromal cells from colon tissues.
• The protocol has been optimized for tissues under inflammatory conditions with compromised cell viability.
• This protocol is suitable for experimental mouse models of colon inflammation and human biopsies.
Graphical overview
Graphical representation of the main steps for the processing of colon tissue from dextran sodium sulfate (DSS)-treated mice (upper panel) and frozen biopsies from Crohn’s patients (lower panel)
Keywords: Colon Epithelium Lamina propria Stem cells Inflammatory bowel disease (IDB)
Background
Inflammatory bowel disease (IBD) is a chronic inflammatory condition of the gastrointestinal (GI) tract that encompasses two distinct pathologies: ulcerative colitis and Crohn’s disease. The global prevalence of IBD has been increasing since 2000 and now affects approximately 1.4 million individuals in America and 2.3 million in Europe (Ng et al., 2017). Current medical treatment for IBD involves the use of immunomodulatory agents to empirically dampen inflammation; however, up to one-third of patients fail initial treatment, and half of the initial responders will eventually relapse.
The interplay between the epithelial and stromal compartments supports homeostasis of the GI tract. Epithelial cells form a dynamic cellular layer with high regeneration potential that separates the luminal content from the underlying stromal tissue while supporting nutrient and water absorption. Simultaneously, the resident stromal cells of the lamina propria provide structural support, immunity against pathogens, and tolerance to the commensal microbiota. Breakdown of this equilibrium by dysregulation of host–microbiome interactions, defects in intestinal epithelial barrier permeability, and loss of immune tolerance can lead to the development of IBD (Maloy and Powrie, 2011). Therefore, it is important to consider the complex cellular heterogeneity when designing new experimental approaches to better understand the molecular mechanisms underlying disease.
The recent development of single-cell technologies has provided a finer picture of complex biology and deciphered heterogeneity present in tissue, enabling the identification of new cell populations, cellular interactions, and signaling pathways associated with pathology. However, these powerful approaches require the isolation of high-quality single cells from tissues to provide a representative and reliable tissue-associated cellular landscape. While the field of single-cell technology is moving fast, scientists have had to adapt tissue dissociation protocols depending on the cell types and contexts of interest. In the context of intestinal biology, the complex cellular heterogeneity of the gut mucosa and sensitivity to detachment-mediated cell death has hindered protocols for tissue dissociation that aim to represent the cellular repertoire of the original tissue. The main limitation is the fragility of the epithelial cells that cannot support most enzymes used to dissociate tissues. This becomes even more challenging when working with tissues under inflammatory conditions where cells have been exposed to a harmful and often cytotoxic environment. Some studies have already developed protocols to obtain epithelial and stromal cells from the same tissues (Smillie et al., 2019; Elmentaite et al., 2020); however, there is variability across different studies and laboratories. Our goal was to establish a detailed and easy-to-follow protocol to isolate simultaneously epithelial and stromal cells from mouse and human colon samples under inflammatory conditions. For the mouse protocol, we used the common dextran sodium sulfate (DSS) model of intestinal damage; for the human, we optimized the protocol for processing cryo-preserved colon biopsies from patients with Crohn’s disease. We validated the protocols using downstream flow cytometry and single-cell RNA sequencing analyses and demonstrated that both recover the main cell populations found in the gut mucosa.
Protocol for processing colon tissues from DSS-treated mice
Materials and reagents
Biological materials
7-week-old female CL57BL/6 mice (Charles River Laboratory) were treated with 3% dextran sodium sulfate (DSS) in water for five consecutive days. Colon tissues were isolated on day 3 after treatment at the peak of the inflammatory response.
Reagents
RPMI (Invitrogen, catalog number: 11875119)
Advanced DMEM/F-12 (Gibco, catalog number: 12-634-028)
HBSS (Cytiva, catalog number: SH3058801)
HEPES (Invitrogen, catalog number: 15630080)
GlutaMAX (Thermo Fisher Scientific, catalog number: 35050061)
Pen/Strep (Invitrogen, catalog number: 15140122)
EDTA (0.5 M, pH 8.0) (Invitrogen, catalog number: AM9260G)
DNase I (Roche, catalog number: 10104159001)
Liberase TM (Sigma, catalog number: 05401127001)
TrypLE (Invitrogen, catalog number: 12605010)
Dextran sodium sulfate (DSS) (Alfa Aesar, catalog number: J62101-22)
Trypan Blue solution (Gibco, catalog number: 15250061)
Flow cytometry antibodies:
LIVE/DEADTM Fixable Aqua Dead Cell Stain kit (Thermo Fisher Scientific, catalog number: L34957)
BV421 Rat anti-mouse CD31 (BD Horizon, catalog number: 562939, clone: MEC 13.3)
APC Anti-mouse CD326 (EP-CAM) (BioLegend, catalog number: 118214, clone: G8.8)
PE Anti-mouse Podoplanin (BioLegend, catalog number: 127408, clone: 8.1.1)
FITC Rat anti-mouse CD45 (BD Pharmingen, catalog number: 553080, clone: 30-F11)
Note: Depending on the flow cytometer available, users can use a different color panel design.
Solutions
Epithelial cell solution (see Recipes)
Lamina propria cell solution (see Recipes)
Epithelial wash buffer (see Recipes)
Recipes
Epithelial cell solution
Prepare it fresh and keep it on ice.
Reagent Final concentration Quantity (for one sample)
HBSS n/a 11,250 μL
HEPES 10 mM 150 μL
EDTA (0.5 M) 10 mM 300 μL
Pen/Strep 100 U/mL 150 μL
FBS 2% 300 μL
DNase I 100 μg/mL 150 μL
Total n/a 15 mL
Lamina propria solution
Prepare it fresh and keep it on ice.
Reagent Final concentration Quantity (for one sample)
RPMI n/a 9,600 μL
Pen/Strep 100 U/mL 100 μL
Liberase TM 100 μg/mL 200 μL
DNase I 100 μg/mL 100 μL
Total n/a 10 mL
Epithelial wash buffer
Once prepared can be stored at 4 °C for one month.
Reagent Final concentration Quantity (for one sample)
Adv-DMEM F12 n/a 4,900 μL
HEPES 10 mM 500 μL
GlutaMAX 10 mM 500 μL
Total n/a 50 mL
Laboratory supplies
15 mL centrifuge tube (CELLTREAT, catalog number: 30411B)
50 mL centrifuge tube (CELLTREAT, catalog number: 229430)
100 mm × 15 mm Petri dishes (CELLTREAT, catalog number: 229693)
100 μm cell strainer (Falcon, catalog number: 352360)
40 μm cell strainer (Falcon, catalog number: 352340)
Gavage needle (Gavage Needle, catalog number: AFN2025S)
Frosted microscope slides (Fisher Scientific, catalog number: 12-550-343)
5 mL Polystyrene round-bottom tube with cell-strainer cap (Falcon, catalog number: 352235)
Filter pipette tips (1,000 μL) (Genesee Scientific, catalog number: 24-430)
Serological pipets (10 and 5 mL) (Genesee Scientific, catalog number: 12-102, 12-104)
Equipment
Surgical scissors
Forceps
Centrifuge 5810 R (Eppendorf)
Countess 3 (Invitrogen)
Water bath (Fisher Scientific, Isotemp GPD 10)
Software and datasets
FlowJo v10.8.1
GraphPad Prism9
Procedure
Harvesting colon tissue and cleaning
Cut the colon from the gastrointestinal tract.
Note: Colons from DSS-treated mice tend to be shorter compared to control mice, as a result of active inflammation (Figure 1).
Figure 1. Colon tissues isolated from control or DSS- treated mice
Place the colon tissue in a Petri dish containing ice-cold HBSS.
Clean the colon tissue by flushing it with HBSS using a 10 mL syringe with a gavage needle.
Cut the tissue and open it longitudinally. Wash it in the same Petri dish several times by replacing it with fresh ice-cold HBSS.
Once clean, cut the tissue into small pieces (approximately 2 cm long) and keep them with HBSS in the same Petri dish on ice until proceeding to the next step.
Note: Clean and cut all colon tissues required for the experiment before proceeding to the next step.
Isolation of crypts and lamina propria
In this step, the epithelial layer containing the crypts will be isolated from the rest of the tissue.
Transfer the colon pieces into a 50 mL conical tube containing 15 mL of epithelial cell solution.
Incubate at 37 °C in a water bath for 15 min and manually shake the tubes gently every 5 min.
After incubation, transfer the tubes to ice for 10 min.
Transfer the tissue to a new 50 mL conical tube containing 15 mL of fresh HBSS.
Note: It is important to proceed with the crypt isolation (steps B4–B7) using fresh HBSS without EDTA.
Manually shake vigorously for 30–40 s.
Take a small sample of the supernatant, transfer it into a glass slide, and look under the microscope to verify the presence of colon crypts (see example in Figure 2). If crypts are present, proceed to step B7.
Note: If crypts are not present, extend the incubation time in EDTA for 10 more minutes on ice and shake again for 30–40 s.
Figure 2. Brightfield image of colon crypts. (A) Colon crypts obtained after EDTA incubation and three consecutive rounds of shaking. (B) Magnification from (A). Squares highlight colon crypts.
Collect the supernatant by filtering it through a 100 μm strainer into a new 50 mL conical tube and place the tissue back into the same 50 mL conical tube.
Repeat steps B4–B7 two more times to pool three fractions (total volume of 45 mL).
Centrifuge the supernatant containing the three crypt fractions at 400× g for 5 min.
Resuspend the crypt pellet with 20 mL of epithelial wash buffer and keep the samples on ice until the epithelial fraction digestion step.
Keep the remaining tissue (lamina propria) in a 50 mL conical tube containing 20 mL of HBSS. This tissue will be digested in the next step.
Digestion
Lamina propria fraction
After epithelial layer separation, take the rest of the tissue and place it in the 50 mL conical tube containing 20 mL of HBSS.
Wash the tissue by gently shaking it for 30 s. Discard the HBSS and add 20 mL of new HBSS. Repeat this step at least three times.
Note: This step is very important, as any remaining EDTA will block the lamina propria digestion enzyme.
After washing, cut the tissue into small pieces (approximately 0.5 cm long) and transfer them into a new 15 mL conical tube containing 10 mL of lamina propria solution.
Incubate at 37 °C in a water bath for 20 min.
After 20 min, start pipetting up and down with the 5 mL pipette until the tissue passes smoothly through the pipette.
Note: You can start with the 10 mL pipette and then move to the 5 mL.
Incubate at 37 °C for 10 min.
Start pipetting up and down with a 1 mL pipette until the tissue gets completely dissociated.
Transfer samples to ice and add 1 mL of FBS and 80 μL of EDTA to stop the digestion reaction.
Pipette up and down with a 1 mL pipette and filter through a 40 μm strainer into a 50 mL conical tube.
Add HBSS until 30 mL of volume.
Spin down at 400× g for 10 min.
Remove the supernatant and resuspend the cell pellet with 2 mL of RPMI.
Count cells and measure viability using trypan blue and a cell counter.
Epithelial fraction
Spin down the epithelial fraction (containing crypts) at 400× g for 5 min.
Inspect the cell pellet after the spin down.
Note: It is very important to adjust the volume of TrypLE enzyme used depending on the crypt pellet. As a reference, a pellet similar to the one shown in Figure 3A will be resuspended in 5 (left) or 3 mL (right) of TrypLE.
Figure 3. Examples of cell pellets during the epithelial fraction dissociation. (A) Crypt pellet obtained from a control (left) or dextran sodium sulfate (DSS)-treated (right) mouse in 50 mL tubes. (B) Example of a floating cell pellet after digestion with TrypLE in a 15 mL tube.
Resuspend the crypt pellet in 5 mL of pre-warmed TrypLE containing DNase I (100 μg/mL).
Pipette gently up and down until the crypt pellet is dissolved and incubate samples for 5 min at room temperature (RT).
After the 5 min, start pipetting again with a 1 mL pipette for a total period of 10 min.
Note: When there are several samples, keep switching from one to the other every 2–3 min.
Inspect the cells under the microscope to see if they are single cells. If there are still clumps of 5–10 cells, keep pipetting for 5 more minutes.
Note: We do not recommend incubating the epithelial fraction with TrypLE for more than 30 min.
Add 1 mL of FBS into the 5 mL of TrypLE to stop the reaction.
Filter through a 40 μm strainer in a 50 mL conical tube. Add epithelial wash buffer up to 50 mL.
Spin down cells at 400× g for 10 min at 4 °C. Critical point: If the cell pellet is not visible or is floating (see example Figure 3B), after this step we recommend increasing the time and the speed of the centrifugation. We have seen that 10 min at 800× g does not compromise cell viability and helps to pellet the cells.
Discard the supernatant by pipetting. Caution: The pellet can easily detach from the bottom of the tube. It is very important to remove the supernatant carefully. If required, leave 1 or 2 mL of volume to avoid losing cells.
Resuspend the cell pellet in 2 mL of epithelial wash buffer.
Count cells and measure viability using trypan blue and a cell counter.
Validation of protocol
Cell viability
Here, we provide the cell viability and cell count of epithelial and stromal cells obtained at the end of the protocol.
Cell count and viability were obtained using trypan blue and an automated cell counter and listed in Table 1.
Table 1. Cell count and viability at end of protocol.
Mouse Cell count (total) Viability
Epithelium Lamina propria Epithelium Lamina propria
Mouse 1_Untreated 520 K 14.1 M 83% 98%
Mouse 2_Untreated 540 K 12.2 M 84% 95%
Mouse 3_DSS 396 K 17.8 M 93% 98%
Mouse 4_DSS 340 K 15 M 93% 96%
Flow cytometry
By flow cytometry, we characterized different cell types within the epithelial and lamina propria fractions harvested from the colon of naive and DSS-treated mice. As seen in Figure 4A and 4C (top), from viable cells, over 80% of cells are epithelial cells (EPCAM+) in the epithelial fraction from naïve mice (left panel), while in DSS-treated mice (right panel), higher CD45+ immune cells are observed along with a decrease in EPCAM+ cells reflective of active inflammation. We also studied the potential presence of endothelial cells and fibroblasts within the epithelial fraction by gating on CD31 and Podoplanin (PDPN), respectively, in stromal cells identified as CD45 and EPCAM double negative. We noticed that in the epithelial fraction, there is no contamination by CD31+ or PDPN+ cells. In the lamina propria fraction (Figure 4B and 4C, bottom), we observed that most cells are stroma (mainly fibroblasts and, to a lesser extent, endothelial cells) in both naïve and DSS mice with higher immune cell infiltration in the last group.
Figure 4. Flow cytometry gating strategy in the colon. (A and B) Representative flow plot and (C) quantification of cellular subsets (pre-gated on live cells) in the epithelial (A) and lamina propria fractions (B) of naive (left panel) compared to dextran sodium sulfate (DSS)-treated mice (right panel). Data are shown as mean ± SD for three mice in each group. CD45+ = immune cells, EPCAM+ = epithelial cells, CD31+ = endothelial cells and PDPN+ = fibroblast.
Protocol for processing colon biopsies from patients with Crohn’s disease
Materials and reagents
Biological materials
This protocol works equally well for both fresh and cryopreserved biopsies, but for consistency and process efficiency, we routinely use cryopreserved tissue (see General notes). We routinely process two biopsies per subject, to have sufficient cell counts for both sequencing and colonoid line establishment. Biopsies were collected from pediatric patients aged 0–17 years (Table 2) undergoing endoscopy at the Children’s Hospital of Philadelphia between January and December 2021. Prior to sample collection, informed consent was obtained in accordance with Institutional Review Board guidelines (protocol 16-013042). Research biopsies were collected by a trained gastroenterologist from the ascending colon in addition to those for clinical care. Patients with a history of bone marrow transplantation or prior diagnosis of any hemorrhagic, coagulopathic, or aggregation disorders were excluded from this study. Expert pathologists confirmed the diagnoses. Control subjects were evaluated for gastrointestinal complaints but were found to have normal endoscopic and histologic findings.
Table 2. Biopsy donor data
ID Age Sex Race Ethnicity
CTR-1 9 Male White Not Hispanic/Latino
CTR-2 8 Male White Not Hispanic/Latino
CTR-3 8 Female White Not Hispanic/Latina
Crohn’s-1 7 Female Asian Not Hispanic/Latina
Crohn’s-2 9 Male White Not Hispanic/Latino
Crohn’s-3 8 Female White Not Hispanic/Latina
Reagents
CryoStor (CS-10, STEMCELL Technologies, catalog number: 07930)
DPBS (Gibco, catalog number: 14190-136)
Sorbitol (Fisher Bioreagents, catalog number: BP439-500)
Sucrose (Sigma, catalog number: S8501)
BSA (Fisher Bioreagents, catalog number: BP1600-100)
Gentamicin (Gibco, catalog number: 15750-060)
DMEM, high glucose (Corning, catalog number: 10-013-CV)
Advanced DMEM/F12 (Thermo Fisher Scientific, catalog number: 12634028)
FBS (Peak Serum, catalog number: PS-FB1)
EDTA (Quality Biological, catalog number: 351-027-721)
HBSS (Gibco, catalog number: 14175-079)
Antibiotic-Antimycotic (Gibco, catalog number: 15240-062)
HEPES (Gibco, catalog number: 15630-080)
GlutaMAX Supplement (Gibco, catalog number: 35050-061)
Liberase TH (Sigma, catalog number: 5401135001)
DNase I (Roche, catalog number: 10104159001)
IntestiCult organoid growth medium, human (STEMCELL Technologies, catalog number: 06010)
TrypLE Express (Gibco, catalog number: 12605-010)
Trypan Blue solution (Gibco, catalog number: 15250061)
Dead cell removal kit (Miltenyi Biotech, catalog number: 130-090-101) and necessary accessories (see Laboratory supplies)
10× Chromium Next GEM Single Cell 3’ v3.1
Solutions
Liberase TH stock is 13 Wunsch Units (WU)/mL (2.5 mg collagenase/mL) in HBSS. Aliquots are stored at -20 °C, and freeze/thaw is limited to two cycles
DNase I stock is 50 U/mL (25 mg/mL) in DPBS. Aliquots are stored at -20 °C, and freeze/thaw is limited to five cycles
HBSS-BSA (1% BSA, sterilized by filtration)
Biopsy collection medium (see Recipes)
Chelation buffer; prepare fresh every time (see Recipes)
Transport medium (see Recipes)
Thaw medium (see Recipes)
Stroma dissociation enzyme (see Recipes)
Epithelium dissociation enzyme (see Recipes)
Recipes
Biopsy collection medium
Reagent Final concentration Quantity
Advanced DMEM/F12 1× 500 mL
Antibiotic-antimycotic 1× 5 mL
HEPES 10 mM 5 mL
GlutaMAX 1× 5 mL
Total n/a 515 mL
Chelation buffer (sterilize by filtration and chill on ice)
Reagent Final concentration Quantity
Sorbitol 2% 0.5 g
Sucrose 1% 0.25 g
BSA 1% 0.25 g
Gentamicin 1% 25 μL
DPBS 1× 25 mL
Total n/a 25 mL
Transport medium
Reagent Final concentration Quantity
DMEM 1× 500 mL
Antibiotic-Antimycotic 1× 5 mL
HEPES 10 mM 5 mL
Total n/a 510 mL
Thaw medium
Reagent Final concentration Quantity
Transport medium n/a 47.5 mL
FBS 5% 2.5 mL
Total n/a 50 mL
Stroma dissociation enzyme (on ice)
Reagent Final concentration Quantity
HBSS n/a 5 mL
Liberase-TH 0.13 WU/mL 50 μL
DNase I 0.5 U/mL 50 μL
Total n/a 5 mL
Epithelium dissociation enzyme
Reagent Final concentration Quantity
TrypLE 1× 1.5 mL
DNase I 0.5 U/mL 15 μL
Total n/a 1.5 mL
Laboratory supplies
Cryopreservation container (CoolCell, Corning, catalog number: CLS432001 or Mr. Frosty, Fisher Scientific, catalog number: 15-350-50)
Dumont SS forceps, two forceps are needed per specimen (Fine Science Tools, catalog number: 11203-25)
15 and 50 mL conical tubes
35 mm round TC dish (alternatively, a well in a 6-well plate can be used)
40 μm cell strainers
FACS tubes
MACS accessories:
MS Columns (Miltenyi Biotec, catalog number: 130-042-201)
MiniMACS Separator (Miltenyi Biotec, catalog number: 130-042-102) or OctoMACS Separator (Miltenyi Biotec, catalog number: 130-042-109)
Equipment
Tube revolver rotator (Thermo Scientific, catalog number: 88881001)
ThermoMixer (Eppendorf, catalog number: 05-412-501)
Vortex mixer (e.g., Vortex Genie-2, Scientific Industries, catalog number: SI-0236)
Stereomicroscope (Nikon, model: SMZ-U)
Centrifuge supporting 15 and 50 mL conicals
Tabletop microcentrifuge
Hemocytometer for cell number and viability counting
Automated cell counting device (e.g., Invitrogen Countess II)
Procedure
Cryopreservation of biopsies
Two colonic biopsies are collected during colonoscopy into Eppendorf tubes containing 1 mL of biopsy collection medium and delivered to the lab immediately for cryopreservation.
Washing of the tissue fragments (do not use vacuum aspirator):
Note: This is particularly important to prevent contamination if establishing primary culture from the specimen (e.g., patient-derived colonoids).
Remove the biopsy collection medium.
Repeat the following three times: add 1 mL of DPBS, pipette up and down 5–10 times, aspirate the DPBS with a P1000 pipette.
Using a P1000 tip, gently transfer both fragments of tissue into a labeled cryovial containing 1 mL of CryoStor (try to carry over as little PBS as possible) and let the tissue settle to the bottom of the cryovial.
Place the cryovials into the cryopreservation jar.
Place the cryopreservation jar in a -80 °C freezer.
After an overnight incubation at -80 °C, samples should be moved to liquid nitrogen storage.
Thaw the cryopreserved tissue
Thaw the biopsy in a water bath and transfer it to a 1.5 mL Eppendorf tube.
Note: If using a bead bath, place a beaker with pre-heated water into the beads and place the cryovial in the water.
Wash three times with 1 mL of thaw medium.
Here and below, follow this procedure for one wash cycle: add 1 mL of appropriate wash solution (thaw medium here) and pipette up and down 10 times to wash the tissue without letting the fragment enter the pipette tip. Remove the spent wash solution.
Isolation of crypts and lamina propria
Wash twice with 1 mL of chelation buffer (Figure 5B) and discard the spent buffer.
Figure 5. Separation of colonic crypts from lamina propria using forceps. (A) Graphical representation of the crypt scraping process, also remonstrated in Video 1. (B) Colonic biopsy, recovered from cryopreservation and suspended in chelation buffer. (C) Colonic biopsy, after the rotation in chelation buffer and EDTA, post-vortexing and transferred into a 35 mm dish for scraping. (D) View of separated lamina propria (LP) and colonic crypts (c) post-scraping. (E) Lamina propria suspended in DPBS prior to enzymatic digestion.
Video 1. Demonstration of crypt scraping
After this, add 80 μL of 0.5 M EDTA to the remaining ~20 mL of chelation buffer.
Add 1 mL of chelation buffer + EDTA (final concentration 2 mM) to the tissue in a 1.5 mL Eppendorf tube and rotate cold (in a tube revolver placed in a cold room, set to 10 rpm) for 15 min. During this time, label and chill on ice:
35 mm round TC dish for scraping;
15 mL conical for crypt collection;
1.5 mL Eppendorf tube for stroma collection.
Vortex the tissue on maximum speed setting for five cycles: vortex for 30 s followed by 30 s rest on ice.
Note: For some samples, crypts begin separating during vortexing.
Transfer the spent chelation buffer (may contain crypts) into the prepared 15 mL crypt collection tube.
Use 1 mL of fresh chelation buffer to gently transfer the tissue into a prepared 35 mm dish (Figure 5C).
Under a stereomicroscope, position the biopsy fragment with the luminal side (epithelium) facing up and scrape off the crypts (Figure 5A and 5D; Video 1).
Note: If it is hard to see which side is up, the easiest way is to begin scraping. If no crypts separate, that indicates the lumen is facing down; flip the fragment over and try again.
Transfer the crypt suspension from the 35 mm dish into the 15 mL crypt collection tube.
Move the remaining tissue into the original Eppendorf tube and mark it with stroma (S).
Use 1 mL of thaw medium to rinse the remaining crypts off the stroma fragments and transfer the suspension to the 15 mL crypt collection tube (leave as little liquid with the stroma as possible).
Submerge the stroma in 1 mL of DPBS and place it on ice (Figure 5E).
Use 2 mL of thaw medium to rinse the remaining crypts of the 30 mm dish and transfer the suspension to the 15 mL crypt collection tube.
Spin the crypt collection tube at 700× g for 1 min.
Note: Here and below, it is beneficial, but not critical, to keep the centrifuges chilled to 4 °C.
Transfer the pellet to a 1.5 mL Eppendorf tube and wash with extra HBSS (~500 μL).
Reconstitute the crypt in 1 mL of cold IntestiCult (with 0.5 U/mL DNase).
Optional: count the crypts (typical yield is 200–500 crypts per biopsy fragment).
Place the crypt suspension to gently rotate in the cold room while processing the lamina propria.
Digestion
Lamina propria fraction
Prepare the enzyme mix that will be used for three cycles of digestion.
Cycle 1: Add 500 μL of enzyme mix to stroma fragments.
Transfer the tube to ThermoMixer for 10 min (37 °C, 800 rpm).
During this time, prime a 40 μm strainer, placed in a 50 mL conical, with 2 mL of FBS; this will be the collection tube.
Force the supernatant through the strainer into the collection tube.
Cycle 2: Add 500 μL of fresh enzyme mix to the remaining tissue fragments.
Slowly pipette the tissue up and down 5–10 times through a P1000 tip to enhance breakdown.
Transfer the tube to ThermoMixer for 10 min (37 °C, 800 rpm).
Force the supernatant through the strainer into the same collection tube.
Cycle 3: Add 500 μL of fresh enzyme mix to stroma fragments.
Slowly pipette the tissue up and down 5–10 times through a P1000 tip to enhance breakdown.
Transfer the tube to ThermoMixer for 10 min (37 °C, 800 rpm).
Force the suspension through the strainer.
Use a plunger from a 1 mL insulin syringe to force the remaining fragments through the strainer.
Wash the strainer with 4 mL of thaw medium.
Transfer the contents from the collection tube to a new 15 mL conical.
Note: This step is optional, but the cell pellet is easier to see after the spin when in a 15 mL conical.
Spin at 700× g for 3 min.
Prime the FACS tube by forcing 50 μL of FBS through the strainer cap.
Reconstitute the pellet in 0.5 mL of thaw Medium and strain into the primed FACS tube.
Count the cells and viability using a hemocytometer and trypan blue exclusion, either manually or in an automated device.
Epithelial fraction
Spin down the crypt suspension at 700× g for 1 min.
Reconstitute the crypt pellet in 1.5 mL of TrypLE + 15 μL of DNase I.
For efficient mixing, split into 3 × 1.5 mL Eppendorf tubes each containing 500 μL of the crypt+enzyme suspension.
Transfer the tube to ThermoMixer for 30 min (37 °C, 800 rpm).
Prime the FACS tube by forcing 1 mL of FBS through the strainer cap. Critical: large clumps could be formed at the end of digestion.
Strain into the primed FACS tube and wash the strainer with 1 mL of thaw medium.
Spin at 700× g for 3 min and reconstitute in 500 μL of HBSS-BSA supplemented with 5 μL of DNase I.
Count the cells and viability using a hemocytometer and trypan blue exclusion, either manually or in an automated device.
For single-cell RNA sequencing, preparation of single-cell cDNA libraries and next-generation sequencing are carried out as outlined in the 10× Chromium Next GEM Single Cell 3’ v3.1 user manual.
Data analysis
We aligned and quantified the reads using 10× Cellranger 3.1.0 and built a gene expression matrix by combining all the available samples. Low-quality cells expressing less than 1,000 genes and mitochondrial UMI (Unique Molecular Identifier) greater than 30% of the cell total were discarded. Only the genes that are expressed in more than 100 cells were used for analysis. The UMI counts for each gene were normalized by the cell totals followed by log transformation after addition of a pseudonumber (Bolstad, 2023) to avoid negative and undefined log values. Single-cell RNA sequencing data metrics and viability counts are listed in Table 3 and Table 4, respectively. Using the top 2,000 genes with highest expression variation across the cells, we performed Principal Component Analysis (PCA) on the log-normalized data with R preprocessCore package (Bolstad, 2023). Top 10 dimensions of the PCA explaining the greatest variation in the data were used as input for non-linear dimensionality reduction with Uniform Manifold Approximation and Projection (UMAP) (Becht et al., 2018). We ran clustering on the UMAP coordinates using density clustering function implemented in Monocle (Qiu et al., 2017). Finally, we assigned the cell types for clusters based on the expression patterns of the known cell markers curated from the literature (Figure 6).
Table 3. Single cell data metrics
ID Total cells sequenced Cells removed due to: Cells post-filtering
Doublets* < 500 genes* > 30% mitochondrial UMI*
CTR-1 6,998 75 1,712 4,318 1,977
CTR-2 11,880 212 1,306 4,393 5,161
CTR-3 9,035 38 5,536 7,833 1,044
Crohn’s-1 6,905 61 1,524 4,514 1,658
Crohn’s-2 3,997 17 2,518 3,382 513
Crohn’s-3 11,587 211 2,285 4,529 5,267
*The total number of cells removed is not always equal to the sum of columns 3–5, because many cells fit several criteria (e.g., the same cell may have < 500 genes and mitochondrial UMI > 30%)
Table 4. Cell and viability counts
Sample % viability Total live cells
Stroma Epithelium Stroma Epithelium
CTR-1 90 48 620,000 58,500
CTR-2 77 76 328,500 70,000
CTR-3 85 42 267,000 123,000
Crohn’s-1 66 57 126,000 141,000
Crohn’s-2 90 57 114,500 50,000
Crohn’s-3 65 78 264,000 45,000
Figure 6. Single-cell RNA-sequencing analysis of tissue biopsies. (A) Uniform Manifold Approximation and Projection (UMAP) visualization of all cells across the six samples color-coded by cell-type compartment. (B) UMAPs divided by disease status. (C) Visualization of the expression of canonical gene markers for epithelial (EPCAM), stromal (PDGFRA), and immune (PTPRC) cells.
Validation of protocol
Cell viability
Single-cell RNA sequencing data metrics and viability counts are listed in Table 3 and Table 4 above.
Note: If the cell viability is lower than 70%, you can use the Dead Cell Removal kit (following the manual provided).
Unsupervised cell clustering
Uniform Manifold Approximation and Projection (UMAP) visualization of all cells across the six samples, as well as visualization of expression of canonical epithelial, stromal, and immune cells are presented above in Figure 6.
General notes and troubleshooting
General notes
The protocols described above are optimized for colon tissues. However, they can also be used for small intestine tissues with some modifications. For murine small intestine, scraping of the villi can be done prior to EDTA incubation to obtain crypt-enriched samples. For human colonic biopsies, the crypt and villi suspension after scraping can be strained through a 100 µm pore membrane to isolate the crypts, while the villi will not go through.
We cryopreserve biopsies prior to processing. Logistically, since we depend on collecting clinical samples from the endoscopy suite, it is difficult to plan the timing and book equipment for fresh tissue processing. Since processing some specimens fresh and some cryopreserved would become a confounding factor, we cryopreserve all specimens and perform the cell isolation protocol in batches.
It is critical that biopsies are cryopreserved as soon as possible, and within 30 min from being procured from a subject.
Mixing back the epithelial cells with stromal cells for single-cell cDNA library generation results in reduced yield of transcriptomes from viable epithelial cells. Therefore, the two fractions must be sequenced separately. To optimize the inevitable cost increase, we sequence biopsies from three subjects at a time: each epithelial fraction results in an independent library, but the stromal fractions can be multiplexed, so there is only one additional library.
Centrifugation steps in the human protocol are short (30 s to 3 min) and thus can be carried out at RT with the sample remaining cool. However, if the centrifuge can be kept at 4 °C, that would further ensure that the sample remains cold.
Troubleshooting
Crypts sticking to forceps: make sure to always dampen the forceps in chelation buffer before touching the tissue.
Poor crypt separation: we find that EDTA stock is the likeliest culprit; opening a new bottle resolves the issue in most cases. Prepare the Chelation buffer fresh every time.
Cell clumps instead of single cells: refresh the enzymes.
Acknowledgments
This publication is part of the Gut Cell Atlas Crohn’s Disease Consortium funded by the Leona M. and Harry B. Helmsley Charitable Trust and is supported by a grant from Helmsley to the Children’s Hospital of Philadelphia. www.helmsleytrust.org/gut-cell-atlas/.
This work is also supported by NIH R01-DK124369 (K.E.H.) and the Children’s Hospital of Philadelphia Gastrointestinal Epithelium Modeling Program.
These protocols were inspired by independent protocols developed by Mahe et al. (2015), Parikh et al. (2019), and Smillie et al. (2019).
Ethical considerations
Animal samples: All animal experiments were performed according to protocols approved by the Institutional Animal Care and Use Committee of the University of Pennsylvania.
Human samples: The institutional review board at the Children’s Hospital of Philadelphia approved the protocol (2014-010826), and all parents of patients provided written informed consent.
References
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Article Information
Copyright
© 2023 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
Cell Biology > Single cell analysis > Flow cytometry
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4,826 | https://bio-protocol.org/en/bpdetail?id=4826&type=0 | # Bio-Protocol Content
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Peer-reviewed
Intraperitoneal Injection of Neonatal Mice
AP Amanda M. Pocratsky *
JS James N. Sleigh *
(*contributed equally to this work)
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4826 Views: 1977
Reviewed by: Vivien J. Coulson-ThomasSudhir Verma Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in JCI Insight Mar 2023
Abstract
Administration of substances into neonatal mice is required for early treatment with pre-clinical therapeutics, delivery of recombination-inducing substances, and dosing with viruses or toxins, amongst other things. Several injection routes into mouse pups are possible, including intravenous and intracerebroventricular, each with their own advantages and limitations. Here, we describe a simple and rapid protocol for the intraperitoneal injection of neonatal mice for systemic dosing. By detaching a 30-gauge needle from its plastic hub and inserting it into polyethylene tubing attached to a Hamilton syringe, small volumes (1–10 μL) can be accurately injected into the peritoneal cavity of pups aged 1–5 days old. The procedure can be completed within a few minutes, is generally safe and well tolerated by both pups and parents, and can be used in combination with alternative administration routes.
Key features
• This protocol provides a simple description to rapidly and efficiently inject mouse pups aged 1–5 days for systemic dosing.
• Allows treatment of neonatal mice with substances such as viruses and compounds for research across disciplines.
Graphical overview
Keywords: Adeno-associated virus (AAV) Gene therapy I.P. administration Lentivirus (LV) Mouse pup Neonate Systemic delivery Tamoxifen
Background
The early post-natal injection of substances into rodents is required for several purposes, including the testing of pre-clinical therapeutics in disease models (Hua et al., 2011), delivering recombination-inducing substances such as tamoxifen (Pitulescu et al., 2010), and performing mechanistic studies with viruses or toxins (MacDonald et al., 2021). Several delivery routes into neonates are available, each with advantages and pitfalls. For instance, injection into the cerebral lateral ventricles (i.e., intracerebroventricular) can bypass the blood–brain barrier and deliver substances throughout the central nervous system via the cerebrospinal fluid, but this injection paradigm will not result in systemic delivery, is not suitable for repetitive treatments, and is technically challenging (Passini and Wolfe 2001). Alternatively, intravenous injections permit greater volumes (~50–100 μL) and result in a more systemic effect; however, this route is tricky to master (Sands and Barker 1999; Kienstra et al., 2007) and becomes more difficult in pups older than two days due to vein visualisation issues (Gombash Lampe et al., 2014). Subcutaneous injections are straightforward and enable larger dosing volumes if needed, but rates of substance absorption are slower compared to other delivery routes (Turner et al., 2011). Intraperitoneal neonatal injections also allow body-wide substance administration via transfer to the circulatory system (thus rapid absorption) and require less expertise and technical ability but are limited by injection volume and are not frequently used in a clinical setting (Al Shoyaib et al., 2019).
Nevertheless, different injection routes into neonatal rodents can result in distinct tissue targeting (Ohshima et al., 2015) and varied temporal pharmacodynamics post-injection (Statler et al., 2007; Foust et al., 2008); Therefore, it is important to master a variety of administration strategies. Indeed, depending on the dosing and targeting requirements, combining injection paradigms in pups is possible and can provide a superior therapeutic effect compared to single-route injections (Nizzardo et al., 2014).
Several published methods for the delivery of substances into neonates via intravenous (temporal/facial vein, jugular vein, or retro-orbitally) and intracerebroventricular routes are available (Sands and Barker 1999; Kienstra et al., 2007; Glascock et al., 2011; Yardeni et al., 2011; Gombash Lampe et al., 2014). However, a similar, detailed protocol for the intraperitoneal injection of mouse neonates is lacking, although approaches have been presented in brief (Ostermann et al., 2013; Xu et al., 2018; MacDonald et al., 2021).
Here, we describe a stepwise method for accurate injections into the peritoneal cavity of mouse pups, which has been used to successfully deliver adeno-associated viruses to drive transgene expression in 1–3-day-old mice (MacDonald et al., 2021; Sleigh et al., 2023). Intraperitoneal injections are highly reproducible and easy to master and can be used for repeated treatments. Success rates will depend on experimenter experience and the substances being injected, but our recent experiments suggest that a near 100% success rate is achievable.
Materials and reagents
Note: Similar materials, reagents, and equipment can be purchased from alternative sources. We note that alternative sourcing of 0.28 mm internal diameter tubing may lead to issues in creating a watertight seal with the 30 G needle (e.g., variance in actual vs. reported internal diameter).
Cannula fabrication
Disposable surgical scalpel (Swan Morton, VWR International, catalog number: 0505)
PE-10/10 Polyethylene tubing, 0.28 mm internal diameter, 0.61 mm external diameter (Warner Instruments, Multi-Channel Systems, catalog number: 64-0750)
30 G × 1/2" needle (BD, catalog number: 305106)
6-well plate, flat bottom (VWR International, catalog number: 734-2777)
Sterile water
10 mL syringe (Fisherbrand, Fisher Scientific, catalog number: 15879152)
10 μL syringe, 701 N, 26 G, 51 mm length, 0.47 mm external diameter (Hamilton, VWR International, catalog number: 549-1135)
Intraperitoneal injection
Disposable surgical drape, e.g., 30 cm × 45 cm (Millpledge, catalog number: SDT0100)
Large weigh boat (Fisherbrand, Fisher Scientific, catalog number: 15758187)
Biohazard sharps bin
Cannula (see above)
Substance(s) for injection (e.g., virus, toxin, compound) in 1.5 mL microfuge tube(s) on ice or disposable heating pad, as needed
Sterile cotton swab (Texwipe, Fisher Scientific, catalog number: 15823886)
Marker pen
P20 pipette tips
Parafilm (VWR International, catalog number: 291-0057)
Paper towel or surgical gauze
Optional:
50 mL centrifuge tube (Falcon, Fisher Scientific, catalog number: 10788561) with 20–40 mL of bleach (for viruses/toxins)
Insulin syringe (BD, Fisher Scientific, catalog number: 13161931) (for toe tattooing)
Tattoo ink (Ketchum, VWR International, catalog number: NASCC01885N) (for toe tattooing)
Equipment
Cannula fabrication
30 cm ruler or tape measure
Straight tip haemostat, Stainless steel, 5" (Newport Spectra-Physics, catalog number: LAB-17) (see Note 1)
Intraperitoneal injection
Animal heating pad (Tonkey Electrical Technology, catalog number: TK-HPP4030) or disposable heating pad (e.g., HotHands Hand Warmers) (see Note 2)
10 μL syringe, 701 N, 26 G, 51 mm length, 0.47 mm external diameter (Hamilton, VWR International, catalog number: 549-1135) (see Note 3)
P20 pipette
Optional:
Class II biosafety cabinet (for viruses/toxins)
Rack for 50 mL centrifuge tubes (for viruses/toxins)
Magnifying light (to aid injection accuracy)
Procedure
Note: Appropriate personal protective equipment should be used, and particular care should be taken when working with toxic/harmful substances.
Cannula fabrication (Figure 1 and Video 1)
Figure 1. Fabrication of cannulas from 30 G needles. A) Cut an approximately 15 cm long piece of polyethylene tubing. B) Using a haemostat, remove the plastic hub from a 30 G needle and brush away any debris that remains on needle shaft. C) Insert ~2–3 mm of the detached needle shaft into the tubing. D) Store the fabricated cannulas in a 6-well plate. See also Video 1.
Video 1. Cannula fabrication for intraperitoneal injection of neonatal mice. 00:00–00:11: Removal of plastic hub from 30 G needle. 00:11–00:27: Clearing away plastic hub debris from needle base. 00:27–00:39: Needle insertion into polyethylene tubing. 00:39–01:08: Testing the cannula seal. 01:08–01:31: Clearing water from the cannula. 01:31–01:44: Cannula storage in a 6-well plate.
Measure and then cut the polyethylene tubing with the scalpel blade into pieces ~15 cm long (Figure 1A). Ensure the blade is sharp to avoid crushing the tubing.
Using the haemostat, clamp the base of the 30 G needle where it attaches to the plastic hub. Grab the plastic hub of the needle with your fingers and quickly rotate it. The plastic hub should pop off with minimal force (see Note 4) (Figure 1B).
Brush away any plastic residue at the base of the needle and then inspect it for any physical damage that may have occurred during the needle extraction (e.g., crushed or bent hub).
Carefully insert ~2–3 mm of the newly exposed base of the needle into the tubing until firmly attached (Figure 1C). Based on our experience, the combined use of 0.28 mm internal diameter tubing with a 30 G needle creates a watertight seal without further modification.
To confirm that there are no leaks in the cannula seal, carefully attach the free end of the polyethylene tubing onto a 30 G needle fixed to a 10 mL syringe containing sterile water and expel a small volume of liquid (see Note 5). To clear the water from the cannula and air-dry it, fix the tubing onto a 30 G needle connected to an empty 10 mL syringe and expel the remaining water. Fresh sterile 10 mL syringes are used to avoid contamination of the Hamilton syringes used for intraperitoneal injections.
Store the cannulas in a 6-well plate until ready to use (Figure 1D) (Pause Point).
Intraperitoneal injection (Figure 2 and Video 2)
Figure 2. Intraperitoneal injection of neonatal mice. A) Insert the 10 μL Hamilton syringe into the open end of the tubing attached to the fabricated cannula. B) Away from the sterile injection area, wait until the pup urinates and then use a cotton-swab to dry the pup. C) Move to the sterile injection area and restrain the pup with the ventral-side facing upwards. D) With the bevel facing upwards, insert the cannula needle at a shallow angle into the peritoneal cavity, before slowly injecting. The animal in these images was aged P2. See also Video 2.
Video 2. Encouragement of bladder voiding, intraperitoneal injection, and toe tattooing. 00:00–00:56: Encouraging voiding of the bladder (N.B., the mouse in the video has already voided). 00:56–01:19: Intraperitoneal injection. 01:19–01:52: Tattooing of the paw for identification. The animal in the video was aged P2.
Optional: If working with viruses/toxins, turn on the class II biosafety cabinet.
Turn on the animal heating pad and allow to warm to 38–40 °C.
Clean the working surface and then place a sterile surgical drape on top.
Sprinkle some home bedding material into the large weigh boat and place the boat on the heating pad (to hold pups post-injection for acute monitoring).
Optional: If working with viruses/toxins, place the 50 mL centrifuge tube containing bleach in the rack and ensure that the biohazard sharps bin is available.
Optional: If tattooing the intraperitoneally injected pups for identification purposes, set aside an insulin syringe and tattoo ink.
When ready for the injections, transfer the parents to a separate cage and include a scattering of their home bedding. Also, ensure they have access to food, water, and cage enrichment. Ideally, the parent cage will be kept in a separate room while the injections are being performed in order to avoid exposure to any potentially distressing vocalisations from the pups.
Place the original home cage with the pups onto the pre-warmed heating pad.
Attach the cannula to the 10 μL Hamilton syringe and lay on the surgical drape (Figure 2A).
Load an appropriate volume of the substance to be injected into the cannula (Table 1). Care must be taken during this step, as the polyethylene tubing can be intractable due to rigidity. Place the cannula needle into the microfuge tube containing the substance to be injected. Provide stability to the polyethylene tubing by gently holding it against the side of the microfuge tube so that the cannula needle tip remains submerged in the substance as you draw up solution into the Hamilton syringe. If this proves tricky, the microfuge tube can be placed on its side (for small volumes in the tube) or, alternatively, a small amount (e.g., 5–10 μL) can be pipetted onto parafilm and loaded from there (see Note 6)—but be wary of evaporation. If pipetting viruses/toxins, dispose of the pipette tips and parafilm in the 50 mL centrifuge tube with bleach.
Table 1. Approximate maximum volumes for intraperitoneal injections using this method from post-natal day 1 (P1) to P5 (see Note 7). For mice older than P5, more conventional intraperitoneal injection methods may be used, even though the method outlined here can also be used (see Note 3). Greater injection volumes via neonatal intraperitoneal injections have been reported (Foust et al., 2008).
Post-natal age (day) Maximum injection volume (μL)
1 2–4
2 4–6
3 6–8
4 8–10
5 > 10
Scent your gloves with home bedding and then transfer a single pup to the sterile surgical drape.
To empty the bladder and thus create more space for injecting substances into the abdominal area, it is encouraged to induce the righting reflex response, which often elicits voiding (Video 2). This can be achieved by placing the pup on a paper towel adjacent to where you will perform the injection. Gently roll the pup on its back with your fingers. The pups will automatically begin rhythmic kicking of the forelimbs and hindlimbs, and this is often concomitant with involuntary voiding. Wipe away the urine using a sterile cotton swab (Figure 2B) and transfer the pup to the sterile injection area. P3 and older pups usually do not need to have their bladders voided prior to injections.
Gently restrain the pup using your thumb and index finger around the abdomen with the ventral side facing upwards (Figure 2C). Carefully gather excess skin between your digits to provide an injection site (see Note 8).
Insert the cannula needle (bevel side up) into the intraperitoneal cavity between the midline and the fat pads (when visible), avoiding the liver and milk-filled stomach (Figure 2D and Video 2). Aim to insert 2–3 mm of the needle and do so at a shallow angle to avoid puncturing any organs. Slowly inject the pup, then grab the base of the needle to hold it stable and wait for 5–10 s to ensure absorption before carefully removing the needle (see Note 9).
Monitor the pup and injection site for potential backflow. Clean the injection site with a new cotton swab if needed (see Note 10) and then transfer the injected pup to the weigh boat on the heating pad. Cover the pup with home bedding.
Optional: To tattoo pups for post-injection identification purposes, coat the tip of an insulin syringe needle with tattoo ink (do this sparingly, as it can get messy). Gently restrain the paw to be tattooed and transdermally insert the needle (Video 2). You should encounter little to no resistance during the needle insertion. Gently insert and retract (but not fully) the needle a few times to deposit the ink under the skin (see Note 11). Once done, dab the paw on a paper towel or surgical gauze to remove excess ink. There should be little to no blood at the insertion site. Tattooing should ideally be performed at P0–P2.
Optional: If injecting viruses/toxins, before disconnecting the cannula from the syringe, draw some of the bleach from the 50 mL centrifuge tube up into the polyethylene tubing. Detach the tubing and needle from the syringe and dip it into bleach for a few seconds before disposing of it in the biohazard sharps bin. Place any used cotton swabs to the side.
Repeat steps B9–B14 or B9–B16 for each pup that is to be injected.
Once all pups have been dosed, transfer the litter back to the home cage and cover with nesting material (see Note 12). After a few minutes, move the parents back to home cage.
To clean up, transfer any virus/toxin-contaminated tools (e.g., pipette tips and cotton swabs) to the centrifuge tube with bleach. Spray the working surface (including surgical drape and weigh boat) with bleach solution and then fold the drape inward to contain all bleach-treated materials. Dispose of materials in the biohazard sharps bin.
Validation of protocol
This protocol or parts of it has been used and validated in the following research article(s):
MacDonald et al. (2021). A central mechanism of analgesia in mice and humans lacking the sodium channel NaV1.7. Neuron 109(9): 1497–1512.e6 (Supplemental Figure 3 and 4).
Sleigh et al. (2023). Boosting peripheral BDNF rescues impaired in vivo axonal transport in CMT2D mice. JCI Insight 8(9): e157191 (Supplemental Figure 9, panels A–C).
Notes
Instead of a haemostat, a benchtop vice may be used to remove the needle hub.
In place of the electric animal heating pad, disposable heating pads (e.g., HotHands hand warmers) or rechargeable heating pads (e.g., Unigear Rechargeable Hand Warmer) can be used. Disposable heating pads can be reused multiple times if stored in Ziploc bags to avoid further air exposure.
Syringes with larger volumes are required if injecting more than 10 μL (see Table 1).
Keeping the needle stationary while twisting and pulling the plastic hub results in clean separation without crushing or bending the needle (Video 1). If the needle base is crushed or bent, there may be insufficient back pressure to draw up solution into the cannula.
An improper seal will result in poor loading of the water and accumulation of bubbles in the tubing, especially if handling viscous solutions.
Similar to when testing the cannula seal during fabrication, ensure that there is a smooth draw of the solution up into the tubing and that there are no bubbles. If this occurs, detach the cannula from the syringe and then try a new one.
P1 is designated as the day after a litter is born.
A magnifying light can aid the injection process. There is no need to pre-clean the injection site.
Sometimes, the cannula seal can fail after the substance to be injected has been loaded. To ensure that this does not occur without you noticing, take care to observe the small bulge that forms beneath the skin upon injection. After the injection, you may also manipulate the syringe plunger to check for residual liquid. If you note that the full volume was not dispensed during the injection, gently wiggle the needle while it is still inserted into the intraperitoneal cavity. We find that doing this relieves the internal pressure within the abdominal cavity to allow for the remaining solution to be absorbed. As an additional optional precaution to aid visualisation, a dye such as trypan blue or filtered green food dye can be pre-mixed with the substance to be injected (Glascock et al., 2011).
Make sure to keep the cotton swabs for urine and virus/toxin leakage separate; a marker pen can be used to aid identification.
For an informative video protocol of paw/toe tattooing, see the JoVE Science Education Database (2021) reference.
If needed and permitted, pups can receive multiple intraperitoneal injections on subsequent days.
Acknowledgments
This work was supported by European Molecular Biology Long-Term Fellowship (ALTF 495-2018 to A.M.P.), the Dystonia Medical Research Foundation (DMRFPRF-2022-3 to A.M.P.), the Medical Research Council (MR/S006990/1 to J.N.S.), and the Rosetrees Trust (M806 to J.N.S.). The graphic abstract was created with www.BioRender.com. The protocol described here has been performed in MacDonald et al. (2021) and Sleigh et al. (2023).
Competing interests
The authors have no competing interest to declare.
Ethical considerations
Experimentation involving mice was performed under license from the UK Home Office in accordance with the Animals (Scientific Procedures) Act (1986) and was approved by the UCL Queen Square Institute of Neurology Ethical Review Committee.
References
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.
Foust, K. D., Poirier, A., Pacak, C. A., Mandel, R. J. and Flotte, T. R. (2008). Neonatal Intraperitoneal or Intravenous Injections of Recombinant Adeno-Associated Virus Type 8 Transduce Dorsal Root Ganglia and Lower Motor Neurons. Hum. Gene Ther. 19(1): 61–70.
Glascock, J. J., Osman, E. Y., Coady, T. H., Rose, F. F., Shababi, M. and Lorson, C. L. (2011). Delivery of Therapeutic Agents Through Intracerebroventricular (ICV) and Intravenous (IV) Injection in Mice. J. Vis. Exp.: e3791/2968-v.
Gombash Lampe, S. E., Kaspar, B. K. and Foust, K. D. (2014). Intravenous Injections in Neonatal Mice. J. Vis. Exp.: e3791/52037.
Hua, Y., Sahashi, K., Rigo, F., Hung, G., Horev, G., Bennett, C. F. and Krainer, A. R. (2011). Peripheral SMN restoration is essential for long-term rescue of a severe spinal muscular atrophy mouse model. Nature 478(7367): 123–126.
JoVE Science Education Database (2021). Lab Animal Research. Rodent Identification II. J Vis Exp.
Kienstra, K. A., Freysdottir, D., Gonzales, N. M. and Hirschi, K. K. (2007). Murine neonatal intravascular injections: modeling newborn disease.J. Am. Assoc. Lab. Anim. Sci. 46(6): 50–54.
MacDonald, D. I., Sikandar, S., Weiss, J., Pyrski, M., Luiz, A. P., Millet, Q., Emery, E. C., Mancini, F., Iannetti, G. D., Alles, S. R., et al. (2021). A central mechanism of analgesia in mice and humans lacking the sodium channel NaV1.7. Neuron 109(9): 1497–1512.e6.
Nizzardo, M., Simone, C., Salani, S., Ruepp, M. D., Rizzo, F., Ruggieri, M., Zanetta, C., Brajkovic, S., Moulton, H. M., Müehlemann, O., et al. (2014). Effect of Combined Systemic and Local Morpholino Treatment on the Spinal Muscular Atrophy Δ7 Mouse Model Phenotype. Clin. Ther. 36(3): 340–356.e5.
Ohshima, M., Taguchi, A., Tsuda, H., Sato, Y., Yamahara, K., Harada-Shiba, M., Miyazato, M., Ikeda, T., Iida, H., Tsuji, M., et al. (2015). Intraperitoneal and intravenous deliveries are not comparable in terms of drug efficacy and cell distribution in neonatal mice with hypoxia–ischemia. Brain Dev. 37(4): 376–386.
Ostermann, E., Macquin, C., Bahram, S. and Georgel, P. (2013). Use of In vivo Imaging to Monitor the Progression of Experimental Mouse Cytomegalovirus Infection in Neonates. J. Vis. Exp.: e3791/50409.
Passini, M. A. and Wolfe, J. H. (2001). Widespread Gene Delivery and Structure-Specific Patterns of Expression in the Brain after Intraventricular Injections of Neonatal Mice with an Adeno-Associated Virus Vector. J. Virol. 75(24): 12382–12392.
Pitulescu, M. E., Schmidt, I., Benedito, R. and Adams, R. H. (2010). Inducible gene targeting in the neonatal vasculature and analysis of retinal angiogenesis in mice. Nat. Protoc. 5(9): 1518–1534.
Sands, M. S. and Barker, J. E. (1999). Percutaneous intravenous injection in neonatal mice. Lab. Anim. Sci. 49(3): 328–330.
Sleigh, J. N., Villarroel-Campos, D., Surana, S., Wickenden, T., Tong, Y., Simkin, R. L., Vargas, J. N. S., Rhymes, E. R., Tosolini, A. P., West, S. J., et al. (2023). Boosting peripheral BDNF rescues impaired in vivo axonal transport in CMT2D mice. JCI Insight 8(9): e157191.
Statler, P. A., McPherson, R. J., Bauer, L. A., Kellert, B. A. and Juul, S. E. (2007). Pharmacokinetics of High-Dose Recombinant Erythropoietin in Plasma and Brain of Neonatal Rats. Pediatr. Res. 61(6): 671–675.
Turner, P.V., Brabb, T., Pekow, C. and Vasbinder, M.A. (2011). Administration of substances to laboratory animals: Routes of administration and factors to consider. J. Am. Assoc. Lab. Anim. Sci. 50(5): 600–613.
Yardeni, T., Eckhaus, M., Morris, H. D., Huizing, M. and Hoogstraten-Miller, S. (2011). Retro-orbital injections in mice. Lab Anim. 40(5): 155–160.
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Article Information
Copyright
© 2023 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
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Biological Sciences > Biological techniques
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4,827 | https://bio-protocol.org/en/bpdetail?id=4827&type=0 | # Bio-Protocol Content
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Peer-reviewed
Identification of Matriglycan by Dual Exoglycosidase Digestion of α-Dystroglycan
Ishita Chandel
KC Kevin P. Campbell
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4827 Views: 429
Reviewed by: Gal HaimovichPhilipp A.M. SchmidpeterQingliang Shen
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Original Research Article:
The authors used this protocol in eLIFE Feb 2023
Abstract
Matriglycan is a linear polysaccharide of alternating xylose and glucuronic acid units [-Xyl-α1,3-GlcA-β1,3]n that is uniquely synthesized on α-dystroglycan (α-DG) and is essential for neuromuscular function and brain development. It binds several extracellular matrix proteins that contain laminin-globular domains and is a receptor for Old World arenaviruses such as Lassa Fever virus. Monoclonal antibodies such as IIH6 are commonly used to detect matriglycan on α-DG. However, endogenous expression levels are not sufficient to detect and analyze matriglycan by mass spectrometry approaches. Thus, there is a growing need to independently confirm the presence of matriglycan on α-DG and possibly other proteins. We used an enzymatic approach to detect matriglycan, which involved digesting it with two thermophilic exoglycosidases: β-Glucuronidase from Thermotoga maritima and α-xylosidase from Sulfolobus solfataricus. This allowed us to identify and categorize matriglycan on α-DG by studying post-digestion changes in the molecular weight of α-DG using SDS-PAGE followed by western blotting with anti-matriglycan antibodies, anti-core α-DG antibodies, and/or laminin binding assay. In some tissues, matriglycan is capped by a sulfate group, which renders it resistant to digestion by these dual exoglycosidases. Thus, this method can be used to determine the capping status of matriglycan. To date, matriglycan has only been identified on vertebrate α-DG. We anticipate that this method will facilitate the discovery of matriglycan on α-DG in other species and possibly on other proteins.
Key features
• Analysis of endogenous matriglycan on dystroglycan from any animal tissue.
• Matriglycan is digested using thermophilic enzymes, which require optimum thermophilic conditions.
• Western blotting is used to assay the success and extent of digestion.
• Freshly purified enzymes work best to digest matriglycan.
Graphical overview
α-Dystroglycan (α-DG) from muscle is shown here modified by a phosphorylated core M3 glycan, which extends further and terminates in a repeating disaccharide of xylose (Xyl) and glucuronic acid (GlcA) called matriglycan. β-glucuronidase (Bgus) and α-xylosidase (Xyls) hydrolyze the β-1,3-linked GlcA and α-1,3 linked-Xyl, starting from the terminal residues.
Keywords: Matriglycan Dystroglycan Exoglycosidases β-Glucuronidase α-Xylosidase Thermophiles Glycosylation Digestion
Background
α-Dystroglycan (α-DG) is an extensively glycosylated and widely expressed transmembrane protein that functions as an extracellular matrix receptor. It is modified with several types of O- and N- glycans. The glycans on α-DG enable it to interact with the extracellular molecules and maintain cell stability and integrity. The most widely studied post-translational modification of α-DG is the addition of the O-mannosylated glycan called core M3. This glycan is defined as a phosphorylated O-mannosyl trisaccharide [GalNAcβ1-3GlcNAcβ1-4(phosphate-6)Man-O-Ser] comprised of a mannose that is extended by the sequential addition of an N-acetylglucosamine and an N-acetyl galactosamine (Yoshida-Moriguchi and Campbell, 2015). The C6 hydroxyl group of the mannose of core M3 is also phosphorylated by a kinase (Yoshida-Moriguchi et al., 2013). Core M3 is further elongated, from the N-acetylgalactosamine, by a glycan modification that terminates in the repeating disaccharide of xylose (Xyl) and glucuronic acid [-Xyl-α1,3-GlcA-β1,3-]n called matriglycan. Notably, over 18 genes are involved in generating the final core M3 structure (Walimbe et al., 2020). Matriglycan acts as a scaffold for laminin-G domain containing proteins (e.g., laminin, agrin, perlecan, and neurexin) in the extracellular matrix. Loss-of-function mutations in any of the 18 genes involved in matriglycan synthesis lead to different types of muscular dystrophies, including severe forms such as Walker-Warburg syndrome.
Owing to the complexity of the final core M3 structure terminating in matriglycan and its low expression levels in cells, it has never been analyzed using mass spectrometry. Therefore, to study and to confirm the presence of matriglycan on dystroglycan in various animal tissues, mutants, and patient samples, we use an enzymatic approach. We identified two exoglycosidases that together can hydrolyze matriglycan in native tissues: β-glucuronidase (Bgus) from Thermotoga maritima (Salleh et al., 2006) and α-xylosidase (Xyls) from Sulfolobus solfataricus (Moracci et al., 2000). In conjunction with mass spectrometry and chemically synthesized matriglycan, we previously demonstrated that these enzymes sequentially remove sugars from the non-reducing end and that Bgus and Xyls specifically cleave the β1,3-linked GlcA and α1,3-linked Xyl, respectively (Briggs et al., 2016). Given that neither enzyme has detectable endoglycosidase activity (Briggs et al., 2016), they enable us to probe the matriglycosylation status of native α-DG. Matriglycan from brain was found to be capped by a 3-O-sulfation of its terminal GlcA, a reaction catalyzed by HNK-1 sulfotransferase. The sulfate cap prevents further elongation of matriglycan and also makes it resistant to digestion by the dual exoglycosidases. It has been previously demonstrated that the matriglycan on α-DG isolated from brain is capped by sulfation and becomes susceptible to digestion by the exoglycosidases after digesting it with a sulfatase (Sheikh et al., 2020). Therefore, this method enables us to probe the tissue-dependent matriglycosylation status of α-DG. To date, matriglycan has been found only on α-DG, although a report suggests it could be present on glypican-4 (Inamori et al., 2016). This digestion protocol is expected to be useful in confirming the presence of matriglycan on other proteins such as glypican and may also aid in discovering new proteins modified with matriglycan. Also, matriglycan has been found only in vertebrate α-DG, e.g., in human, mouse, and zebrafish (Praissman et al., 2016; Liu et al., 2020), and this protocol can be used to determine whether matriglycan exists in other species. Such analyses have the potential to broaden the use of other species as a model organism for the study of muscular dystrophies arising from changes in matriglycosylation of α-DG. Although matriglycan can be detected using antibodies, those currently available sometimes detect nonspecific bands; therefore, confirming the presence of matriglycan independently will be important, especially when working with new proteins or organisms. In summary, the technique described here is expected to be useful for definitively detecting matriglycan (in contrast to antibodies) on new proteins and in new organisms.
One limitation of using thermophilic enzymes for digestion is that they require high temperatures and acidic pH to function, which could be detrimental to some substrate proteins. We have found α-DG to remain remarkably stable at high temperatures and acidic pH, making the use of the dual thermophilic exoglycosidases feasible (Briggs et al., 2016).
Materials and reagents
Biological materials
BL21(DE3) chemically competent E. coli cells (Invitrogen, One Shot, C600003)
C57BL/6J mice (The Jackson Laboratory, 000664)
Reagents
BGUS_pET-28a(+) (GenScript, SC1691, custom-made)
XylS_pET-28a(+) (GenScript, SC1691, custom-made)
IPTG (Isopropylthio-β-galactoside) (IBI Scientific, catalog number: IB02105)
Kanamycin (IBI Scientific, catalog number: IB02120)
TALON superflow metal affinity resin (Takara Bio, catalog number: 635506)
Econo-Column chromatography columns, 1.5 cm × 10 cm, glass (Bio-Rad, catalog number: 7371512)
Nuclease, Pierce Universal nuclease (Thermo Scientific, catalog number: 88701)
30 KD cutoff concentrators, Amicon Ultra centrifugal filters (Sigma-Aldrich, catalog number: UFC803024)
Coomassie stain, G-250 stain (Bio-Rad, catalog number: 1610786)
2-mercaptoethanol (VWR, catalog number: 97064-880)
Wheat germ agglutinin (WGA), agarose bound (Vector Laboratories, catalog number: AL-1023-10)
Core DG antibody (Bio-Techne, R&D systems, catalog number: AF6868)
Anti-matriglycan antibody (DSHB, catalog number: IIH6 C4)
SOC medium (Invitrogen, One Shot, catalog number: C600003)
Acrylamide, 30% Acrylamide/Bis solution, 37.5:1 (Bio-Rad, catalog number: 1610158)
Ammonium persulfate (APS) (Sigma-Aldrich, catalog number: 248614-500G)
TEMED (RPI, catalog number: T18000-0.05)
N-Acetylglucosamine (Sigma-Aldrich, catalog number: A8625)
Bacto-Agar (BD Difco, catalog number: 214010)
Bacto-Tryptone (BD Difco, catalog number: 211705)
NaCl (IBI Scientific, catalog number: IB07071)
Yeast extract (BD Difco, catalog number: 212750)
Tris (Roche, catalog number: 11814273001)
Triton X-100 (Fisher Chemicals, catalog number: BP151)
Pepstatin (EMD-Millipore, catalog number: 516481)
PMSF (Phenylmethylsulfonyl fluoride) (Sigma-Aldrich, catalog number: P7626)
Benzamidine hydrochloride hydrate (MP Biomedicals, catalog number: 195068)
Calpeptin (EMD-Millipore, catalog number: 03-34-0051)
Calpain inhibitor (Sigma-Aldrich, catalog number: A6185)
Leupeptin (Sigma-Aldrich, catalog number: EI8)
Aprotinin (Sigma-Aldrich, catalog number: A3886)
Glycine (Bio-Rad, catalog number: 1610718)
Methanol (Sigma-Aldrich, catalog number: 179337)
KCl (potassium chloride) (Fisher Chemicals, catalog number: P217)
Na2HPO4 (sodium dibasic monophosphate) (Fisher Chemicals, catalog number: S374)
KH2PO4 (potassium dihydrogen phosphate) (Sigma-Aldrich, catalog number: P5655)
CaCl2·2H2O (calcium chloride) (Sigma-Aldrich, catalog number: C8106)
MgCl2·6H2O (magnesium chloride hexahydrate) (Fisher Chemicals, catalog number: M33)
Sodium acetate (Fisher Chemicals, catalog number: BP334)
SDS (sodium dodecyl sulfate) (SERVA, catalog number: 20765)
EDTA (ethylenediaminetetraacetic acid) (PRIMA, catalog number: KCE14000)
Filtration system PES 0.45 µm (Cole-Parmer, catalog number: UX-07630-08)
Solutions
Luria-Bertani (LB) medium (see Recipes)
Luria-Bertani (LB) agar plates-Kanamycin (see Recipes)
Lysis buffer (see Recipes)
Wash buffer 1 (see Recipes)
High salt wash buffer (see Recipes)
Elution buffer (see Recipes)
Solubilization buffer (see Recipes)
WGA wash buffer (see Recipes)
WGA elution buffer (see Recipes)
10× Phosphate buffered saline (PBS) pH 7.4 (see Recipes)
Sodium acetate buffer (see Recipes)
Two (3%–15%) SDS-PAGE separating gradient gels (see Recipes)
Two stacking gels (see Recipes)
Separating gel buffer (see Recipes)
Stacking gel buffer (see Recipes)
10× Western blot transfer buffer (see Recipes)
1× Western transfer tank buffer (see Recipes)
Recipes
Luria-Bertani (LB) medium
Reagent Final concentration Quantity
Bacto-Tryptone 1% 10 g
Yeast extract 0.5% 5 g
NaCl 1% 10 g
H2O n/a 1,000 mL
Total n/a 1,000 mL
Autoclave for 30 min
Store at room temperature
Luria-Bertani (LB) agar plates-Kanamycin
Reagent Final concentration Quantity
Bacto-Tryptone 1% 10 g
Yeast extract 0.5% 5 g
NaCl 1% 10 g
Agar 2% 20 g
Kanamycin 50 μg/mL 50 μL (from 50 mg/mL stock)
H2O n/a 1,000 mL
Total n/a 1,000 mL
Autoclave for 30 min
Note: Add Kanamycin after media has cooled down to 65–70°C. Store plates at 4 °C
Lysis buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 100 mM 1.25 mL 4 M
Triton X-100 1% 2.5 mL 20%
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.75 M 0.375 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
Wash buffer 1
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 5 mL 1 M
NaCl 100 mM 2.5 mL 4 M
Triton X-100 0.1% 0.5 mL 20%
Pepstatin A 0.6 μg/mL 0.06 mL 1 mg/mL
Aprotinin 0.5 μg/mL 0.01 mL 5 mg/mL
Leupeptin 0.5 μg/mL 0.01 mL 5 mg/mL
Calpain I inhibitor 5 μM 0.1 mL 5 mM
Calpeptin 5 μM 0.1 mL 5 mM
PMSF 0.1 mM 0.1 mL 0.1 M
Benzamidine 0.75 mM 0.750 mL 0.1 M
Total n/a 100 mL
Always prepare fresh
High salt wash buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 500 mM 6.25 mL 4 M
Triton X-100 0.1% 0.25 mL 20%
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.75 mM 0.375 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
Elution buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 100 mM 1.25 mL 4 M
Triton X-100 0.1% 0.25 mL 20%
Imidazole 300 mM 15 mL 1 M
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.75 mM 0.375 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
Solubilization buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 100 mM 1.25 mL 4 M
Triton X-100 1% 2.5 mL 20%
EDTA 10 mM 1 mL 0.5 M
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.75 mM 0.375 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
WGA wash buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 100 mM 1.25 mL 4 M
Triton X-100 0.1% 0.25 mL 20%
EDTA 10 mM 1 mL 0.5 M
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.76 mM 0.376 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
WGA elution buffer
Reagent Final concentration Quantity Stock concentration
Tris-HCl 50 mM 2.5 mL 1 M
NaCl 100 mM 1.25 mL 4 M
Triton X-100 0.1% 0.25 mL 20%
EDTA 10 mM 1 mL 0.5 M
N-acetylglucosamine 0.3 M 3.31 grams n/a
Pepstatin A 0.6 μg/mL 0.03 mL 1 mg/mL
Aprotinin 0.5 μg/mL 5 μL 5 mg/mL
Leupeptin 0.5 μg/mL 5 μL 5 mg/mL
Calpain I inhibitor 5 μM 0.05 mL 5 mM
Calpeptin 5 μM 0.05 mL 5 mM
PMSF 0.1 mM 0.05 mL 0.1 M
Benzamidine 0.77 mM 0.377 mL 0.1 M
Total n/a 50 mL
Always prepare fresh
10× Phosphate buffered saline (PBS)
Reagent Final concentration Quantity
NaCl 1.37 M 80 g
KCl 27 mM 2 g
Na2HPO4 100 mM 14.4 g
KH2PO4 18 mM 2.4 g
CaCl2·2H2O 10 mM 1.33 g
MgCl2·6H2O 5 mM 1.0 g
H2O n/a 1,000 mL
Adjust pH to 7.4 with HCl
Store at room temperature
Sodium acetate buffer
Reagent Final concentration Quantity
Sodium acetate 150 mM 10.2 g
Adjust pH to 5.5
H2O n/a 500 mL
Total n/a 500 mL
Store at room temperature
Two (3%–15%) SDS-PAGE separating gradient gels
Reagent 3% 15%
Water 19.3 mL 7.3 mL
Separating gel buffer 7.5 mL 7.5 mL
Acrylamide 3 mL 15 mL
TEMED 11 μL 11 μL
10% APS 180 μL 180 μL
Prepare fresh 10% APS
Two stacking gels
Reagent Quantity
Water 13.3 mL
Stacking gel buffer 5 mL
Acrylamide 2.3 mL
TEMED 15 μL
10% APS 150 μL
Prepare fresh 10% APS
Separating gel buffer
Reagent Final concentration Quantity
Tris
Adjust pH to 8.8 with HCL
1.5 M 90 g
SDS 0.4% 2 g
H2O n/a 500 mL
Total n/a 500 mL
Sterilize through 0.45 μm filter
Store at room temperature
Stacking gel buffer
Reagent Final concentration Quantity
Tris
Adjust pH to 6.8 with HCL
0.5 M 30 g
SDS 0.4% 2 g
H2O n/a 500 mL
Total n/a 500 mL
Sterilize through 0.45 μm filter
Store at room temperature
10× Western blot transfer buffer
Reagent Quantity
Glycine 288 g
Tris base 60.6 g
Water 2,000 mL
Store at room temperature
1× Western transfer tank buffer
Reagent Quantity
10× transfer buffer 400 mL
100% methanol 800 mL
Water 2,800 mL
Total 4,000 mL
Laboratory supplies
Rattler plating beads, 5 mm (Zymo Research, catalog number: S1001)
Liquid scintillation vials (RPI, catalog number: 121043)
Glass homogenizer (Cole-Parmer, catalog number: EW-44468-18)
Plain plunger (Cole-Parmer, EW-44468-06)
PVDF membrane (Immobilon-FL, Millipore, catalog number: IPFL00010)
Blotting paper (Cytiva Whatman 3MM CHR, catalog number: 3030-917)
Petri dishes (Fisher Scientific, catalog number: FB0875713)
Scissors (FST, catalog number: 14090-11)
Forceps (FST, catalog number: 91150-20)
Mortar and pestle (Fisher Scientific, catalog number: S39831)
Equipment
Thermomixer (Eppendorf, catalog number: 5382000023)
Thermo Top (Eppendorf, catalog number: 5308000003)
Polytron homogenizer (Fisher Scientific, Kinematica Polytron PT 2500 E homogenizer, catalog number: 08-451-320)
Generator/probe for Polytron homogenizer (Fisher Scientific, Kinematica, catalog number: 05-400-263)
Spectrophotometer (Eppendorf Bio photometer, model: 6131)
Water bath (Fisher Scientific, Isotemp model: 205)
Incubator (VWR, New Brunswick, Innova 42, catalog number: 75875-638)
Rotator (ATR Rotamix, model: RKVSD)
Motorized overhead stirrer (Fisher Scientific, DWK Life Sciences Wheaton, catalog number: 22-244382)
SDS-PAGE gradient gel system (Hoefer SE 600 model, 18 cm × 16 cm)
Western blot transfer system (Hoefer, TE 42 model)
Procedure
Gene cloning
β-glucuronidase (Bgus) from Thermotoga maritima and α-xylosidase (Xyls) from Sulfolobus solfataricus were cloned into pET-28a(+) vector between NheI/XhoI sites in frame with the N-terminal 6x-His tag by GenScript (see supplemental file S1).
Cell transformation
Thaw BL21DE3 competent cells on ice (50 μL aliquot for each plasmid).
Add 20 ng of each plasmid to the competent cells (mix by swirling the pipette tip three times).
Incubate cells in ice for 30 min.
Heat shock the cells at 42 °C for 30 s.
Incubate on ice again for 5 min. Critical: This step is required to obtain high efficiency of transformation.
Add 950 μL of SOC medium (at room temperature) and incubate at 37 °C for 60 min with shaking at 350 rpm in a thermomixer.
For each sample, spread 50 μL from each tube onto LB agar plates with Kanamycin (use plating beads or spreader).
Incubate the plates at 37 °C for 16 h.
Purification of Bgus and Xyls
Pick a single colony from each plate and inoculate 20 mL of LB medium (with Kanamycin, 50 μg/mL). Incubate at 37 °C for 16 h with shaking at 200 rpm.
Use 10 mL of the overnight culture to inoculate 1 L of LB medium (with Kanamycin, 50 μg/mL). Incubate at 37 °C with shaking at 200 rpm. Keep checking the OD600.
Once the OD600 reaches 0.6, add 1 mM IPTG (1.6 mL from a 600 mM stock solution) to the culture to induce gene expression and incubate at 16 °C for 16 h with shaking at 200 rpm.
Centrifuge the cells at 5,000× g for 10 min at 4 °C.
Store the cell pellets at -80 °C until ready for the purification step. Pause point: Cell pellets can be stored long term if necessary.
Dissolve the cell pellets in lysis buffer (20 mL of buffer per liter of starting culture).
Store the cells at -80 °C overnight to allow ice crystals to form; this step aids in cell lysis.
Thaw the cells at room temperature for purification.
Add nuclease at 6.25 kU (5 μL) concentration and sonicate cells for 5 s (power 4–5) four times with 10 s intervals at 4 °C.
Centrifuge cells at 15,000× g for 20 min at 4 °C. Place supernatant into a new tube.
Place the supernatant into a 75 °C water bath for 10 min. Critical: This step is essential to enrich the thermophilic enzymes.
Spin cells at 15,000× g for 30 min at 4 °C. Transfer supernatant to fresh tube. This is the cell extract.
Note: Perform steps C13–C17 in a cold room.
Prepare 2 TALON superflow metal affinity columns, one for each enzyme, by packing 3 mL of resin into a Econo-Column chromatography column and equilibrating with 10 mL of wash buffer 1.
Apply the cell extract obtained in step C12 to the column three times. Each time, incubate the column with the extract for 15–20 min on a gentle rocking platform. Save all flowthroughs.
Wash the column with 10 mL of wash buffer 1 three times. Save all washes.
Wash the column with high salt wash buffer to remove nonspecific interactions. Save the wash.
Elute the bound proteins with 3 mL elution buffer in five fractions.
Run 100 μL of each saved wash, flowthrough, and elution on an SDS-PAGE gel and visualize with Coomassie blue.
Combine the relevant fractions (usually 1 and 2, see Figure 1) and buffer exchange with PBS pH 7.4 using a 30 kD Amicon concentrator. Use a total dilution factor of 200 and bring the total volume down to 1 mL. Confirm pH is approximately 7.4 with strips.
Aliquot the combined fractions into PCR tubes (50 μL per tube) and flash freeze in liquid nitrogen. Transfer aliquots to a -80 °C freezer for long-term storage.
Figure 1. Purification of Bgus and Xyls. A. Column-purified Bgus is shown here in Eluates 1 and 2 and runs at ~66 kD in a 3%–15% gradient SDS-PAGE gel stained with Coomassie blue. B. Column-purified Xyls is shown here in Eluates 1 and 2 and runs at ~80 kD in a 3%–15% gradient SDS-PAGE gel stained with Coomassie blue.
Digestion of α-Dystroglycan (α-DG)
Use wheat germ agglutinin (WGA) agarose to enrich α-DG from animal tissues, as follows:
Harvest 1 g of skeletal muscle from C57BL/6J mice. Pour liquid nitrogen into a mortar and pestle and crush the muscle in it.
Note: Take muscles from the legs after euthanizing mice by cervical dislocation. Briefly, remove the skin by making a small cut in the mid-dorsal or ventral region and peel it off the mouse; then, using scissors and forceps, take all muscles from the legs including quadriceps, tibialis anterior, and soleus, until bone is visible.
Transfer powdered muscle into a vial and allow to warm for 3–5 min, either on ice or in a cold room.
Note: Vial should be able to withstand cold liquid nitrogen. We use plastic liquid scintillation vials.
Add 10 mL of solubilization buffer and mince the tissue in a polytron homogenizer for 15 s at speed 4. Repeat this three times.
Transfer solution to a 50 mL glass homogenizer and macerate tissue by moving the plunger in and out of the solution at least 10 times.
Note: For macerating tissue with an automated overhead stirrer (instead of doing it by hand), use speed 4.
Transfer homogenized material to a 50 mL Falcon tube and incubate on a rotator for 1 h at 4 °C.
Spin the material for 30 min at 20,000× g at 4 °C. Transfer supernatant to new tube.
Equilibrate 500 μL of WGA beads with 5 mL of solubilization buffer. Spin down beads at 1,500× g for 5 min. Remove buffer.
Combine material obtained in step D1e with the equilibrated WGA beads and incubate for 16 h on rotator at 4 °C.
Spin down beads at 1,500× g for 5 min at 4 °C and save the supernatant as WGA-void fraction.
Wash WGA beads with 5 mL of WGA wash buffer by incubating for 5 min on a rotator and then spinning beads at 1,000× g for 10 min at 4 °C. Repeat this wash step four times.
α-DG is now on the WGA beads. Add 1 mL of WGA elution buffer to elute α-DG from 500 μL of WGA beads and incubate at 4 °C for 1 h on a rotator.
Spin down beads at 1,500× g for 5 min at 4 °C. α-DG is now in the supernatant or eluate.
Buffer exchange 500 μL of the WGA eluate with sodium acetate (pH 5.5) using a 30 kD Amicon concentrator. Use a total dilution factor (DF) of 80 for successfully changing the pH. Critical: This step is essential for obtaining the pH (5.5) at which both enzymes work.
Note: For buffer exchange, follow these steps:
Add 1.5 mL of sodium acetate buffer to 500 μL of WGA eluate in the upper reservoir and spin at 2,000× g for 4 min at 4 °C (DF1 = 4). Approximately 700 μL remains in the upper reservoir. Discard the solution collected in the centrifuge tube.
Add 2.1 mL of sodium acetate buffer to the 700 μL solution from the previous step (DF2 = 4). Spin at 2,000× g for 4 min at 4 °C. Approximately 500 μL remains in the upper reservoir.
Add 2 mL of sodium acetate buffer to the 500 μL solution obtained in the previous step (DF3 =5). Spin at 2,000× g for 4 min at 4 °C. The solution in the upper reservoir (~ 500 μL) is now at pH 5.5 and the total dilution factor used is 80 (DF1 × DF2 × DF3 = 4 × 4 × 5 = 80)
Add 10 mM β-mercaptoethanol (~ 0.7 μL from 12.8 M stock) to the buffer-exchanged eluate and heat for 5 min at 99 °C. Critical: This step is essential to remove disulfide bridges and unfold the protein to make matriglycan accessible for digestion.
Add all protease inhibitors (at final concentrations given in the Recipes) after the solution is cooled down.
Thaw the 50 μL of enzyme aliquots on ice and add them to the above mixture.
Aliquot 100 μL as the initial timepoint (To) and incubate the rest of the mixture at 75 °C for 16 h with shaking at 600 rpm in a thermomixer.
Note: Aliquots from other timepoints can also be taken (e.g., at 4, 6, and 10 h, for T4, T6, and T10, respectively) to check the rate of progression of the digestion. However, for complete digestion of matriglycan on DG, we recommend incubating the samples for 16–23 h (Figure 2E).
Figure 2. Digestion of matriglycan by Bgus and Xyls. A, B. Wheat germ agglutinin (WGA) eluate from mouse skeletal muscle digested with Bgus and Xyls at 65 °C and 75 °C. TON indicates overnight digestion of 16 h. The digestion products are immunoblotted with (A) core DG antibody AF6868 1:200 and (B) anti-matriglycan antibody, IIH6, 1:100. C, D. WGA eluate of mouse skeletal muscle digested with Bgus and Xyls at 75 °C overnight (TON, 16 h) and immunoblotted with (C) AF6868 1:200 and (D) IIH6 1:100. E. Progress of digestion over time, as monitored at 2 (T2), 4 (T4), 6 (T6), and 20 h (T20); immunoblotting is with antibody against matriglycan IIH6 (1:100).
SDS-PAGE and western blotting
We analyze the progression and effectiveness of digestion by running the samples in 3%–15% SDS-PAGE gradient gels and performing immunoblotting on PVDF membranes, using antibodies against matriglycan (IIH6, 1:100), against core α-DG (AF6868 1:200) (Figure 2), and/or laminin overlay and solid phase laminin binding assay (Walimbe et al., 2020).
Data analysis
Upon successful digestion, clear shifts in the molecular weights of α-DG and matriglycan are observed. We also measure changes in laminin binding by performing laminin overlay and solid phase binding analysis (Walimbe et al., 2020, Figure 5D and 5E). Blots are scanned using a Li-Cor Odyssey imaging system and fluorescence detection, and images are analyzed using the Image Studio software. A detailed description of data analysis was provided previously (Briggs et al., 2016, Supplementary figure 1).
Validation of protocol
We have routinely used this method to identify matriglycan and found it to be robust and reproducible (Okuma et al., 2023; Briggs et al., 2016; Sheikh et al., 2020; Walimbe et al., 2020). Additional data validating our method is provided in Figure 1 and Figure 2.
General notes and troubleshooting
General notes
We have found that these enzymes are most effective when freshly purified.
Troubleshooting
If the enzymes are 4–5 months old and do not seem to work, it is best to purify fresh samples.
Acknowledgments
Research reported in this publication was supported by the National Institute of Neurological Disorders and Stroke of the National Institutes of Health under Award Number P50NS053672. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. We are thankful to Dr. Christine Blaumueller of the Scientific Editing and Research Communication Core at the University of Iowa Carver College of Medicine for her critical reading of the manuscript. We also thank Briggs et al. (2016) for first demonstrating the utility of the thermophilic enzymes in digesting matriglycan. K.P.C. is an investigator of the Howard Hughes Medical Institute.
Competing interests
The authors declare that no competing interests exist.
Ethical considerations
All mice were maintained in a barrier-free, specific pathogen-free grade facility and had access to normal chow and water ad libitum. All animals were manipulated in biosafety cabinets and change stations using aseptic procedures. The mice were maintained in a climate-controlled environment at 25 °C on a 12:12 h light/dark cycle. Animal care, ethical usage, and procedures were approved and performed in accordance with the standards set forth by the National Institutes of Health and the University of Iowa Animal Care and Use Committee (IACUC).
References
Briggs, D. C., Yoshida-Moriguchi, T., Zheng, T., Venzke, D., Anderson, M. E., Strazzulli, A., Moracci, M., Yu, L., Hohenester, E., Campbell, K. P., et al. (2016). Structural basis of laminin binding to the LARGE glycans on dystroglycan. Nat. Chem. Biol. 12(10): 810–814.
Inamori, K. I., Beedle, A. M., de Bernabe, D. B., Wright, M. E. and Campbell, K. P. (2016). LARGE2-dependent glycosylation confers laminin-binding ability on proteoglycans. Glycobiology 26(12): 1284–1296.
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Supplementary information
The following supporting information can be downloaded here:
File S1.
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© 2023 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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4,828 | https://bio-protocol.org/en/bpdetail?id=4828&type=0 | # Bio-Protocol Content
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Genome-wide Mapping of 5′-monophosphorylated Ends of Mammalian Nascent RNA Transcripts
Michael A. Cortázar
NF Nova Fong
DB David L. Bentley
Published: Vol 13, Iss 18, Sep 20, 2023
DOI: 10.21769/BioProtoc.4828 Views: 352
Reviewed by: Gal HaimovichMarion HoggRohini Ravindran Nair
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Original Research Article:
The authors used this protocol in Genes & Development Nov 2022
Abstract
In eukaryotic cells, RNA biogenesis generally requires processing of the nascent transcript as it is being synthesized by RNA polymerase. These processing events include endonucleolytic cleavage, exonucleolytic trimming, and splicing of the growing nascent transcript. Endonucleolytic cleavage events that generate an exposed 5′-monophosphorylated (5′-PO4) end on the growing nascent transcript occur in the maturation of rRNAs, tRNAs, and mRNAs. These 5′-PO4 ends can be a target of further processing or be subjected to 5′-3′ exonucleolytic digestion that may result in termination of transcription. Here, we describe how to identify 5′-PO4 ends of intermediates in nascent RNA metabolism. We capture these species via metabolic labeling with bromouridine followed by immunoprecipitation and specific ligation of 5′-PO4 RNA ends with the 3′-hydroxyl group of a 5′ adaptor (5′-PO4 Bru-Seq) using RNA ligase I. These ligation events are localized at single nucleotide resolution via highthroughput sequencing, which identifies the position of 5′-PO4 groups precisely. This protocol successfully detects the 5′monophosphorylated ends of RNA processing intermediates during production of mature ribosomal, transfer, and micro RNAs. When combined with inhibition of the nuclear 5′-3′ exonuclease Xrn2, 5′-PO4 Bru-Seq maps the 5′ splice sites of debranched introns and mRNA and tRNA 3′ end processing sites cleaved by CPSF73 and RNaseZ, respectively.
Key features
• Metabolic labeling for brief periods with bromouridine focuses the analysis of 5′-PO4 RNA ends on the population of nascent transcripts that are actively transcribed.
• Detects 5′-PO4 RNA ends on nascent transcripts produced by all RNA polymerases.
• Detects 5′-PO4 RNA ends at single nucleotide resolution.
Keywords: 5′-PO4 Bru-Seq 5′-monophosphorylated ends Nascent transcripts RNA Bromouridine labeling Immunoprecipitation
Background
Tracking genome-wide active transcription and its regulation has been made possible by several complementary approaches (Wissink et al., 2019). These include isolation of chromatin-associated RNAs (Bhatt et al., 2012, Mayer et al., 2015, Weber et al., 2014), RNA polymerase–associated RNAs (Churchman and Weissman, 2011, Nojima et al., 2015, Fong et al., 2017), or nascent transcripts pulse-labeled with nucleoside analogs such as 4-thiouridine (Schwalb et al., 2016, Kenzelmann et al., 2007, Rabani et al., 2011, Herzog et al., 2017, Muhar et al., 2018, Schofield et al., 2018), 5-ethenyluridine (Jao and Salic, 2008), or bromouridine (Paulsen et al., 2014).
In eukaryotic cells, endonucleolytic cleavage of the nascent transcript is used to release fully transcribed RNAs from chromatin or to release small RNAs from longer precursors during transcription [i.e., micro RNAs (miRNAs) and intron-encoded small nucleolar RNAs (snoRNAs)]. Most of these cleavage events are carried out by nucleases that leave 5′-phosphate (5′-PO4) and 3′ OH ends, including CPSF73, Int11, RNaseP, RNaseZ, and Drosha.
Recently, POINT-5 technology identified 5′ ends generated at cleavage sites of RNA pol II–associated transcripts from runoff products of 5′ RACE in reverse transcription reactions (Sousa-Luis et al., 2021), but this approach does not inform on the identity of the chemical group at these 5′ RNA ends. Previously, methods have been developed to specifically map 5′-PO4 RNA ends by virtue of their ability to be ligated to adaptors by RNA ligase I (Harigaya and Parker, 2012, German et al., 2008).
Here, we describe 5′-PO4 Bru-Seq for direct detection of 5′-PO4 groups in nascent transcripts produced by all RNA polymerases in the cell (Figure 1). The method was previously validated and used to identify targets of the Xrn2 exonuclease (Cortazar et al., 2022). In this method, total RNA is fragmented with Micrococcal Nuclease (MNase) that produces 5′ OH and 3′ PO4 ends, and nascent transcripts are enriched by immunoprecipitation of 5-bromouridine pulse-labeled molecules, followed by repair of 3′ phosphates with T4 polynucleotide kinase, ligation of 5′ and 3′ adaptors, and PCR amplification of sequencing libraries. Only RNA fragments with native 5′-PO4 ends can be incorporated into these libraries (Harigaya and Parker, 2012). Unique molecular identifiers (UMIs) are included to allow removal of PCR duplicates. The molecular events that result in 5′-PO4 groups on nascent RNA ends detected by this method potentially include cleavage by endonucleases, intron debranching, and decapping.
Figure 1. The 5′-PO4 Bru-Seq protocol. 5′-PO4 Bru-Seq enriches transcripts being extended by actively transcribing polymerases via metabolic labeling with bromouridine, fragmentation by micrococcal nuclease (MNase), and immunoprecipitation. The 3′-ends generated by MNase cleavage events are repaired by T4 polynucleotide kinase (PNK) in the absence of ATP. 5′-PO4 ends of nascent transcripts are then ligated to the 3′-hydroxyl group of a 5′ adaptor, amplified by PCR, and deep sequenced using Illumina sequencing technology.
Materials and reagents
Mammalian cell type of interest (e.g., HEK293, ATCC, catalog number: CRL-1573)
Cell culture media supplemented with 10% serum (e.g., DMEM supplemented with 10% FBS; DMEM, Thermo Fisher Scientific, catalog number: 11995040; FBS, VWR, catalog number: 97068-085)
10 cm tissue culture dishes (e.g., Genesee Scientific, catalog number: 25-200)
5/10 mL sterile serological pipets (e.g., Genesee Scientific, catalog numbers: 12-102, 12-104)
15/50 mL conical bottom centrifuge tubes (e.g., Corning, catalog numbers: 05-538-59A, 05-526B)
1.7 mL microcentrifuge tubes (e.g., Genesee Scientific, catalog number: 22-282)
TRIzol reagent (Thermo Fisher Scientific, catalog number: 15596018)
Chloroform (Fisher Scientific, catalog number: BP1145-1)
Isopropanol (Fisher Scientific, catalog number: BP2618-1)
80% ethanol, molecular biology grade (e.g., Thermo Fisher Scientific, catalog number: T08204K7)
GeneRuler 1 kb Plus DNA ladder (Thermo Fisher Scientific, catalog number: FERSM1333)
5 M NaCl, RNase-free (Thermo Fisher Scientific, catalog number: AM9760G)
Micrococcal nuclease (MNase) (2,000,000 gel units/mL) (New England Biolabs, catalog number: M0247S)
10× MNase reaction buffer (New England Biolabs, catalog number: B0247S)
0.5 M EGTA, pH 8.0, DNase and RNase Free (Thermo Fisher Scientific, catalog number: NC1874048)
Nuclease-free water (Thermo Fisher Scientific, catalog number: AM9939)
10× phosphate-buffered saline (PBS), pH 7.4, RNase-free (Thermo Fisher Scientific, catalog number: AM9624)
5-bromouridine (Sigma-Aldrich, catalog number: 850187)
Triton X-100 (Sigma-Aldrich, catalog number: X100)
1 M DTT (Thermo Fisher Scientific, Catalog number: P2325)
Superase-InTM RNase inhibitor (Invitrogen, AM2696)
Protein G magnetic beads (Pierce Thermo Scientific, catalog number: 88848)
Purified mouse anti-BrdU antibody (clone 3D4, RUO) (BD Pharmingen, catalog number: 555627)
QubitTM RNA High Sensitivity Assay kit (Thermo Fisher Scientific, catalog number: Q32852)
InvitrogenTM QubitTM assay tubes (Thermo Fisher Scientific, catalog number: Q32856)
T4 polynucleotide kinase (Thermo Fisher Scientific, catalog number: EK0031)
10× reaction buffer A (Thermo Fisher Scientific, catalog number: EK0031)
RNA Clean & ConcentratorTM-5 kit (ZYMO RESEARCH, catalog number: R1013, R1014)
QIAseq miRNA library kit (Qiagen, catalog number: 331502)
QIAseq miRNA Index kit IL UDI (e.g., UDI-B-96, Qiagen, catalog number: 331625)
10× TBE Buffer (Thermo Fisher Scientific, catalog number: AM9863)
1:20 dilution of MNase (100,000 gel units/mL) (see Recipes)
50 mM 5-bromouridine (see Recipes)
IP buffer (see Recipes)
Software
Removal of UMI duplicates: UMI-tools (Smith et al., 2017)
Removal of adaptor sequences: Cutadapt (Martin, 2011)
Genomic alignment of sequencing reads: Bowtie2 (Langmead and Salzberg, 2012)
BAM file processing: Samtools (Li et al., 2009)
Integrative Genomics Viewer (IGV) (Robinson et al., 2011)
Conversion of BAM files to BED files: bedtools (Quinlan and Hall, 2010)
Conversion of bedGraph files to BigWig: bedGraphToBigWig (Kent et al., 2010)
Equipment
1.5 mL microcentrifuge magnet stand (e.g., Thermo Fisher Scientific, catalog number: 12321D)
Centrifuge 5430 R (e.g., Eppendorf, catalog number: 022620601)
Thermal cycler (e.g., Bio-Rad, model: C1000 Touch, catalog number: 1851148)
Qubit fluorometer (Thermo Fisher Scientific, catalog number: Q33238)
Procedure
Labeling of nascent transcripts
The following steps achieve labeling of nascent transcripts with bromouridine to enable isolation of these transcripts via immunoprecipitation in Section D. 5-bromouridine is incorporated by all RNA polymerases actively transcribing during incubation of cells with 5-bromouridine.
Grow cells in 10 cm culture plates in a total of 10 mL of culture medium. For section C, 250 μg of total RNA is required. Scale the number of plates according to the type of cell line and yield from the protocol of total RNA extraction. For HEK293 cells, use a total of 3 × 10 cm plates or a single 15 cm plate per condition (~20 × 106 cells).
When the cell population has reached approximately 70% confluency, remove the cell medium and replace it with 10 mL of fresh cell medium containing 400 μL of 50 mM 5-bromouridine (2 mM 5-bromouridine final concentration). If using a 15 cm plate, scale up accordingly by using 20 mL of fresh cell medium containing 800 μL of 50 mM 5-bromouridine.
Place plate with cells in a humidifying incubator at 37 °C and 5% CO2 for 30 min (see Note 1).
Obtain a no-bromouridine negative control sample to rule out unspecific enrichment of RNA after immunoprecipitation. Perform the labeling protocol with the number of plates used per condition, excluding 5-bromouridine from the fresh cell medium in step A2.
Remove cell medium, add 1 mL of TRIzol reagent to dissociate cells from the culture plate, and transfer to a 1.5 mL microcentrifuge tube. If using a 15 cm plate, harvest in at least 3 mL of TRIzol reagent.
Pipette the lysate up and down to homogenize.
The following sections should be performed with special care to not contaminate RNA samples, which could result in RNA degradation by RNases. Use RNase-free certified plasticware and filter tips.
Total RNA extraction
Add 0.2 mL of chloroform per 1 mL of TRIzol lysate, securely cap the tube, and thoroughly mix by shaking for ~30 s.
Centrifuge the sample at 12,000× g for 15 min at 4 °C.
Transfer the aqueous phase containing the RNA to a new 1.5 mL microcentrifuge tube by angling the tube at 45° and pipetting the solution out.
Add an equal volume of isopropanol to the aqueous phase, mix by pipetting up and down, and incubate for 10 min at 4 °C.
Centrifuge at 12,000× g for 10 min at 4 °C.
Total RNA precipitate forms a white, gel-like pellet at the bottom of the tube.
Remove the solution by pipetting. Avoid removing the RNA pellet, which should be located below the hinge of the microcentrifuge tube as a white pellet.
Gently add 1 mL of ice-cold 80% ethanol and centrifuge at 12,000× g for 5 min at 4 °C.
Remove the 80% ethanol solution by pipetting out and pulse-spin the 1.7 mL microcentrifuge tube in a mini-centrifuge, to bring any residual ethanol from the sides of the tube down to the bottom.
Using a P20 pipette and tip, remove the remaining solution.
Resuspend the RNA pellet in 100 μL of RNase-free water, or in 300 μL if starting from a 15 cm plate.
Quantify the concentration of RNA using a Nanodrop spectrophotometer.
We suggest ruling out RNA fragmentation during the extraction of total RNA by analyzing RNA quality by your method of choice (i.e., Bioanalyzer, TapeStation, or RNA gel electrophoresis analysis). For RNA gel electrophoresis, we load 2 μL of the total RNA sample and 3 μL of GeneRuler 1 kb Plus DNA ladder into a 1% agarose gel in 1× TBE buffer and apply 140 V for 1 h (Figure 2A) (see Note 2). Store the RNA samples in a freezer at -80 °C or continue with Section C.
Figure 2. Gel electrophoresis of MNase digested total RNA. Nascent transcripts of HEK293 cells were pulse-labeled with 5-bromouridine for 30 min and total RNA was extracted. Shown is the purified total RNA sample before (A) and after fragmentation with MNase (B), for three biological replicates.
RNA fragmentation with MNase
Fragmentation of total RNA with MNase before immunoprecipitation results in enrichment of only the region of pre-mRNA molecules that is labeled with 5-bromouridine. In addition, MNase digestion creates RNA molecules of a suitable size for library preparation in Section G.
The procedures described in Sections C and D have been performed uninterrupted. Before starting, consider that this protocol does not include a stopping point between these sections. If a stopping point is necessary, we propose to complete step C4 in this section and place the fragmented RNA on ice until this sample is used in step D6 within 24 h.
Mix 250 μg of total RNA, 25 μL of 5 M NaCl, and 50 μL of 10× MNase reaction buffer in a 1.7 mL tube and bring to a total volume of 500 μL with nuclease-free water. Mix by pipetting up and down.
Incubate samples at 37 °C for 5 min.
Add 5 μL of a 1:20 dilution of MNase (100,000 gel units/mL), mix by pipetting up and down, and incubate for 1 min.
Stop the reaction by adding 10 μL of 0.5 M EGTA. Mix well and place sample on ice until completing step D5.
Evaluate RNA fragmentation by performing a gel electrophoresis using 2–4 μL of fragmented RNA (1–2 μg). RNA fragments should range from ~1.5 kb to ~100 bp in length. (Figure 2B).
Immunoprecipitation of nascent transcripts
The following steps enrich bromouridine-labeled nascent RNA transcripts via specific binding to a mouse BrdU antibody conjugated to protein G magnetic beads while washing away unlabeled RNA.
Wash protein G magnetic beads with IP buffer. Transfer 50 μL of well-mixed protein G magnetic beads–containing solution into a 1.7 mL tube and follow the steps below to wash the beads:
Place the 1.7 mL tube on the magnet stand to immobilize the beads.
When the solution has cleared completely and all beads are immobilized on the wall of the tube, remove the supernatant by pipetting out and add 500 μL of IP buffer.
Remove the tube from the magnet stand and resuspend beads by pipetting up and down gently until all beads are dissociated and no clumps of beads are observed. Place back on the magnet stand.
When the solution has cleared completely and all beads are immobilized on the wall of the tube, remove the supernatant by pipetting out.
Add 1 mL of IP buffer to the washed beads and remove from the magnet stand.
Conjugate mouse anti-BrdU antibody to protein G magnetic beads. Add 8 μL of antiBrdU antibody (4 μg) to the 1 mL of IP buffer containing the protein G magnetic beads, mix by pipetting up and down, and incubate the sample at 4 °C rotating end-to-end for 1 h. If the stopping point proposed in section C is included, this incubation can be performed overnight.
Wash conjugated anti-BrdU antibody and beads with IP buffer by performing the wash procedure described in step D1 (a–d) three times.
Add 1 mL of IP buffer to the beads and remove tube from the magnet stand.
Add the fragmented RNA from step C4 to the tube containing the beads, mix by pipetting up and down, and incubate the sample at 4 °C rotating end-to-end for 1 h (see Note 3).
Wash beads with IP buffer by performing the wash procedure described in step D1 (a–d) three times. Leave the tube on the magnet stand after the last wash.
Without disturbing the beads, add 500 μL of RNase-free 1× PBS, incubate for 30 s, and remove the supernatant by pipetting out.
Remove tube from the magnet stand, add 20 μL of nuclease-free water, resuspend beads by pipetting up and down, and transfer to a PCR tube.
Place the tube on a thermal cycler and run the following program to elute the antibody and nascent transcripts from the beads: 90 °C for 5 min, 12 °C for 30 s, with heated lid at 105 °C.
Vortex and transfer the sample to a 1.5 mL tube.
Place the 1.5 mL tube on the magnet stand and, when the solution has cleared, transfer the supernatant into a clean 1.5 mL tube.
Measure the RNA concentration of the sample using the QubitTM RNA High Sensitivity Assay kit. The expected concentration of specific enrichment of nascent transcript from HEK293 cells following this protocol is ~9–15 ng/μL with a total RNA yield of ~180–300 ng. The no-bromouridine negative control sample should not contain detectable signal by the Qubit Assay (< 0.2 ng/μL). Detection of RNA in this negative control suggests low efficiency of the washes after the immunoprecipitation.
Repair of MNase-fragmented RNA 3′-end
RNA fragmentation by MNase creates 3′ phosphate groups (Alexander et al., 1961). These can be converted to 3′OH RNA ends by T4 polynucleotide kinase, required for ligation to a 3′ adaptor during library preparation in section G. This reaction is performed in the absence of ATP, precluding installation of non-native 5′-PO4 groups on RNA molecules by this enzyme.
Transfer 150 ng of nascent transcripts into a 1.5 mL tube, add 5 μL of 10× reaction buffer A, add RNase-free water to bring the solution up to 49 μL, and mix by pipetting up and down.
Add 1 μL of T4 polynucleotide kinase (10 U) and incubate at 30 °C for 30 min.
In-column RNA purification
Purify the RNA from the in vitro reaction using any affinity micro-column purification protocol that elutes the RNA in a small quantity (< 20 μL). We used the RNA Clean & ConcentratorTM-5 cleanup kit described below.
Add two volumes of RNA binding buffer to each sample and mix by pipetting up and down.
Add an equal volume of ethanol (95%–100%) and mix by pipetting up and down.
Transfer the sample to the Zymo-SpinTM IC column in a collection tube and centrifuge at 12,000× g for 30 s. Discard the flowthrough.
Skip the DNase I treatment step, add 400 μL of RNA prep buffer to the column, and centrifuge at 12,000× g for 30 s. Discard the flowthrough.
Add 700 μL of RNA wash buffer to the column and centrifuge at 12,000× g for 30 s. Discard the flowthrough.
Add 400 μL of RNA wash buffer to the column and centrifuge at 12,000× g for 1 min. Ensure complete removal of the wash buffer. Carefully, transfer the column into a RNase-free tube.
Add 15 μL of DNase/RNase-free water directly to the column matrix and centrifuge at 12,000× g for 1 min to elute the RNA.
Measure the RNA concentration of the sample using the QubitTM RNA High Sensitivity Assay kit (expected to be approximately 6 ng/μL). Continue to section G or freeze the RNA sample at -80 °C.
Preparation of 5′-PO4 Bru-Seq libraries
Preparation of 5′-PO4 Bru-Seq libraries can be achieved using a library preparation protocol to detect miRNAs. In this protocol, we employed the QIAseq miRNA library kit, including 10-base UMIs. Transcripts with 5′ ends other than a monophosphate, including cap structures, are excluded because they cannot be ligated to the 5′ adaptor. Follow the steps in the manufacturer’s protocol with the specific conditions described below:
Prepare reagents required for the 3′ ligation reactions. Thaw QIAseq miRNA NGS 3′ adapter, QIAseq miRNA NGS 3′ buffer, 2× miRNA ligation activator, and nuclease-free water at room temperature (15–25 °C). Mix each solution by flicking the tubes. Centrifuge the tubes briefly to collect any residual liquid from the sides of the tubes and keep at room temperature.
Use 10–50 ng of nascent RNA transcripts using a 1:5 dilution of the 3′ adapter according to the QIAseq miRNA library kit Table 5 (Dilution of the QIAseq miRNA NGS 3′ Adapter).
On ice, prepare the 3′ ligation reaction according to Table 1. Briefly centrifuge, mix by pipetting up and down 15–20 times, and centrifuge briefly again.
Table 1. Setup of 3′ ligation reactions
Components Volume/reaction
Nuclease-free water Variable
QIAseq miRNA NGS 3′ adapter 1 μL
QIAseq miRNA NGS RI 1 μL
QIAseq miRNA NGS 3′ ligase 1 μL
QIAseq miRNA NGS 3′ buffer 2 μL
2x miRNA ligation activator 10 μL
Template RNA (added in step G4) Variable
Total volume 20 μL
Add template RNA to each tube containing the 3′ ligation master mix. Briefly centrifuge, mix by pipetting up and down 15–20 times, and centrifuge briefly again.
Incubate for 1 h at 28 °C.
Incubate for 20 min at 65 °C.
Hold at 4 °C for at least 5 min.
Prepare reagents required for the 5′ ligation reactions. Thaw QIAseq miRNA NGS 5′ adapter and QIAseq miRNA NGS 5′ buffer at room temperature. Mix by flicking the tube. Centrifuge the tube briefly to collect residual liquid from the sides of the tube and keep at room temperature.
Use a 1:2.5 dilution of the 5′ adapter according to the QIAseq miRNA library kit Table 7 (Dilution of the QIAseq miRNA NGS 5′ Adapter).
On ice, prepare the 5′ ligation reaction according to Table 2, adding the components in the order listed. Briefly centrifuge, mix by pipetting up and down 10–15 times, and centrifuge briefly again.
Table 2. Setup of 5′ ligation reactions
Component Volume/reaction
3′ ligation reaction (already in the tube) 20 μL
Nuclease-free water 15 μL
QIAseq miRNA NGS 5′ buffer 2 μL
QIAseq miRNA NGS RI 1 μL
QIAseq miRNA NGS 5′ ligase 1 μL
QIAseq miRNA NGS 5′ adapter 1 μL
Total volume 40 μL
Incubate for 30 min at 28 °C.
Incubate for 20 min at 65 °C.
Hold at 4 °C and proceed immediately to step G14.
Prepare reagents required for the reverse transcription reactions. Thaw QIAseq miRNA NGS RT initiator, QIAseq miRNA NGS RT buffer, and QIAseq miRNA NGS RT primer at room temperature. Mix by flicking the tube. Centrifuge the tubes briefly to collect residual liquid from the sides of the tubes and keep at room temperature.
Add 2 μL of QIAseq miRNA NGS RT initiator to each tube. Briefly centrifuge, mix by pipetting up and down 15–20 times, and centrifuge briefly again.
Incubate the tubes as described in Table 3.
Table 3. Incubation of tubes with QIAseq miRNA NGS RT initiator
Time Temperature
2 75 °C
2 70 °C
2 65 °C
2 60 °C
2 55 °C
5 37 °C
5 25 °C
∞* 4 °C
*Hold until setup of the RT reaction
Use a 1:5 dilution of the QIAseq miRNA NGS RT primer according to the QIAseq miRNA library kit Table 10 (Dilution of the QIAseq miRNA NGS RT Primer).
On ice, prepare the reverse transcription reaction according to Table 4. Briefly centrifuge, mix by pipetting up and down 12 times, and centrifuge briefly again.
Table 4. Setup of reverse transcription reactions
Component Volume/reaction
5′ ligation reaction + QIAseq miRNA NGS RT initiator (already in the tube) 42 μL
QIAseq miRNA NGS RT primer 2 μL
Nuclease-free water 2 μL
QIAseq miRNA NGS RT buffer 12 μL
QIAseq miRNA NGS RI 1 μL
QIAseq miRNA NGS RT enzyme 1 μL
Total volume 60 μL
Incubate for 1 h at 50 °C.
Incubate for 15 min at 70 °C.
Hold at 4 °C for at least 5 min.
Prepare QIAseq miRNA NGS beads (QMN beads). Thoroughly vortex QIAseq beads and QIAseq miRNA NGS bead binding buffer to ensure that the beads are in suspension and homogenously distributed. Do not centrifuge the reagents. Important: QIAseq beads need to be homogenous. This necessitates working quickly and thoroughly resuspending the beads immediately before use. If a delay in the protocol occurs, simply vortex the beads again.
Carefully add 400 μL of QIAseq beads (bead storage buffer is viscous) to a 2 mL microfuge tube. This quantity of beads is sufficient to perform “Protocol: cDNA Cleanup” and the cleanup associated with library amplification for one sample. Briefly centrifuge and immediately separate beads on a magnet stand.
When beads have fully migrated, carefully remove and discard the supernatant.
Remove the tube from the magnet stand and carefully pipette (buffer is viscous) 150 μL of QIAseq miRNA NGS bead binding buffer onto the beads. Thoroughly vortex to completely resuspend the bead pellet. Briefly centrifuge and immediately separate the beads on a magnet stand.
When beads have fully migrated, carefully remove and discard the supernatant.
Remove the tube from the magnet stand and carefully pipette 400 μL of QIAseq miRNA NGS bead binding buffer onto the beads (buffer is viscous). Thoroughly vortex to completely resuspend the bead pellet. Preparation of the QMN beads is now complete. If the beads will not be used immediately, store them on ice or at 2–8 °C.
Perform a cDNA cleanup. Centrifuge the tubes containing the cDNA reactions and add 143 μL of QMN beads to tubes containing the cDNA reactions. Vortex for 3 s and centrifuge briefly.
Incubate for 5 min at room temperature.
Place the tubes on a magnet stand for ~4 min or until the beads have fully migrated.
Discard the supernatant and keep the beads.
With the beads still on the magnet stand, add 200 μL of 80% ethanol. Immediately remove and discard the ethanol wash.
Repeat the wash by adding 200 μL of 80% ethanol. Immediately remove and discard the second ethanol wash. Important: completely remove all traces of ethanol after the second wash. Briefly centrifuge and return the tubes to the magnetic stand. Remove the ethanol with a 200 μL pipette first, and then use a 10 μL pipette to remove any residual ethanol.
With the beads still on the magnetic stand, air-dry at room temperature for 10 min. Residual ethanol can hinder amplification efficiency in the subsequent library amplification reactions. Depending on humidity, extended drying time may be required.
With the beads still on the magnetic stand, elute the DNA by adding 17 μL of nuclease-free water to the tubes. Subsequently close/cover and remove the tubes/plates from the magnetic stand.
Carefully pipette up and down until all the beads are thoroughly resuspended, briefly centrifuge, and incubate at room temperature for 2 min.
Return the tubes to the magnetic stand for ~2 min or until the beads have fully migrated. The completed cDNA cleanup product can be stored at -20 °C; alternatively, continue with the steps below.
Prepare reagents required for the library amplification reactions. Thaw QIAseq miRNA NGS library buffer, QIAseq miRNA NGS ILM library forward primer, and required index primer(s). Mix by flicking the tube. Centrifuge the tubes briefly to collect residual liquid from the sides of the tubes.
On ice, prepare the library amplification reaction according to Table 5. Briefly centrifuge, mix by pipetting up and down 12 times, and centrifuge briefly again.
Table 5. Setup of library amplification reactions when using tube indices
Component Volume/reaction
Product from “Protocol: cDNA Cleanup” 15 μL
QIAseq miRNA NGS library buffer 16 μL
HotStarTaq DNA polymerase 3 μL
QIAseq miRNA NGS ILM library forward primer 2 μL
QIAseq miRNA NGS ILM IPD1 through IPD48 (Index Primer) 2 μL
Nuclease-free water 42 μL
Total volume 80 μL
Program the thermal cycler according to Table 6.
Table 6. Library amplification protocol
Step Time Temperature
Hold 15 min 95 °C
3-step cycling (18 cycles)
Denaturation 15 s 95 °C
Annealing 30 s 60 °C
Extension 15 s 72 °C
Hold 2 min 72 °C
Hold ∞ 4 °C
Place the library amplification reaction in the thermal cycler and start the run. Upon completion of the protocol, hold at 4 °C for at least 5 min.
Add 75 μL of QMN beads to tubes. Ensure the QMN beads are thoroughly mixed at all times. This necessitates working quickly and resuspending the beads immediately before use. If a delay in the protocol occurs, simply vortex the beads.
Briefly centrifuge the 80 μL library amplification reactions and transfer 75 μL to the tubes containing the QMN beads. Vortex for 3 s and briefly centrifuge.
Incubate for 5 min at room temperature.
Place tubes on a magnet stand for approximately 4 min or until the beads have fully migrated.
Keep the supernatant and transfer 145 μL of the supernatant to new tubes. Discard the tubes containing the beads. Important: do not discard the supernatant at this step.
To the 145 μL supernatant, add 130 μL of QMN beads. Vortex for 3 s and briefly centrifuge.
Incubate at room temperature for 5 min.
Place the tubes on a magnet stand until beads have fully migrated.
Discard the supernatant and keep the beads.
With the beads still on the magnet stand, add 200 μL of 80% ethanol. Immediately remove and discard the ethanol wash.
Repeat the wash by adding 200 μL of 80% ethanol. Immediately remove and discard the second ethanol wash. It is important to completely remove all traces of the ethanol wash after the second wash. Briefly centrifuge and return the tubes to the magnetic stand. Remove the ethanol with a 200 μL pipette first, and then use a 10 μL pipette to remove any residual ethanol.
With the beads still on the magnetic stand, air-dry at room temperature for 10 min.
With the beads still on the magnetic stand, elute the DNA by adding 17 μL of nuclease-free water to the tubes. Subsequently close and remove the tubes from the magnetic stand.
Carefully pipette up and down until all beads are thoroughly resuspended; briefly centrifuge and incubate at room temperature for 2 min.
Place the tubes on the magnetic stand for ~2 min (or until beads have cleared).
Transfer 15 μL of eluted DNA to new tubes. This is the 5′-PO4 Bru-Seq sequencing library. Store sequencing libraries at -20 °C.
Submit libraries for Illumina sequencing. It is recommended to use paired-end sequencing. The 5′ end of the sequenced Read 1 informs on the position of the ligation event (5′-PO4 RNA end) and the first 12 nucleotides at the 5′ end of Read 2 inform on the identity of the UMI.
Data analysis
Extract UMIs from the Illumina sequencing run files (e.g., Read1.fastq, Read2.fastq) and obtain a new Read1.UMI.fastq file using UMI-tools (see Note 4).
umi_tools extract \
-I Read2.fastq \
--extract-method=string \
--bc-pattern=NNNNNNNNNNNN \
--read2-in=Read1.fastq \
--read2-out=Read1.UMI.fastq \
> Read2.UMI.fastq
Remove adaptor sequences using Cutadapt.
cutadapt \
-a 'AACTGTAGGCACCATCAAT' \
-A 'GATCGTCGGACTGTAGAACTCTGAAC' \
-o Read1.trimmed.fastq \
-p Read2.trimmed.fastq \
Read1.UMI.fastq \
Read2.UMI.fastq \
Use Bowtie (e.g., Bowtie 2) to align reads in the Read1.trimmed.fastq file to the human genome index (e.g., GRCh37/hg19).
bowtie2 \
-x bowtie_index \
-U Read1.trimmed.fastq \
-S Read1.sam
Convert the “Read1.sam” file to BAM format.
samtools view -S -b Read1.sam > Read1.bam
Sort the Read1.bam file.
samtools sort Read1.bam -o Read1.sorted.bam
Remove UMI duplicates using UMI-tools.
umi_tools dedup \
--method unique \
--read-length \
-I Read1.sorted.bam \
-S Read1.filtered.bam
Use the Read1.filtered.bam and the Read1.filtered.bam.bai files to visualize sequence reads on IGV. Figure 3 below shows sequenced reads mapped to the poly(A) site of the ACTB gene, where the first 5′ nucleotide in the sequenced read contains the single nucleotide coordinates of the 5′-PO4 RNA end in the nascent transcript immediately downstream of the poly(A) cleavage site (red arrows).
Figure 3. Integrative Genomics Viewer (IGV) screenshot showing sequenced 5′-PO4 Bru-Seq reads mapped to the 3′ end of the ACTB gene, where the localized 5′-monophosphorylated nucleotides in the nascent transcript are indicated by read arrows. The poly(A) signal is indicated by a red line above the sequence track.
Convert the “Read1.filtered.bam” file to BED format using bedtools.
bedtools bamtobed \
-i Read1.filtered.bam \
> Read1.filtered.bed
Collapse read coordinates in the “Read1.filtered.bed” file to the 5′ single nucleotide coordinate, which corresponds to the 5′-PO4 RNA single nucleotide coordinate, and create coverage BEDGRAPH files with strand specificity using the “chrom.sizes” file associated with the reference genome that was used for mapping.
bedtools genomecov \
-strand + \
-5 \
-i Read1.filtered.bed \
-bg chrom.sizes \
> Read1.5p.positive.bg
bedtools genomecov \
-strand - \
-5 \
-i Read1.filtered.bed \
-bg chrom.sizes \
> Read1.5p.negative.bg
Sort the output BEDGRAPH files.
sort -k1,1 -k2,2n Read1.5p.positive.bg \
> Read1.5p.sorted.positive.bg
sort -k1,1 -k2,2n Read1.5p.negative.bg \
> Read1.5p.sorted.negative.bg
Use bedGraphToBigWig to create BigWig files for visualization of genomic data on the University of California Santa Cruz (UCSC) Genome Browser.
bedGraphToBigWig \
Read1.5p.sorted.positive.bg \
chrom.sizes \
Read1.5p.positive.bw
bedGraphToBigWig \
Read1.5p.sorted.negative.bg \
chrom.sizes \
Read1.5p.negative.bw
Confirm enrichment of 5′-PO4 Bru-Seq signal by detection of 5′-PO4 RNA ends at microprocessor cleavage sites of the MIR17HG miRNA cluster (Figure 4A), at the 5′ end of the SNORA14B snoRNA (Figure 4B), the 5′ end of the primary tRNA transcript prior to RNaseP cleavage (Figure 4C, red arrow), and 5′-PO4 ends generated after cleavage at the poly(A) site on the ACTB gene (Figure 4D). 5′-PO4 signal can be further validated by detection of the 5′ end of the 47S rRNA precursor (Wang and Pestov, 2011) (Figure 4E, red arrow). Ribosomal signal can be visualized by mapping to the human ribosomal repeating unit reference sequence (GenBank accession no. U13369). Finally, relative quantification of peak intensities across libraries can be made by normalizing to the total number of mapped reads or by normalization to an appropriate internal control region [e.g., the mitochondrial genome (Cortazar et al., 2022)].
Figure 4. 5′-PO4 Bru-Seq maps known 5′-PO4 RNA ends of nascent transcripts. Shown are UCSC genome browser screenshots of 5′-PO4 Bru-Seq signal at: (A) the MIR17HG miRNA cluster, (B) the SNORA14B snoRNA, (C) a tRNA gene where the 5′ end of the primary tRNA transcript is indicated by a green arrow, and (D) the 3′ end of the ACTB gene where the poly(A) signal is underlined upstream of the poly(A) cleavage site. The peaks represent the nucleotide with the 5′-PO4 group in the RNA fragment downstream of the poly(A) cleavage site, and (E) the ribosomal RNA transcription unit zoomed in y-axis values (0–8,000), to visualize the 5′ end of the 47S rRNA precursor indicated by a black arrow. The y-axis indicates the number of mapped reads per one million of mitochondrial 5′-PO4 Bru-Seq sequenced reads.
Notes
Incubation of cells with 2 mM 5-bromouridine for a period of 30 min has been previously validated to result in mainly nascent transcript signal (Paulsen et al., 2013). Shorter incubation times can be performed (~20 min), although the yield of nascent transcripts is substantially decreased. Longer incubation times are not recommended, given that labeled nascent transcripts are processed, accumulate in the population of mature RNA, and contaminate nascent transcript signal.
The population of intact RNA molecules in high-quality samples show distinct bands that correspond to the 28S and 18S rRNA species, respectively. A fuzzy and faint band at the bottom of the gel corresponding to the population of tRNAs can also be visualized depending on the amount of RNA loaded (Figure 2A). The qualitative 28S/18S band intensity ratio value of 2 has been historically considered to indicate intact RNA.
Shorter incubations during immunoprecipitation of bromo-labeled RNA reduce the potential for RNA fragmentation.
By default, UMI_tools extracts UMIs from the “Read1.fastq” input file using argument “-I.” Given that the UMI is contained in Read 2, you can simply switch the input file provided for argument “--read2-in.”
Recipes
1:20 dilution of MNase (100,000 gel units/mL)
Mix 1 μL of MNase (2,000,000 gel units/mL) with 19 μL of 1× MNase reaction buffer.
50 mM 5-bromouridine
Dissolve 808 mg of 5-bromouridine in 50 mL of RNase-free 1× PBS and store at -20 °C.
IP Buffer
0.05% Triton X-100, 1 mM DTT, supplemented with Superase-InTM RNase inhibitor in RNase-free 1× PBS.
Mix 25 μL of Triton X-100, 50 μL of 1 M DTT, 50 μL of Superase-InTM RNase inhibitor, and 50 mL of RNase-free 1× PBS.
Acknowledgments
Michael A. Cortázar was a scholar of the University of Colorado at Denver RNA Bioscience Initiative. Labeling and immunoprecipitation steps in this protocol were modified from the Bru-Seq protocol (Paulsen et al., 2014). This work was supported by a NIH grant R35 GM144336 to D.B.
Competing interests
The authors declare no competing interests.
References
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4,829 | https://bio-protocol.org/en/bpdetail?id=4829&type=0 | # Bio-Protocol Content
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Peer-reviewed
Mouse Corneal Epithelial and Stromal Cell Isolation and Culture
YZ Yingnan Zhang
LZ Lijun Zhang
YL Yongqing Liu
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4829 Views: 1105
Reviewed by: Pilar Villacampa AlcubierreMarina Sánchez Petidier Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in Communications Biology Apr 2023
Abstract
Corneal epithelium and stroma are the major cellular structures for ocular protection and vision accuracy; they play important roles in corneal wound healing and inflammation under pathological conditions. Unlike human, murine corneal and stromal fibroblast cells are difficult to isolate for cell culture. In our laboratory, we successfully used an ex vivo culture procedure and an enzymatic procedure to isolate, purify, and culture mouse corneal epithelial and stromal fibroblast cells.
Key features
• Primary cell culture models of a disease are critical for cellular and molecular mechanism studies.
• Corneal tissues with the limbus contain stem cells to generate both epithelial and stromal cells.
• An ex vivo corneal culture provides a constant generation of primary corneal cells for multiple passages.
• The isolated cells are validated by the corneal epithelial cell markers Krt12 and Cdh1 and the stromal fibroblast marker Vim.
Keywords: Mouse cornea Epithelial cells Keratocytes Corneal cell isolation Primary cell culture Ex vivo culture
Background
The cornea is a transparent avascular tissue; it acts as a barrier to protect the eye and has a 2/3 focal power to project an image onto the retina. Mouse cornea consists of three cellular layers: the epithelium, the stroma, and the endothelium (Figure 1). The epithelium consists of a differentiated superficial squamous cell, a transit wing cell, and a proliferative basal cell type, and is the outmost ocular tissue that contains 80% corneal cells. The innermost corneal tissue is the endothelium, consisting of a single unrenewable cell sheet. The epithelium layer and the endothelium layer are separated from the stroma by the Bowman’s membrane and the Descemet’s membrane, respectively. The stroma is the thickest and toughest part of the cornea in between the epithelium and the endothelium. Although the stroma contains only approximately 20% of the corneal cells, it occupies two thirds of the corneal space. Both epithelial and stromal cells are embedded in a tough collagen-rich extracellular matrix (ECM) [1]. As the epithelial cells can be regenerated by a group of stem cells, also known as limbal epithelial stem cells, at the limbus (the border area between the cornea and the sclera) for cellular homeostasis and wound repair, it is relatively easy to isolate and culture them [2]. However, the corneal stromal fibroblast (keratocyte) may be regenerated at a relatively slower pace; its regeneration is supposed to be either local and/or come from the progenitor at the limbus region. Isolation and culture of corneal cells, i.e., epithelial cells and stromal keratocytes, are critical for the study of corneal diseases, particularly wound healing [3] and neovascularization [4], as well as their underlying molecular mechanisms [5]. Human corneal primary epithelial and stromal cells and murine primary corneal epithelial cells are commercially available; however, no murine primary stromal fibroblast cells can be purchased. It is difficult to isolate and culture murine primary corneal cells, particularly murine keratocytes. A protocol to isolate and culture murine keratocytes is available for newborn pups, where the separation of the stroma from the epithelium is very difficult and the resultant cell identification was not verified by a specific marker [6]. We have developed a relatively simple and reliable method to isolate both murine corneal epithelial cells and keratocytes, and culture them in vitro for multiple passages.
Figure 1. Mouse corneal structure and cell types
Materials and reagents
Biological materials
Animals: 4–8-week-old B6 mice (The Jackson Laboratory, catalog number: 000664)
Consumable materials
Cell culture plate: 12-well cell culture plates (Thermo Fisher Scientific, catalog number: 150628)
Eppendorf tubes (Thermo Fisher Scientific, catalog number: 3451)
Tipone® pipette tips (USA Scientific, catalog number: 1126-7810)
Pipettes (Corning, Falcon®, catalog number: 357543)
Cryopreservation vials (Corning, catalog number: 430659)
Reagents
Ethanol
Phosphate-buffered saline (PBS) (Sigma-Aldrich, catalog number: P4417-100TAB)
Penicillin-streptomycin (5,000 U/mL) (Thermo Fisher Scientific, catalog number: 15070063)
Dulbecco’s modified Eagle’s medium (DMEM, high glucose) (Thermo Fisher Scientific, catalog number: 11965)
Fetal bovine serum (FBS) (GE Healthcare, HyClone, catalog number: SH30071.03)
Gelatin (Sigma-Aldrich, catalog number: G1890-100G)
Collagenase A (Millipore Sigma, catalog number: 10103578001)
Trypsin ethylenediaminetetraacetic acid (EDTA) solution (Mediatech, Cellgro®, catalog number: 25-053-CI)
Common epithelial marker E-cadherin (Cdh1) (BD Transduction Lab, catalog number: 610181)
Specific epithelial marker Keratin 12 (Krt12) (Abclonal, catalog number: A9642)
Fibroblast marker vimentin (Sigma, catalog number: 17-10195)
100× penicillin-streptomycin solution (Thermo Fisher Scientific, catalog number: 15140122)
Corneal epithelial cell basal medium (ATCC, catalog number: PCS-700-030)
Corneal epithelial cell growth kit (ATCC, catalog number: PCS-700-040)
Solutions
Corneal epithelial cell culture medium (see Recipes)
Corneal stromal fibroblast culture medium (see Recipes)
0.1% gelatin solution (see Recipes)
Recipes
Corneal epithelial cell culture medium
Prepare the complete corneal epithelial growth medium according to the manufacturer’s instructions. The final concentration for each component in ATCC complete corneal epithelial growth medium is as follows:
5 μg/mL Apo-transferrin
1.0 μM epinephrine
0.4% extract P
100 ng/mL hydrocortisone
6 mM L-glutamine
5 μg/mL recombinant human insulin
CE Growth Factor: proprietary formulation
Corneal stromal fibroblast culture medium
450 mL of DMEM
50 mL of FBS
5 mL of penicillin/streptomycin (5,000 U/mL)
0.1% gelatin solution
0.1 g gelatin powder
100 mL of PBS
Autoclave and store at 4 °C for three months.
Equipment
Mcpherson-Vannas curved iris scissors (Storz Ophthalmic Instruments, catalog number: E3347)
Castroviejo suturing forceps 0.12 mm (Storz Ophthalmic Instruments, catalog number: E1796)
Martinez double ended corneal dissector (Storz Ophthalmic Instruments, catalog number: E3001)
Algerbrush II (Katena Products, Inc., Denville, NJ)
Cell culture incubator (Thermo Fisher Scientific, Thermo Scientific, model: 3250)
Water bath (VWR, model: 1545)
Pipettes (Eppendorf)
Pipet-aid (Drummond)
Benchtop centrifuge (Beckman Coulter, model: Allegra® X-15R)
Lab rotating shaker (Barnstead Thermolyne LabQuake, model: 4152110)
Inverted microscope (Nikon, Eclipse TS100)
Confocal microscope (Nikon, Eclipse Ti)
40 μm nylon net filter (Fisher Scientific, Leicestershire, UK)
Procedure
Corneal epithelial cell isolation and culture
All materials used in this procedure must be sterile or autoclaved to prevent contamination (Figure 2).
Figure 2. Surgery tools used in the procedure must be autoclaved
Euthanize mice by carbon dioxide followed by cervical dislocation to confirm death.
Enucleate both eyeballs using tweezers and place them in 1% penicillin-streptomycin PBS (Figure 3A).
Carefully trim off the cornea of full thickness from the eyeball with a pair of surgery scissors under a microscope (Figure 3B).
Thereafter, cut the entire cornea with the limbus quarterly into a butterfly tie-like shape using scissors (Figure 3C) and place it in 1% penicillin-streptomycin PBS.
Figure 3. The procedure of mouse corneal isolation. (A) The eyeball is enucleated and placed in PBS buffer. (B) The entire cornea with the limbus is cut off, and (C) quarterly divided into a butterfly tie-like shape.
To coat a 12-well culture plate, prepare 0.1% gelatin solution (Recipe 3) and sterilize it through filtration. Add 100 μL of this coating solution to each well and leave at room temperature for 10 min.
Aspirate the coating solution from the plate and place 2–4 corneas with the epithelium side down on each well in order to have enough epithelial cells moving out of the tissues.
Gently press the tissue piece with tweezers to make the tissue fully contact the surface of the plate; add 100 μL of corneal epithelial culture medium (Recipe 1) to cover the entire corneal tissue.
Incubate the plate with cultured corneas at 37 °C under 100% humidity and 5% CO2.
On the second day, very slowly add 500 μL of the corneal epithelial culture medium along the well wall to prevent the tissue from floating.
When most epithelial cells move out of the epithelial tissue onto the plate, approximately at day 7 after seeding (Figure 4A–4D), remove the corneal tissue with tweezers and add 1 mL of fresh corneal epithelial culture medium.
Replace the old medium with 1.5 mL of new medium every 3–4 days until cell confluence. Reuse the ex vivo corneal tissue at least five times.
After removing the culture medium, detach the confluent murine corneal epithelial cells from the plate and from each other by incubating them with 0.3 mL of 0.15% trypsin EDTA solution at 37 °C for 3 min.
Split the cells once at 1:2 ratio into two separate wells with 3 mL of the corneal stromal fibroblast culture medium (Recipe 2) overnight to inactivate the trypsin and help cells adhere to the plate.
On the next day, replace the corneal stromal fibroblast culture medium with 1.5 mL of corneal epithelial culture medium.
The corneal epithelial cells are confirmed by their small, compact, round, or elliptical morphology (Figure 4E) and immunostaining (Figure 4F–4G) with the corneal epithelial marker keratin 12 (Krt12) and the common epithelial marker E-cadherin (Cdh1).
Figure 4. Mouse corneal epithelial cell isolation and culture. The isolated mouse cornea is placed upside down on a cell culture plate coated with 0.1% gelatin for (A) 0, (B) 1, (C) 3, and (D) 7 days. (E) The monolayer-cultured corneal epithelial cells and their immunofluorescence with (F) the corneal epithelial marker keratin 12 (Krt12) and (G) the common epithelial maker E-cadherin (Cdh1).
Corneal stroma cell isolation and culture
Euthanize mice by carbon dioxide followed by cervical dislocation to confirm death.
Remove the epithelial layer down to the basement membrane by mechanical scraping using an Algerbrush II and a cotton swab (Video 1 shows the head tip of Algerbrush II scrapping corneal epithelial cells into a cell debris mass; after the entire epithelium area is debrided, the cell debris mass is cleaned by a cotton swab).
Video 1. Debridement of the corneal epithelium using Algerbrush II
Enucleate both eyeballs and place them in 1% penicillin-streptomycin PBS.
Carefully trim off the cornea of full thickness from the eyeball with a pair of surgery scissors under a microscope (Figure 3B).
Thereafter, cut the entire cornea with the limbus quarterly into a butterfly tie-like shape (Figure 3C) and place it in 1% penicillin-streptomycin PBS.
Remove the endothelial layer using Martinez double ended corneal dissector (Video 2 shows that the stroma of cornea noticeably becomes clearer after the removal of the endothelium) and cut the remainder of the stromal tissue into small pieces using a pair of surgery scissors and a razor blade.
Video 2. Removal of the corneal endothelium using Martinez dissector
Further digest each minced corneal tissue in 0.5 mL of 0.1 mg/mL collagenase A in an Eppendorf tube at 37 °C for 1–2 h. Vortex the tube every 10 min until the digestion solution appears cloudy, a sign of thorough tissue digestion.
Filtrate the digested corneal tissue with a 40 μm nylon net filter to remove bigger debris and to collect the flowthrough stromal cells in a 15 mL tube.
Add the corneal stromal fibroblast culture medium to the flowthrough solution and centrifuge at 1,000× g for 5 min.
Decant the supernatant, resuspend the cell pellet in 3 mL of the corneal stromal fibroblast culture medium (Recipe 2), and place it in one well of a 6-well culture plate.
Refresh the medium every 3–4 days until cell confluence.
After removal of the culture medium, detach the confluent murine corneal stromal fibroblast cells from the plate and from each other by incubating with 0.3 mL of 0.15% trypsin solution at room temperature for 3 min.
Split the cells once at 1:2 ratio into two separate wells with 3 mL of the corneal stromal fibroblast culture medium to inactivate the trypsin and help cells adhere to the plate.
The corneal fibroblasts (keratocytes) are confirmed by their morphology (Figure 5A–5C) and immunostaining (Figure 5D) with the fibroblast marker vimentin.
Figure 5. Mouse corneal stromal fibroblast cell isolation and culture. The epithelium and the endothelium are first removed mechanically from the isolated mouse cornea; the remaining stroma is minced and then further digested with collagenase type A. Upon a passage through a nylon net filter, the isolated cells are cultured in a plate coated with 0.1% gelatin for (A) 1, (B) 3, and (C) 7 days. (D) Monolayer-cultured mouse corneal stromal cells stained with the fibroblast cell marker vimentin (Vim).
Validation of protocol
We used this protocol to isolate and culture mouse corneal epithelial and stromal cells for our previous studies and had reliable results [3].
General notes and troubleshooting
These protocols for murine corneal epithelial and stromal fibroblast cell isolation and culture are relatively simple and easy for research laboratories in need, though their isolation yield and proliferation rates in culture are still low. We do not know why it is so technically difficult. Compared to human, mouse life span is very short; we believe that the renewability of mouse primary ocular cells, such as RPE and Müller cells to our best knowledge, is probably not as strong as human primary ocular cells.
Acknowledgments
This protocol was recently adopted in the laboratory [4]. The work was supported by National Institute of General Medical Sciences (P20GM103453 to Y.L.), University of Louisville School of Medicine (E0819 to Y.L.), James Graham Brown Cancer Center of University of Louisville Directed Gift Pilot Project Program (G1779 to Y.L.), and the National Natural Science Foundation of China (82171032 L.Z.), the Natural Science Foundation of Liaoling Province (201800209, 201602210, and 20180550976 to L.Z.).
Competing interests
The authors declare no competing interests.
Ethical considerations
Use of animals in this protocol was conducted according to the policies and guidelines set forth by the Institutional Animal Care and Use Committee (IACUC) and approved by the University of Louisville, Kentucky, USA.
References
Espana, E. M. and Birk, D. E. (2020). Composition, structure and function of the corneal stroma. Exp. Eye Res. 198: 108137. doi: 10.1016/j.exer.2020.108137
Kobayashi, T., Yoshioka, R., Shiraishi, A. and Ohashi, Y. (2009). New technique for culturing corneal epithelial cells of normal mice. Mol. Vis. 15: 1589–1593. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2728571/
Zhang, Y., Do, K. K., Wang, F., Lu, X., Liu, J. Y., Li, C., Ceresa, B. P., Zhang, L., Dean, D. C., Liu, Y., et al. (2023). Zeb1 facilitates corneal epithelial wound healing by maintaining corneal epithelial cell viability and mobility. Commun. Biol. 6(1): e1038/s42003-023-04831-0. doi: 10.1038/s42003-023-04831-0
Jin, L., Zhang, Y., Liang, W., Lu, X., Piri, N., Wang, W., Kaplan, H. J., Dean, D. C., Zhang, L., Liu, Y., et al. (2020). Zeb1 promotes corneal neovascularization by regulation of vascular endothelial cell proliferation. Commun. Biol. 3(1): e1038/s42003-020-1069-z. doi: 10.1038/s42003-020-1069-z
Liang, W., Zhang, Y., Zhou, L., Lu, X., Finn, M. E., Wang, W., Shao, H., Dean, D. C., Zhang, L., Liu, Y., et al. (2022). Zeb1 regulation of wound-healing-induced inflammation in alkali-damaged corneas. iScience 25(4): 104038. doi: 10.1016/j.isci.2022.104038
Zhang, Y., Wang, Y. C., Yuka, O., Zhang, L. and Liu, C. Y. (2016). Mouse Corneal Stroma Fibroblast Primary Cell Culture. Bio Protoc 6(19): e1960. doi: 10.21769/bioprotoc.1960
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© 2023 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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Improve Research Reproducibility
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Applying AnchorWave to Address Plant Genome Alignment
SW Shuai Wang
QX Qinlin Xu
BS Baoxing Song
Published: Oct 5, 2023
DOI: 10.21769/BioProtoc.4830 Views: 338
Reviewed by: Yi Zheng Anonymous reviewer(s)
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Abstract
With the massive development of long-read sequencing technologies, more and more complex genomes are being assembled. Genome alignment is an essential step for the majority of genomics research. Sensitively and accurately aligning complex genomes is critical to gain information from those genomes. In this protocol, we introduce AnchorWave (Anchored Wavefront alignment), which uses conserved sequences as anchors to identify collinear blocks and performs global sequence alignment at the nucleotide level using a 2-piece affine gap cost strategy. Here, we give two examples including maize (Zea mays), sorghum (Sorghum bicolor), and different cultivar maize lines. Maize went through an extra round of whole-genome duplication evolutionary process compared to sorghum. Moreover, maize has abundant transposable elements (TEs), genome rearrangement, and gene losses. AnchorWave provides a significant improvement compared to previous methods when aligning plant genomes with dispersed repeats, active TEs, high sequence diversity, and whole-genome duplication.
Keywords: AnchorWave Genome alignment Whole-genome duplication Collinear regions Transposable elements
Background
Recent technological advancements lead to a huge number of genomes being sequenced. We propose to unlock the secret of genome by genome comparison. Genome alignment is a fundamental step of genomics analysis, providing a set of intermediate results for downstream analysis. However, with previous genome alignment tools, it is difficult to deal with long (> 50 bp) inserts and deletions (InDels) and whole-genome duplication (WGD) that have taken place between two plant genomes. In addition, plants have a lower gene length and more complex structure variations and chromosome rearrangement than mammals. Software developed by the mammal research community cannot sensitively and accurately align plant genomes. This protocol illustrates AnchorWave, which can deal well with dispersed repeats, active TEs, high sequence diversity, and WGD (Song et al., 2022). AnchorWave combines a collinear region identification approach with a 2-piece affine gap cost global alignment strategy. This software firstly identifies collinear regions with conserved anchors and then performs base-pair resolved global sequence alignment for each anchor and inter-anchor region. AnchorWave provides a significant improvement for base-pair-resolved plant genome alignment.
Software and datasets
Software
AnchorWave (Song et al., 2022; v1.0.1; https://github.com/baoxingsong/AnchorWave; MIT License)
Samtools (Li et al., 2009; v1.6; http://www.htslib.org)
minimap2 (Li, 2018; v2.17-r941; https://github.com/lh3/minimap2)
ggplot2 (Wickham, 2016; v3.3.5; https://ggplot2.tidyverse.org)
Input data
Reference genome in FASTA format and annotation in GFF(3) format
Query genome in FASTA format
Procedure
The process generally includes three steps:
1. Extract coding sequences (CDS) as anchors.
2. Lift over to the query and reference genome.
3. Perform WGD and other steps to visualize the relationship between two genomes.
Case study 1: Align two genomes with relocation variation, chromosome fusion, or WGD variation
In the first case study, we align the sorghum genome to the maize genome via AnchorWave for a step-by-step illustration.
Download the reference genome and the query genome.
As inputs, we use reference genome, query genome, and the reference genome annotation that are downloaded from public databases. Do not forget to decompress them.
#Bash
#Installation using conda
conda install -c bioconda -c conda-forge anchorwave
#Download and decompress genome and GFF file of reference genome
wget ftp://ftp.ensemblgenomes.org/pub/plants/release-34/fasta/\
zea_mays/dna/Zea_mays.AGPv4.dna.toplevel.fa.gz
wget ftp://ftp.ensemblgenomes.org/pub/plants/release-34/gff3/\
zea_mays/Zea_mays.AGPv4.34.gff3.gz
wget http://ftp.ensemblgenomes.org/pub/plants/release-54/fasta/\
sorghum_bicolor/dna/Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa.gz
gunzip *.gz
Check the collinearity and WGD.
Access the level of chromosome.
We learn that maize has ten chromosomes by looking into “Zea_mays.AGPv4.dna.toplevel.fa.fai” generated by Samtools and that sorghum also has ten chromosomes by looking into “Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa.fai.” We also learn the name and sequence length for each sequence record.
#Bash
#Access the level of chromosome and collineraity
samtools faidx Zea_mays.AGPv4.dna.toplevel.fa
samtools faidx Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa
less Zea_mays.AGPv4.dna.toplevel.fa.fai
less Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa.fai
Extract reference CDS as anchors and lift over reference and query genome.
At this step, we visually check the WGD difference and chromosome rearrangements between the reference genome and query genome. We use the gff2seq function of the AnchorWave software to extract reference CDS. This command takes the reference genome and reference annotation as inputs. The reference CDS are mapped to the reference genome and query genome by minimap2. x splice function represents long-read splice alignment, t represents the number of threads, k represents k-mer size, p represents minimal secondary-to-primary score ratio to output secondary mappings, and a represents the output file in SAM formation. We can use R to visualize the full-length CDS mapping result (see Data analysis). The alignmentToDotplot.pl script that is used to convert the SAM file into a table is available from GitHub (https://github.com/Bio-protocol/anchorwave_protocol/blob/master/workflow1-case-study1/alignmentToDotplot.pl).
#Bash
#Extract CDS
anchorwave gff2seq -i Zea_mays.AGPv4.34.gff3 -r Zea_mays.AGPv4.dna.toplevel.fa -o cds.fa
#Mapping reference CDS to reference genome
minimap2 -x splice -a -t 10 -k 12 -p 0.4 -N 20 Zea_mays.AGPv4.dna.toplevel.fa cds.fa > ref.sam
#lift over to query genome
minimap2 -x splice -a -t 10 -k 12 -p 0.4 -N 20 Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa cds.fa > cds.sam
# Visualization the result of collinearity and all anchors.
perl alignmentToDotplot.pl Zea_mays.AGPv4.34.gff3 cds.sam > cds.tab
#Use R to draw a dotplot.
library(ggplot2)
library(svglite)
#Transform Coordinates using follow function.
changetoM <- function ( position ){
position=position/1000000;
paste(position, "M", sep="")}
#Read gene position, belong to which chromosome and so on
data =read.table("cds.tab")
#Select all euchromosomes as factor.
data = data[which(data$V1 %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data = data[which(data$V3 %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data$V1 = factor(data$V1, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
data$V3 = factor(data$V3, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
#Using ggplot2 to plot a dotplot and beautify it.
figure1 <- ggplot(data=data, aes(x=V4, y=V2)) +geom_point(size=0.5, aes(color=V5)) +
facet_grid(V1 ~ V3, scales="free",space="free") +labs(x="sorghum", y="maize")+
scale_x_continuous(labels=changetoM) + scale_y_continuous(labels=changetoM) +
theme(axis.line = element_blank(),
panel.background = element_blank(),
panel.border = element_rect(fill=NA,color="black", size=0.5, linetype="solid"),
axis.text.y = element_text( colour = "black"),
legend.position='none',
axis.text.x = element_text(angle=300, hjust=0, vjust=1, colour = "black"))
png("figure1.png")
figure1
dev.off()
pdf("figure1.pdf")
figure1
dev.off()
svglite("figure1.svg")
figure1
dev.off()
Use AnchorWave to identify collinear regions and plot collinear anchors.
By looking at the above plot, there are chromosome fusions, translocation variations, and an unshared WGD. We use the proali function to identify collinear regions. Maize went through an extra WGD event than sorghum, so we set -R as 1 and -Q as 2. The reference and query genome sequence, reference genome annotation file, mapping CDS to reference, and query genome files in SAM format are used as inputs. In addition, novel anchors do not help to identify collinear blocks; by setting -ns, we do not identify novel anchors for this step. We use R package ggplot2 to plot the result. Draw the graph with query start location of blocks as x-axis and reference start location of blocks as y-axis (see Data analysis). The user needs to modify the chromosome names manually in the R code if different genomes are being analyzed.
#Bash
#Applying AnchorWave to extract collinear anchors
anchorwave proali -r Zea_mays.AGPv4.dna.toplevel.fa -i Zea_mays.AGPv4.34.gff3 -a cds.sam -as cds.fa -ar ref.sam -s \
Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa -n align1.anchors -R 1 -Q 2 -ns
#Using R to plot a dotplot.
library(ggplot2)
library(svglite)
changetoM <- function (position){
position=position/1000000;
paste(position, "M", sep="")}
data =read.table("align1.anchors", header=TRUE)
data = data[which(data$refChr %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data = data[which(data$queryChr %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data$refChr = factor(data$refChr, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
data$queryCh = factor(data$queryChr, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
figure2 <- ggplot(data=data, aes(x=queryStart, y=referenceStart))+
geom_point(size=0.5, aes(color=strand)) +
facet_grid(refChr~queryChr, scales="free", space="free") +
labs(x="sorghum", y="maize")+scale_x_continuous(labels=changetoM) +
scale_y_continuous(labels=changetoM) +
theme(axis.line = element_blank(),
panel.background = element_blank(),
panel.border = element_rect(fill =NA,color="black", size=0.5, linetype="solid"),
axis.text.y = element_text(colour = "black"),
legend.position='none',
axis.text.x = element_text(angle=300,hjust=0, vjust=0.5, colour = "black") )
png("figure2.png")
figure2
dev.off()
pdf("figure2.pdf")
figure2
dev.off()
svglite("figure2.svg")
figure2
dev.off()
Perform WGD.
To align the sorghum genome against the maize genome, we use the proali command of AnchorWave; as represents anchor sequence files, and i, r, and o represent input reference GFF(3), FASTA file, and output MAF file. a and ar represent SAM files generated by mapping conserved sequences to reference and query genome, s represents query genome file, and n represents output anchor files. We set parameters -R 1 -Q 2 to utilize the knowledge that the maize lineage has been through a WGD since its divergence with sorghum. We perform these steps using a workstation with random access memory (RAM) of 128 GB.
#Bash
anchorwave proali -i Zea_mays.AGPv4.34.gff3 \
-r Zea_mays.AGPv4.dna.toplevel.fa -a cds.sam -as cds.fa -ar ref.sam \
-s Sorghum_bicolor.Sorghum_bicolor_NCBIv3.dna.toplevel.fa \
-n align.anchors -o align.maf -t 1 -R 1 -Q 2 -f align1.f.maf
Case study 2: Align two maize genomes without translocation rearrangement while with inversions
In this case study, we align different maize genomes between B73 and Mo17 to illustrate genoAli function. We can directly align two genomes using the following steps if we know the relationship between two genomes.<
Download the reference genome and the query genome.
wget https://ftp.ensemblgenomes.ebi.ac.uk/pub/plants/release-34/gff3/zea_mays/Zea_mays.AGPv4.34.gff3.gz
wget https://ftp.ensemblgenomes.ebi.ac.uk/pub/plants/release-34/fasta/zea_mays/dna/Zea_mays.AGPv4.dna.toplevel.fa.gz
wget https://download.maizegdb.org/Zm-Mo17-REFERENCE-CAU-1.0/Zm-Mo17-REFERENCE-CAU-1.0.fa.gz
gunzip *.gz
# transform format
sed -i 's/>chr/>/g' Zm-Mo17-REFERENCE-CAU-1.0.fa
Extract CDS as anchors and lift over to the query and reference genome.
For AnchorWave, gff2seq function is used for extracting CDS sequences; r, i, and o represent input reference FASTA, GFF(3) file, and output files. For minimap2, x splice function represents long-read splice alignment, t represents the number of threads, k represents k-mer size, p represents minimal secondary-to-primary score ratio to output secondary mappings, and a represents the output file in SAM format. Other parameters are default. We can use R to visualize the full-length CDS mapping result (see Data analysis). We use the proali function to identify collinear regions and use R package ggplot2 to plot the result. Draw the graph with start location of query anchors as x-axis and start location of reference anchors as y-axis (see Data analysis).
#extract CDS as anchors
anchorwave gff2seq -i Zea_mays.AGPv4.34.gff3 -r Zea_mays.AGPv4.dna.toplevel.fa -o cds.fa
#map CDS to the reference genome and query genome
minimap2 -x splice -t 10 -k 12 -a -p 0.4 -N 20 Zm-Mo17-REFERENCE-CAU-1.0.fa cds.fa > cds.sam
minimap2 -x splice -t 10 -k 12 -a -p 0.4 -N 20 Zea_mays.AGPv4.dna.toplevel.fa cds.fa > ref.sam
# visualize all anchors
perl alignmentToDotplot.pl Zea_mays.AGPv4.34.gff3 cds.sam > maizecds.tab
#Use R to draw a dotplot.
library(ggplot2)
library(svglite)
changetoM <- function (position){
position=position/1000000;
paste(position, "M", sep="")
}
data =read.table("maizecds.tab")
data = data[which(data$V1 %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data = data[which(data$V3 %in% c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10")),]
data$V1 = factor(data$V1, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
data$V3 = factor(data$V3, levels=c("1", "2", "3", "4", "5", "6", "7", "8", "9", "10"))
figure3 <- ggplot(data=data, aes(x=V4, y=V2))+geom_point(size=0.5, aes(color=V5))+facet_grid(V1~V3, scales="free", space="free" ) +
labs(x="Mo17", y="B73")+scale_x_continuous(labels=changetoM) + scale_y_continuous(labels=changetoM) +
theme(axis.line = element_blank(),
panel.background = element_blank(),
panel.border = element_rect(fill=NA,color="black", size=0.5, linetype="solid"),
axis.text.y = element_text( colour = "black"),
legend.position='none',
axis.text.x = element_text(angle=300, hjust=0, vjust=1, colour = "black") )
png("figure3.png")
figure3
dev.off()
pdf("figure3.pdf")
figure3
dev.off()
svglite("figure3.svg")
figure3
dev.off()
#identify the collinear anchors using AnchorWave.
anchorwave genoAli -i Zea_mays.AGPv4.34.gff3 -as cds.fa -r Zea_mays.AGPv4.dna.toplevel.fa -a cds.sam -ar ref.sam -s Zm-Mo17-REFERENCE-CAU-1.0.fa -n anchors.anchors -IV
#Use R to draw a dotplot.
library(ggplot2)
library(svglite)
changetoM <- function (position){
position=position/1000000;
paste(position, "M", sep="")
}
data =read.table("align1.anchors", header=TRUE)
data = data[which(data$refChr %in% c("1", "2", "3", "4","5", "6", "7", "8", "9", "10")),]
data = data[which(data$queryChr %in% c("1", "2", "3", "4","5", "6", "7", "8", "9", "10")),]
data$refChr = factor(data$refChr, levels=c("1", "2", "3", "4","5", "6", "7", "8", "9", "10"))
data$queryCh = factor(data$queryChr, levels=c("1", "2", "3", "4","5", "6", "7", "8", "9", "10"))
figure4 <- ggplot(data=data, aes(x=queryStart, y=referenceStart))+
geom_point(size=0.5, aes(color=strand)) +
facet_grid(refChr~queryChr, scales="free", space="free") +
labs(x="Mo17", y="B73")+scale_x_continuous(labels=changetoM) +
scale_y_continuous(labels=changetoM) +
theme(axis.line = element_blank(),
panel.background = element_blank(),
panel.border = element_rect(fill =NA,color="black", size=0.5, linetype="solid"),
axis.text.y = element_text( size=5,colour = "black"),
legend.position='none',
axis.text.x = element_text(angle=300,size=6,hjust=0, vjust=0.5, colour = "black"))
png("figure4.png")
figure4
dev.off()
pdf("figure4.pdf")
figure4
dev.off()
svglite("figure4.svg")
figure4
dev.off()
Perform WGD.
For AnchorWave, genoAli function is used for whole-genome alignment without WGD or translocation, as represents anchor sequence files, and i, r, and o represent input reference GFF(3), FASTA file, and output MAF file. a and ar represent SAM files generated by mapping conserved sequences to reference and query genome, s represents query genome file, and n represents output anchor files. IV represents calling inversion. Other parameters are default.
anchorwave genoAli -i Zea_mays.AGPv4.34.gff3 -as cds.fa -r Zea_mays.AGPv4.dna.toplevel.fa -a cds.sam -ar ref.sam -s Zm-Mo17-REFERENCE-CAU-1.0.fa -n anchors.anchors -o b73tomo17.maf -f b73tomo17.f.maf -IV
Data analysis
Result interpretation
The final base-pair-resolved whole-genome alignment result is in MAF format, which is fundamental to downstream analysis.
Four figures generated by anchors are obtained to show collinearity. Figure 1 and Figure 2 are used to visualize collinearity between maize and sorghum. Figure 3 and Figure 4 are utilized to illustrate collinearity between maize lines.
From this plot, we can learn the relationship between maize and sorghum. There are chromosome fusions, translocation variations, and an unshared WGD. We will base on this figure to perform genome alignment (Figure 1).
Figure 1. All identified anchors between the maize and sorghum genome. Each dot was plotted based on the start coordinate on the reference genome and query genome of an anchor. Anchors on the same strand between the reference genome and query genome are shown in blue; otherwise, they are shown in red.
AnchorWave implements a longest-path dynamic programming algorithm to identify collinear anchors that guide global genome alignment (Figure 2).
Figure 2. Identified collinear anchors between the maize and sorghum genome
The full-length CDS mapping result is visualized, and collinearity between two maize lines is acquired. B73 genome is consistent with Mo17 genome but some inversions happened (Figure 3).
Figure 3. All identified anchors between the maize B73 and maize Mo17 genome
Figure 4 shows that the genoAli is suitable for genome alignment without translocation rearrangement while with inversions.
Figure 4. Identified collinear anchors between the maize B73 and Mo17 genome
Discussion
We provide detailed guidance on how to perform genome alignment for maize and sorghum using AnchorWave. To run this process successfully, we need to know the WGD variation, translocation variation, or chromosome fusion between genomes. In this protocol, we draw some plots to figure out that information. There are two functions implemented in AnchorWave for genome alignment: genoAli and proali. genoAli is suitable for genome alignment without translocation or chromosome fusion; it is designed to align the genomes from different accessions of the same species. genoAli is also recommended to be used for closely related species. proali is suitable for genome alignment with translocation variation, chromosome fusion, or even WGD. Moreover, base-pair-resolved global sequence alignment may take a couple of CPU days for highly diverse genomes. This step could be parallelized, as long as the computer has enough RAM available. If the two genomes have very similar sequences, time and memory costs are significantly reduced. At present, the AnchorWave team is improving its computational efficiency and extending the AnchorWave to perform multiple genomes alignment.
The genome annotation is essential to provide candidate anchors for AnchorWave, while the quality of genome annotation varies from species to species. We also tested AnchorWave with low quality genome annotation (only ab initio predictions) and the result is available at https://github.com/baoxingsong/genomeAlignment/tree/main/AnnotationCompare.
Acknowledgments
We thank the reviewers for their help in improving the protocol. This work is supported by the National Natural Science Foundation of China No. 31900486 and Shandong Provincial Natural Science Fund for Excellent Young Scientists Fund Program (Overseas) No. 2023HWYQ-109.
Competing interests
The authors declare no conflicts of interest.
References
Song, B., Marco-Sola, S., Moreto, M., Johnson, L., Buckler, E. S. and Stitzer, M. C. (2022). AnchorWave: Sensitive alignment of genomes with high sequence diversity, extensive structural polymorphism, and whole-genome duplication. Proc. Natl. Acad. Sci. U. S. A. 119(1): e2113075119.
Li, H., Handsaker, B., Wysoker, A., Fennell, T., Ruan, J., Homer, N., Marth, G., Abecasis, G., Durbin, R. and Genome Project Data Processing, S. (2009). The Sequence Alignment/Map format and SAMtools. Bioinformatics 25(16): 2078–2079.
Li, H. (2018). Minimap2: pairwise alignment for nucleotide sequences. Bioinformatics 34(18): 3094–3100.
Wickham, H. (2016). ggplot2: Elegant Graphics for Data Analysis, 2nd Edn. New York: Springer.
Supplementary information
Data and code availability: all data and code have been deposited to GitHub: https://github.com/Bio-protocol/anchorwave_protocol.
Article Information
Copyright
© 2023 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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GutMap: A New Interface for Analysing Regional Motility Patterns in ex vivo Mouse Gastrointestinal Preparations
TA Tanya Abo-Shaban *
CL Chalystha Y. Q. Lee *
SH Suzanne Hosie
GB Gayathri K. Balasuriya
MM Mitra Mohsenipour
LJ Leigh A. Johnston
EH Elisa L. Hill-Yardin
(*contributed equally to this work)
Published: Vol 13, Iss 19, Oct 5, 2023
DOI: 10.21769/BioProtoc.4831 Views: 552
Reviewed by: Durai SellegounderBruno Mazet Anonymous reviewer(s)
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Original Research Article:
The authors used this protocol in ACS Applied Materials & Interfaces Jun 2022
Abstract
Different regions of the gastrointestinal tract have specific functions and thus distinct motility patterns. Motility is primarily regulated by the enteric nervous system (ENS), an intrinsic network of neurons located within the gut wall. Under physiological conditions, the ENS is influenced by the central nervous system (CNS). However, by using ex vivo organ bath experiments, ENS regulation of gut motility can also be studied in the absence of CNS influences. The current technique enables the characterisation of small intestinal, caecal, and colonic motility patterns using an ex vivo organ bath and video imaging protocol. This approach is combined with the novel edge detection script GutMap, available in MATLAB, that functions across Windows and Mac platforms. Dissected intestinal segments are cannulated in an organ bath containing physiological saline with a camera mounted overhead. Video recordings of gut contractions are then converted to spatiotemporal heatmaps and analysed using the GutMap software interface. Using data analysed from the heatmaps, parameters of contractile patterns (including contraction propagation frequency and velocity as well as gut diameter) at baseline and in the presence of drugs/treatments/genetic mutations can be compared. Here, we studied motility patterns of female mice at baseline and in the presence of a nitric oxide synthase inhibitor (Nω-Nitro-L-arginine; NOLA) (nitric oxide being the main inhibitory neurotransmitter of gut motility) to showcase the application of GutMap. This technique is suitable for application to a broad range of animal models of clinical disorders to understand underlying biological pathways contributing to gastrointestinal dysfunction.
Key features
• Enhanced video imaging analysis of gut contractility in rodents using a novel software interface.
• New edge detection algorithm to accurately contour curvatures of the gastrointestinal tract.
• Allows for output of high-resolution spatiotemporal heatmaps across Windows and Mac platforms.
• Edge detection and analysis method makes motility measurements accessible in different gut regions including the caecum and stomach.
Graphical overview
Keywords: Gastrointestinal motility Enteric nervous system Ex vivo gut motility Edge detection Software Spatiotemporal heatmap
Background
Gut motility is critical for digestive function, including fermentation and mixing, stool formation, and microbial composition. This contractile activity is predominantly regulated by the intrinsic network of neurons within the gut, the enteric nervous system (ENS) (Furness, 2012). Motility patterns vary in the different regions of the gastrointestinal (GI) tract. In the small intestine, contractile activity serves to maximise exposure to digestive enzymes and absorption of nutrients, whereas the colon is responsible for the formation and expulsion of faeces and absorption of water and electrolytes. Several studies have reported characteristics of contractile patterning in the small and large intestine of small animal species, e.g., rat, mouse, and guinea pig (Lyster et al., 1995; Tonini et al., 1996; Costa et al., 2013 and 2015; Swaminathan et al., 2016; Spencer et al., 2018; Li et al., 2019). However, contractile patterning of the caecum (the equivalent of the human appendix) is not well characterised. Caecal motility has been described in studies of chicken (Janssen et al., 2009; van Staaveren et al., 2020), guinea pig (Schulze‐Delrieu et al., 1996), and rabbit (Hulls et al., 2012 and 2016), but is yet to be modelled in mice or studied in humans.
GI dysmotility is a common comorbidity in many disorders including autism spectrum disorder (ASD; autism) (Gorrindo et al., 2012; McElhanon et al., 2014; Vuong and Hsiao, 2017; Lee et al., 2020) and Parkinson’s disease (Kupsky et al., 1987; Singaram et al., 1995; Cersosimo et al., 2013; Giancola et al., 2017). Given that changes in gut motility patterns can contribute to gastrointestinal symptoms that impact quality of life, such as constipation, diarrhoea, visceral pain, and/or bacterial overgrowth, the accurate measurement of motility patterns could assist in identifying potential therapeutic targets to treat these issues. It is therefore necessary to design and utilise unbiased methods to study gut motility patterns both to gain fundamental knowledge about the neural regulation of motility and to determine changes in these patterns in pre-clinical animal models of disease. Ex vivo motility assays enable the investigation of gut motility patterns in animal models (Roberts et al., 2007; Swaminathan et al., 2016). These techniques were first developed to investigate small intestinal peristalsis in guinea pigs (Hennig et al., 1999; Gwynne and Bornstein, 2007) and then expanded for use in the mouse colon (Swaminathan et al., 2016). This method has been primarily applied to segments of mouse and guinea pig colon (Roberts et al., 2007; Hosie et al., 2019; Leembruggen et al., 2020) and, in some studies, to segments of small bowel (Hennig et al., 1999; Gwynne et al., 2004; Neal et al., 2009). As mentioned previously, studies of caecal motility are restricted to rabbit and chicken (Schulze‐Delrieu et al., 1996; Janssen et al., 2009; Hulls et al., 2012 and 2016). Even though external innervation from the central nervous system (CNS) plays a role in modulating gut motility (Browning and Travagli, 2014), the ENS is capable of independently regulating GI function. Hence, a major advantage of the ex vivo video imaging technique reported here is the ability to measure gut motility in the absence of CNS inputs.
Video imaging combined with spatiotemporal mapping enables the quantitative assessment of gastrointestinal motility in animal models. Using these techniques, multiple contractility parameters such as propagation speed, magnitude, length, duration, diameter, and frequency of gastrointestinal contractions can be measured. However, the software platform we previously utilised for this purpose (Swaminathan et al., 2016) had limited functionality. A lack of compatibility with current software platforms and closed source code that prevented flexibility were major limitations. We found that although suitable for assessing intestinal motility patterns, the edge detection function within the previous version of the MATLAB-based software (Analyse2) could not accurately detect edges of tissue segments from irregular shaped gut regions, such as the caecum, due to the curved/nonlinear anatomy of this organ. In addition, the quality of the heatmaps generated using the previous edge detection function (Swaminathan et al., 2016) was lower, as 16-bit unsigned integer arrays led to a quantisation and pixelation of the resultant spatiotemporal heatmaps. Importantly, this edge detection code was not compatible across different operating systems (i.e., Windows and Macintosh), which reduced user accessibility. Here, we highlight the utility of the new MATLAB-based software interface, entitled GutMap, that enables i) accurate edge detection in multiple gut regions, including in combination with a novel mouse caecal motility protocol (which can additionally be applied to stomach motility), ii) output of high-resolution spatiotemporal heatmaps, and iii) usage across different operating systems.
GutMap is a user-friendly enhanced software interface (available on request) that enables a sensitive and robust approach for visualising and analysing regional gut motility patterns in rodents (with potential applications in other preclinical models following some modification). Data from video recordings of gut contractility patterns acquired from different regions of the GI tract (i.e., the small intestine, caecum, and colon) are converted to high-resolution spatiotemporal heatmaps. The GutMap resolution, measured in μm/pixel or mm/pixel as appropriate to the length scale of the experimental preparation, is entirely dependent on the resolution of the input video file. The spatiotemporal heatmaps generated are high quality as the edge data is stored in double precision arrays, providing a near-continuum of edge locations and thus greater precision of gut width measurements.
Spatiotemporal heatmaps display gut diameter and gut position plotted as a function of time and assign warm colours denoting gut contractions and cool colours indicating gut relaxation. GutMap can be used to measure multiple contractile parameters to characterise subtle changes in motility patterns and ENS activity. Novel features of GutMap include a video calibration function, real-time video tagging information that shows the video properties and edge detection dimensions, as well as a file queueing function for edge detection processing. These features ensure increased accuracy and consistency of measures (i.e., based on the initial tissue size calibration) for each video recording file, as well as streamlined heatmap generation. Heatmap analysis using GutMap enables full functionality of the previous Analyse2 module (Swaminathan et al., 2016), plus an additional novel analysis function that enables measurement of overall contractility of the gut tissue segment.
Here, we provide a comprehensive guide for investigating ex vivo gastrointestinal motility in rodent models and highlight the novelty of our caecal motility measurement protocol in female mouse GI preparations, using the novel video analysis software, GutMap. A detailed explanation is provided for the experimental setup and execution, which are adapted from studies performed in mouse colon (Swaminathan et al., 2016) and rabbit caecum (Hulls et al., 2012 and 2016). Here, we also outline detailed steps for generating and analysing spatiotemporal heatmaps using GutMap. We demonstrate that these processes enable the comparison of motility patterns in different gut regions (i.e., the small intestine, caecum, and colon) both in physiological conditions and in response to drug administration [i.e., the nitric oxide synthase (NOS) inhibitor drug, Nω-Nitro-L-arginine (NOLA)].
The method can be further extended for use in various conditions including examining gut motility in response to different treatments and in other preclinical models of disease.
Materials and reagents
Two-chamber organ bath [manufactured by The University of Melbourne, Department of Physics (Swaminathan et al., 2016)] or equivalent, such as glass Petri dish, 14 cm diameter (e.g., Sterilin 140 mm Petri dish, Thermo Scientific, catalog number: 501V)
Organ bath tubing
Krebs solution inflow tubes
i. Polyethylene tubing 3.00 mm × 2.00 mm (Microtube Extrusions, North Rocks NSW, catalog number: PE300200)
ii. 1 cm of Masterflex Platinum-cured silicone tubing (John Morris Group, catalog number: 96410-14; L/S 14, 25 ft)
iii. Silicone rubber tubing 0.078" ID × 0.125" OD × 50 ft Sil-Med Corporation
iv. Tube connector (200 μL pipette tip) (Axygen, catalog number: AX-T-200-Y)
Vacuum and carbogen tubes
Laboratory tubing 1.02 mm ID × 2.16 mm OD (Silastic, catalog number: 508-005)
Front and back pressure cannulation inflow tubes
i. Single lumen polyethylene tube OD 2.00 mm × ID 1.00 mm used as cannula (catalog number: 112074)
ii. 0.2 cm of laboratory tubing 1.02 mm ID × 2.16 mm OD to create grip at the end of cannula (Silastic, catalog number: 508-005)
iii. Masterflex L/S® precision pump tubing, platinum-cured silicone (John Morris Group, catalog number: L/S 16 96410-16) to connect to 3-way stopcock and silicone tube connector
iv. Masterflex L/S® precision pump tubing, platinum-cured silicone (John Morris Group, catalog number: L/S 14 96410-14) to connect cannula and tube connector
v. Tube connector (200 μL pipette tip) (Axygen, catalog number: AX-T-200-Y)
Circulating water bath tubes
i. Platinum-cured silicone Masterflex L/S® precision pump tubing (John Morris Group, catalog number: L/S 15 96410-15)
Luer Lock tip syringe, 60 mL (Livingstone, catalog number: DSL050MLLCL)
Luer Slip tip tuberculin syringe, 1 mL (Livingstone, catalog number: DSL001MLSC)
Luer Slip tip syringe, 10 mL (Livingstone, catalog number: DS100MTL)
Rubber stopper (Mad About Science, 10 pack, SKU: MAS-03121-1)
Capillary glass tubing, 8 mm diameter, 1.5 mm wall thickness
Multi-purpose sealant [732 Dow Corning (clear) or equivalent]
Suture thread (white cotton thread, Woolworths Supermarket, Woolworths Group, Australia)
Nω-Nitro-L-arginine (NOLA) (Sigma-Aldrich, catalog number: N5501-5G)
Sylgard
184 Silicone elastomer base (DOWSIL)
184 Silicone elastomer curing agent (DOWSIL)
3-way stopcocks (B Braun Discofix, catalog number: 16494C)
Glass media bottle, clear, 500 mL with hose connection at base
Activated charcoal (White Glo)
Filter flask 2 L (Kimax Kimble, catalog number: 27060)
Sodium chloride (NaCl) (Chem-Supply, catalog number: SA046-5KG)
Potassium chloride (KCl) (Ajax Finechem, catalog number: AJA383-500G)
Sodium dihydrogen orthophosphate dihydrate (NaH2PO4·2H2O) (Chem-Supply, catalog number: SA328-500G)
Magnesium sulphate heptahydrate (MgSO4·H2O) (Chem-Supply, catalog number: MA048-500G)
Calcium chloride dihydrate (CaCl2·2H2O) (Ajax Finechem, catalog number: 127-500G)
D-glucose anhydrous (Chem-Supply, catalog number: GA018-500G)
Sodium hydrogen carbonate (NaHCO3) (Chem-Supply, catalog number: SA001-5KG)
Hydrochloric acid (HCl) (Merck Millipore, SKU: 100313)
Distilled H2O (dH2O)
Krebs 10× stock solution in 2 L (4 °C) (see Recipes)
1× Krebs solution in 2 L (4 °C) (see Recipes)
NOLA solution 100 mM (4 °C) 5 mL (see Recipes)
100 μM NOLA-Krebs solution in 500 mL (see Recipes)
Recipes
Krebs 10× stock solution in 2 L (4 °C)
Reagent Final concentration Quantity
Sodium chloride (NaCl) 1.18 M 138 g
Potassium chloride (KCl) 47.6 mM 7.1 g
Sodium dihydrogen orthophosphate dihydrate (NaH2PO4·2H2O) 10 mM 3.1 g
Magnesium sulphate heptahydrate (MgSO4·H2O) 12 mM 5.9 g
Calcium chloride dihydrate (CaCl2·2H2O) 32.99 mM 9.7 g
dH2O n/a 2 L
1× Krebs solution in 2 L (4 °C)
Reagent Final concentration Quantity
Krebs 10× stock solution 10% 200 mL
dH2O n/a 2 L
D-glucose anhydrous 11.1 mM 4.0 g
Sodium hydrogen carbonate (NaHCO3) 25 mM 4.2 g
NOLA solution 100 mM (4 °C) 5 mL
Reagent Final concentration Quantity
Nω-Nitro-L-Arginine (NOLA) 100 mM 109.6 mg
Hydrochloric acid (HCl) 1 M 400 μL
dH2O n/a 4.6 mL
100 μM NOLA-Krebs solution in 500 mL
Reagent Final concentration Quantity
NOLA solution 100 μM 500 μL
1×Krebs l solution n/a 500 mL
Equipment
Webcam (Logitech Carl Zeiss Tessar HD 1080p or equivalent)
Dry block heater (Ratek, model PB)
Circulating water bath 8.5 L (ELMI TW-2.03)
Carbogen, BOC Limited
Vacuum (e.g., laboratory vacuum system)
Retort-stand with clamp (e.g., Australian Scientific, catalog number: SKU: 91055006, 91055028, 120015)
Dumont tweezer, style 1 (Fine Science Tools, catalog number: T01-111)
Dumont forceps, style 5 (Fine Science Tools, catalog number: 11252-30)
Curved forceps (Fisher Scientific, catalog number: 17-467-201)
Fine dissecting scissors (Fine Science Tools, catalog number: 14060-09)
Vannas spring scissors (Fine Science Tools, catalog number: 15000-08)
Amber scintillation 250 μL glass vial (Agilent, catalog number: 5183-2085) for light-sensitive NOLA solution 100 mM
Insect pins (Australian Entomological Supplies, catalog number E-157)
Thermometer (Westlab, catalog number: 072002-0131)
Software
VirtualDub version 1.10.4 64-bit
GutMap (must be downloaded into the MATLAB program files folder)
MATLAB 2021b (or any version of MATLAB that is compatible with Mac or Windows devices):
Curve fitting toolbox
Image processing toolbox
Logitech HD Pro Webcam C920 Driver
Procedure
Dissection and tissue collection
Cull the mouse via cervical dislocation. Exposure to drugs, such as ketamine/xylazine, should be avoided to reduce the potential influence on gut motility.
Weigh and record mouse body weight.
Disinfect the site of incision by spraying the abdomen with 70% ethanol.
Using curved dissecting forceps and small scissors, hold the skin at the mid-lower abdominal area and make an incision. Starting from this initial incision, cut a straight line to the sternum.
Make lateral incisions to open the abdominal cavity and expose the intestinal tract. If required, pour 1× Krebs solution onto the tissue immediately after opening the abdomen to prevent the tissue from drying out during the dissection process.
Lift the liver to expose the stomach and snip the oesophagus to separate it from the stomach. Next, orientate the fine dissecting scissors parallel to the tissue and carefully cut the mesentery along the gastrointestinal tract, while continuing to grasp the gastrointestinal tract by the stomach (using curved forceps), taking care to avoid puncturing the tissue preparation.
Cut the pelvic bone using fine dissecting scissors and carefully remove the mesentery with spring scissors to expose the colon.
Submerge the excised gut in a Petri dish containing approximately 10 mL of 1× Krebs solution and carefully unravel the gastrointestinal tract and cut the required segments for motility. Handle carefully to avoid stretching gut as this can skew results.
Identify the duodenum segment by cutting a 5 cm length of the gastrointestinal tract distal to the stomach. From that point, cut 6 cm of proximal jejunum tissue.
Measure 2 cm proximal to the ileo-caecal junction and 2 cm distal to the caeco-colonic junction to excise the caecum.
From the distal end of the small intestine, measure and cut 5 cm of ileum tissue.
Immediately place the dissected segments in a 100 mL beaker filled with approximately 30 mL of 1× Krebs solution on ice and bubbled with carbogen gas (95% O2, 5% CO2) until the setup is ready for the cannulation step. For best motility results, it is recommended that gastrointestinal segments are cannulated, and video recording begins within 1 h of dissection.
Setting up the organ bath
The organ bath used for the analysis of jejunum, ileum, and colon motility (Figure 1A) was made to order by the Physics Department, The University of Melbourne, Victoria, Australia (specifications as described in Swaminathan et al., 2016). The base of the organ bath is lined with black silicone-based elastomer (Sylgard), and the top surfaces of the chamber walls are sealed with a layer of silicone (Multi-purpose Sealant 732; Dow Corning). The presence of silicone ensures that the bath is watertight and provides a base to pin tubing firmly in place on the setup.
Set up the organ bath by attaching the inlet and outlet tubes using pins embedded into the silicone of the bath. The required tubes include one front pressure (FP) and one back pressure (BP) cannula, a carbogen inflow tube, and a vacuum suction tube. The Krebs solution inflow tube is pre-inserted inside the dual chambered organ bath, and silicone is applied to each inlet to avoid water leaks. Ensure that the Krebs solution inflow tube is a minimum of 45 cm in length to connect to the inflow reservoir.
Attach the Masterflex L/S® precision pump tubing (or an equivalent rubber hose) with an outer diameter of 15 mm to the inlet and outlet pipes of the organ bath and connect the opposite end of the tube to the external water bath to allow heated distilled water to flow inside the dual chambers.
Figure 1. Organ baths to assess motility in (A) small intestine (SI) and colon preparations and (B) caecum. (A) The colon/small intestinal organ bath has two chambers enabling two tissue preparations to be assayed simultaneously. (B) The caecum organ bath is a Petri dish that allows for the three-way cannulation of the caecum preparation. The temperature of setup (A) is regulated by an external water bath. The caecum setup (B) is placed on a heat block to ensure the temperature is kept constant throughout video recording. White arrows indicate the flow of Krebs solution and therefore the pressure exerted on the gut preparations.
For caecum motility (Figure 1B), a 14 cm diameter glass Petri dish (with the base coated with black Sylgard) is used as an organ bath; edges of the Petri dish are also lined with silicone.
Set up the organ bath by attaching the tubes with steel insect pins. Tubes required include one FP cannula, two BP cannulas, a carbogen inflow tube, a vacuum suction tube, and a Krebs solution inflow tube. All FP and BP cannulas require a short (2–3 mm length) rubber sleeve at the cannulating tip. The cannulating tip should be cut at an angle (approximately 45°) for easier cannulation.
Using suture, tie loose knots around the FP and BP cannulas.
Note: Perform this step before the organ bath is filled with 1× Krebs solution because dry thread is easier to handle.
If working with a pre-made organ bath, switch on the external water bath. To ensure that the temperature of 1× Krebs solution in the organ bath is maintained between 33 and 36 °C, the temperature of the external water bath should be between 50 and 60 °C.
If working with the custom-built caecum organ bath, place the Sylgard-lined Petri dish atop a heat block and set to desired temperature (i.e., between 70 and 90 °C, depending on the environmental temperature) to ensure that the 1× Krebs solution temperature is maintained between 33 and 36 °C.
Set up the Krebs solution inflow bottle by fastening it to the retort stand. Connect the in-flow tube attached to the organ bath to the Krebs solution bottle and twist the 3-way stopcock to allow Krebs solution to flow. Ensure that Krebs solution is flowing into the organ bath (flow rate of 8 mL/min or 500 mL/h) by observing bubbles in the capillary tube of the bottle stopper.
Ensure that the Krebs solution in the organ bath is constantly perfused with carbogen gas.
Switch on the vacuum pump and ensure that vacuum tubes are positioned such that the organ bath does not overflow or leak.
Using a 10 mL syringe, flush 1× Krebs solution into the FP and BP inflow valves to clear any debris potentially blocking the FP and BP tubes. Blockages in these tubes will affect the intraluminal pressure.
Pin a laminated printout of a 2 cm ruler onto the Sylgard-lined base of the intestinal or caecal organ bath and ensure that it is visible in the video frame. This will ensure that the gut length and width can be calculated when the user generates a spatiotemporal heatmap.
Cannulating intestinal preparations
Transfer the dissected gut segment to the organ bath. The organ bath should now be filled with warmed 1× Krebs solution. When handling the tissue, take care to hold it by the remaining mesentery and avoid touching the gut tissue, as this will affect motility.
Small intestine and colon:
Using fine forceps, cannulate the proximal end to the FP inflow tube and tie a knot above the rubber sleeve. This will ensure the preparation does not slip off. Tie a second knot to secure the preparation.
Twist the FP stopcock to allow Krebs solution to flow through the lumen and gently push content to clear the lumen.
Cannulate the distal end to the BP outlet tube and tie a knot above the rubber sleeve. This will ensure the preparation does not slip off. Tie a second knot to secure the preparation.
Once cannulated, carefully remove remaining mesentery using fine dissecting scissors to ensure the edges of the intestinal segment are clearly visible during video recording.
Push the stopper of the capillary rod in the FP reservoir to create a pressure seal. Adjust the height so that the meniscus height in the capillary rod is at 6 cm.
Determine the meniscus height in FP and BP tubes by subtracting the height of the FP or BP meniscus from the height of the organ bath.
Cannulating the caecum:
Using fine forceps, cannulate the ileum onto the left cannula that connects to the FP. Tighten the knot above the rubber sleeve to ensure that the preparation does not slip off. Tie a second knot to secure the preparation.
Using fine forceps, carefully grab hold of the caecal tip and snip the tip off (1–2 mm) using fine spring scissors.
Slowly twist open the FP stopcock and allow Krebs solution to flow through the cannula. The FP will activate contractions in the caecum, which will push the caecal contents out through the hole at the caecal tip or colon. To obtain a cleaner preparation (which allows for clearer videos downstream), gently exert a higher FP by using a small 5 mL syringe with a 20 μL pipette tip attached (cut at a 45° angle). Try to clean out the caecum as best as possible; however, avoid exerting too much pressure on the tissue as this will affect motility.
Once the body of the caecum is cleaned out, cannulate the preparation at the caecal tip by tightening the knot on the BP (caecal tip) cannula. Tie a second knot to secure the preparation. Use pins to pin down the cannula in position on the Sylgard-coated surface of the organ bath.
If there is still content in the proximal colon, gently exert pressure from the FP to flush out contents via the colon.
Once the proximal colon is cleaned, cannulate the colon via the BP (colon) cannula. Tighten the knot and tie a second knot to secure the preparation. Use pins to secure the position of the cannula.
Ensure that no blockages are present in each of the cannulas or the caecal preparation by gently exerting pressure via the FP cannula and observing a rising meniscus in the BP tubes.
Remove external FP exertion and allow the FP and BP solutions to stabilise to 4 cm height. The meniscus in the FP capillary tube and BP tubes may bob up and down as the caecum contracts but will stabilise at 4 cm during the quiescence period between contraction complexes.
If the caecum floats or bobs around as it contracts, use a small insect pin to gently pin it down to the Sylgard-coated organ bath surface by the mesentery. Ensure that the entire caecal preparation is submerged beneath 1× Krebs solution in the organ bath.
Check organ bath temperature, FP and BP readings, bubbling carbogen, and position of vacuum suction prior to starting video recording.
Video recording
Open VirtualDub video capture software. Select File followed by Capture AVI… and enter a file name under File and then Set capture file.
To ensure that the videos do not use up a large amount of storage space, compress the video files by selecting Compression under the Video tab, followed by Xvid MPEG-4 Codec.
Uncheck Enable audio capture under the Audio tab to disable audio recording.
Set desired stop conditions by changing the settings under the Capture tab.
Place a covering (i.e., cardboard box) over the organ bath to reduce glare and flickering from the reflection of the bath solution occurring during the video recording.
Adjust the brightness, contrast, exposure, and gain settings of the video using the webcam driver software (e.g., Logitech driver) to further reduce glare or noisy signals, which may interfere with the recording.
In VirtualDub, start video recording by selecting Capture video under the Capture tab.
Record 15 min videos according to the experimental protocol (e.g., 2 × 15 min video recordings during tissue equilibrium and 4 × 15 min video recordings for control, drug, and washout, i.e., generating a total of 14 video files per experiment). It is recommended to capture 15 min videos to enable periodical assessment of the tissue and experimental setup during the intervals between recordings; however, recording duration can be extended for specific experimental protocols as required. Video files showcasing gastrointestinal motility can be viewed in these published articles: Swaminathan et al. (2016); Hosie et al. (2019); Lee et al. (2023).
At the beginning of each video, record the temperature of the organ bath using a thermometer probe and measure the height of FP and BP meniscus. Record information in a table.
Repeat video recording in various conditions
In this example, the nitric oxide synthase (NOS) inhibitor, NOLA 100 mM, was diluted 1:9 with 1× Krebs solution and perfused in the organ bath system to investigate changes in gut motility. NOS is the rate-limiting enzyme in the production of nitric oxide (NO). The main role of NO is to relax the GI tract by inhibiting contractions (Toda and Herman, 2005). Therefore, when NOLA is applied, a reduction in inhibitory tone is expected to result in increased gut constriction and frequency of contractions.
Replace the Krebs solution inflow bottle with a second inflow bottle containing 100 μM NOLA in Krebs solution and repeat steps D5–D7 to visualise motility patterns in the presence of NOLA.
Once recordings in the presence of NOLA are completed, replace with regular Krebs solution, and repeat steps D5–D7 to visualise motility patterns during washout of the drug (NOLA) from the organ bath system.
Cleaning up
Remove the intestinal preparation by snipping the suture with dissecting scissors. Discard the tissue appropriately. Please note: although it is not recommended to use tissue samples following motility experimentation for histological analysis, there is potential to use tissue samples following experimentation to assess for neuronal populations via immunofluorescence techniques.
Using a 50 mL syringe, pass distilled water through all cannulas and Krebs solution inflow tube; repeat this three times per tube.
Use the vacuum suction tube to remove the distilled water from the organ bath.
Finally, use a 50 mL syringe to push out any residual water in the tubing.
Obtaining spatiotemporal heatmaps
Open MATLAB and select Set Path. Add the GutMap folder to the MATLAB path and save.
To generate a spatiotemporal map, type GutMap at the MATLAB command prompt and select Edge Detection (for video files) from the pop-up window.
Load .AVI format video recordings and select Calibrate to define video resolution and the length of the gut preparation. This action will open a new window, including an image of the tissue preparation alongside a physical ruler placed within the organ bath. In this window, use the magnify symbol to zoom in. Use the cursor to select two points 1 cm apart on the ruler that is visible in the window. From these points, GutMap will calculate a pixel/mm calibration metric specific to the current video file, ensuring a spatiotemporal heatmap with accurate dimensions (Figure 2A).
Figure 2. GutMap edge detection control panel. (A) To calibrate the resolution of the current video file, select Calibrate. This will open the calibration figure where the user will be directed to zoom in using the magnifying tool and click on the orange bar. The user selects two points 1 cm apart on the ruler. The calibration figure will then close, and the calibrated resolution value will appear in the Edge Detection Control Panel. Edge detection contours for (B) small intestine and (C) caecum. The selected region of interest contains the upper and lower edges of the gut segment. A red contour delineates the upper edge of the gut, and the green contour identifies the lower edge of the gut. Specifications of the region selected appear in the Video Information area, which can be used to match region-of-interest dimensions between videos. When the gut tissue is adequately contoured, Generate Heatmap is selected by the user. The user is then prompted for the output file name (D) and is next prompted to add the file to the Edge Detection Queue to begin edge detection. For batch processing of multiple files, see instructions in text. A Wait dialogue box appears and indicates the status of the heatmap generation to the user. Once this process is complete, the user must exit.
Using the mouse, draw a rectangle around the region of interest of the gut tissue. A red contour line will define the upper edge and a green contour line will delineate the lower edge of the gut preparation in the image (Figure 2B, 2C).
Use the time slider function to view a selection of frames and ensure gut tissue is adequately contoured prior to proceeding to the heatmap generation step. If there are issues, adjust brightness, contrast, and edge smoothness to enhance edge detection accuracy.
Click Generate Heatmap. This will prompt the user to name the output file. Select a directory to save the file, and then click Add to queue to process the video.
Repeat steps G3–G6 to edge detect additional videos. Set box dimensions to match between videos if desired. Select Begin Detection to edge-detect the videos listed in the queue (Figure 2D).
Data analysis
Heatmap analysis
Click To Heatmap Analysis on the Edge Detection Control Panel to access the Heatmap analysis control panel.
Set the minimum and maximum colour display values. This will display contractions and dilations in warm and cool toned colours, respectively. Click Lock Color Range to set the colour range. It is important to keep the colour range constant throughout a dataset to enable comparison between samples and experiments.
Cross-sectional heatmap analysis
To analyse cross-sections of the spatiotemporal heatmap, select the Take Cross-Sections button (Figure 3A).
Figure 3. GutMap cross-section analysis. (A) In the Heatmap Analysis Window, lock colour range and select the Take Cross-Sections button. This will open a new window (B) where the user can add cross-sectional lines to specific positions on the spatiotemporal heatmap. Output is provided from these cross-sectional lines including gut width, power spectra, and histogram plots of contractile activity. (C) By selecting Pop-Out Plots, these temporal cross-sections can be annotated for further analysis.
Click Add and place a line at a specific position on the heatmap. Three temporal plots will appear below the spatiotemporal heatmap displaying gut width data over time, a frequency power spectrum, and a histogram of contractility frequency (Figure 3B).
The gut width temporal plot displays a range of peaks; maximum peaks represent resting gut diameter, and minimum peaks represent constricted gut diameter.
The frequency temporal plot displays frequency power; lower frequency peaks represent neurogenic input, and higher frequency peaks represent myogenic input.
The histogram plot displays the frequency of contractility.
Temporal plots can be annotated by selecting Pop-Out Plots from the Heatmap Annotation Window to create individual figure windows of the temporal plots. Use the magnification tools to zoom in or out of the temporal plots, and hover over the peaks to place xy-coordinates (Figure 3C).
Cross-sectional heatmap analysis: gut width over time temporal plot
Place xy-coordinates on the gut width vs. time temporal plot to obtain the maximal resting gut diameter, maximal contracted gut diameter, duration of contractile complexes, and quiescent period between contractile complexes.
Cross-sectional heatmap analysis: power spectrum frequency temporal plot
Place data tips on peaks to obtain myogenic and neurogenic input measures.
Cross-sectional heatmap analysis: histogram temporal plot
The histogram plots contractile activity from the selected cross-section line.
Click Export Lines to export data values from the cross-section line in a text file format.
Annotation of contraction waves
To analyse contractions, select Annotate Contraction Waves from the heatmap analysis control panel (Figure 4A).
Figure 4. GutMap contraction wave annotation windows. (A) In the Heatmap Analysis Window, lock the colour range and select Annotate Contraction Waves. A new window will open, allowing the user to manually annotate the contractions or groups of contractions. Annotated examples are shown for (B) colonic, (C) jejunal, and (D) caecal heatmaps [annotated region depicts caecal contractions propagating from the colon-caecal junction (upper red region of map) towards the caecal tip]. The Contraction Wave Annotation tool can be used to measure velocity, duration, and length of the contraction by selecting Manually Annotate and drawing a line following the slope of the contraction or contractile cluster. The data obtained in the results table can be exported by selecting Export Data in the Heatmap Analysis Window.
Use the magnification tools in the Annotate Contraction Waves window to zoom in to the contractions to be analysed.
Select Manually Annotate to draw a line from the start to the end of the contraction (see Figure 4B–4D for examples).
Metrics of the annotated line (velocity, duration of contraction, and length of the annotated contraction) will appear in the table below the heatmap. Note: Contraction Name can be modified.
Select the Update Label button to make changes to the annotated contraction. The results table will update accordingly.
To remove an annotation, select the individual check box within the results table. This will make the selected annotation appear as a red line on the heatmap. Then, click Remove Selected Annotation.
To export the results, return to the heatmap analysis control panel and select Export data (Figure 4A).
Validation of protocol
To compare control and experimental conditions (i.e., motility patterns in the presence of NOLA) data, data tips from each heatmap (four control and four NOLA heatmaps) were averaged. Averaged values can then be inputted to GraphPad Prism version 9.5.0 (or alternative statistical software), where an unpaired t-test is performed to compare various parameters (frequency, diameter, velocity, and duration). This analysis process can be applied to various gastrointestinal tissue segments (i.e., small intestine, caecum, and colon). For example, here we show colonic motility data obtained from n = 3 female mice during baseline and NOLA conditions (Figure 5). We observed a reduction in colonic diameter (constriction) at rest upon NOLA treatment; however, there were no changes to frequency, velocity, or duration of colon migrating motor complexes.
Figure 5. Analysis of colon migrating motor complexes (CMMCs) in n = 3 female mice. Representative colonic heatmaps in (A) control and (B) NOLA conditions display contractile activity in warm tones and relaxation in cool tones. (C) CMMCs that propagated for greater than 50% of the length of the colon were counted and compared between control and NOLA groups. (D) Data tips placed at the peaks of the temporal cross-section plot were averaged to obtain the maximal resting diameter between control and NOLA groups. (E) and (F) The Manually Annotate tool was used to annotate the slope of the CMMCs propagating for greater than 50% of the length of the colon. Velocity and duration results are automatically outputted and then averaged and compared between control and NOLA groups using an Unpaired student t-test.
Acknowledgments
All authors approved the manuscript for submission. ELH-Y was supported by an ARC Future Fellowship (FT160100126) and an RMIT Vice Chancellor’s Senior Research Fellowship. TAS and CYQL received the RMIT Research Stipend Scholarship (RRSS). This protocol describes a significantly enhanced approach based on our previously published methods protocol: Swaminathan et al. (2016).
Competing interests
The authors declare no conflict of interest.
Ethical considerations
All animal handling and euthanasia procedures were performed according to strict guidelines as approved by the RMIT Animal Ethics Committee (AEC#: 21268).
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© 2023 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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Computational Biology and Bioinformatics
Biological Engineering > Biomedical engineering
Neuroscience > Peripheral nervous system
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